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Mutations in Hydin impair ciliary motility in mice
Correspondence to K.-F. Lechtreck: Karl.Lechtreck{at}umassmed.edu; or G.B. Witman: George.Witman{at}umassmed.edu
Chlamydomonas reinhardtii hydin is a central pair protein required for flagellar motility, and mice with Hydin defects develop lethal hydrocephalus. To determine if defects in Hydin cause hydrocephalus through a mechanism involving cilia, we compared the morphology, ultrastructure, and activity of cilia in wild-type and hydin mutant mice strains. The length and density of cilia in the brains of mutant animals is normal. The ciliary axoneme is normal with respect to the 9 + 2 microtubules, dynein arms, and radial spokes but one of the two central microtubules lacks a specific projection. The hydin mutant cilia are unable to bend normally, ciliary beat frequency is reduced, and the cilia tend to stall. As a result, these cilia are incapable of generating fluid flow. Similar defects are observed for cilia in trachea. We conclude that hydrocephalus in hydin mutants is caused by a central pair defect impairing ciliary motility and fluid transport in the brain.
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The CP apparatus consists of two microtubules displaying several projections and connectors (Smith and Lefebvre, 1997). In Chlamydomonas reinhardtii, mutants with a defective central apparatus swim slowly, have abnormal flagellar waveforms, or are paralyzed. The structural defects range from lack of individual projections to loss of the entire CP. Several components of the central apparatus of C. reinhardtii have been identified (Witman et al., 1978; Dutcher et al., 1984; for review see Smith and Yang, 2004), including the 
540-kD protein hydin (Lechtreck and Witman, 2007). Hydin was found in the flagellar proteomes of the protists C. reinhardtii (Pazour et al., 2005) and Trypanosoma brucei (Broadhead et al., 2006), and comparative genomics indicates that the encoding gene is present broadly in organisms with the ability to assemble motile 9 + 2 cilia (Li et al., 2004). The knockdown of hydin in C. reinhardtii resulted in the loss of a specific projection from the central apparatus (Lechtreck and Witman, 2007). Hydin-deficient flagella exhibited paralysis with arrest at the end of the effective or recovery stroke; those displaying residual motility often stopped for extended periods of time at these same positions, where the direction of the beat is reversed. Based on these observations, it was postulated that hydin is a component of a CP projection involved in switching the activity of dynein arms between opposite halves of the axoneme during the transitions between effective and recovery strokes. Knockdown of hydin in T. brucei similarly resulted in CP defects and the loss of flagellar motility (Dawe et al., 2007).
Mice defective in Hydin develop hydrocephalus with early perinatal onset, and most animals die by 3 wk after birth (Raimondi et al., 1976; Davy and Robinson, 2003). Two mutant alleles of Hydin have been characterized. hy3, a spontaneous mutation first described by Gruneberg (1943), carries a single base pair deletion that causes a premature stop that would result in the loss of 89% of the full-length gene product (Davy and Robinson, 2003). The insertional mutation OVE459 is characterized by genomic rearrangement around the insertion site within the Hydin gene (Robinson et al., 2002; Davy and Robinson, 2003). The two alleles do not complement each other and Northern analysis failed to detect hydin transcripts in these mutants. In the wild type, Hydin is expressed in developing spermatocytes and in epithelia lining the brain ventricles, the oviduct, and the airways (Davy and Robinson, 2003). This expression pattern correlates with the presence of motile cilia. This, together with the results from C. reinhardtii and T. brucei, suggests that hydrocephalus in hydin mutants is caused by defects in the ependymal cilia of the brain. Indeed, hydrocephalus has been reported for mice, rats, dogs, and humans with PCD, a disorder impairing ciliary motility (Torikata et al., 1991; Daniel et al., 1995; Afzelius, 1999). In humans, the HYDIN gene is located within a 1.2-Mb fragment to which a hydrocephalus-associated translocation has been mapped (Callen et al., 1990; Doggett et al., 2006), which suggests that defects in HYDIN also may cause hydrocephalus in humans.
Here, we analyzed the structure and movement of motile cilia from the brain and airways of hydin mutant mice and observed the specific loss of a projection from one of the central microtubules. Mutant cilia were unable to bend properly and frequently stalled, which is indicative of a defect in the regulation of dynein arm activity. As a consequence, cilia-generated flow was severely impaired. We conclude that hydrocephalus in hydin mouse mutants is caused by a CP defect and predict that humans with HYDIN mutations will have a higher than normal risk of developing hydrocephalus because of similar defects.
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55%) diminished or absent. Cilia in the trachea of hydin mutants similarly lacked the C2b projection and had altered C1b and C2c projections (n = 13 from two mice; Fig. 2, k and l). The general ultrastructure of the CP and the defects observed in mutant animals are revealed clearly in image averages (Fig. 2, f, h, j, and l). In summary, Hydin deficiency in mice results in the loss of CP projection C2b with accompanying changes to the two adjacent projections.
Ciliary bending is impaired in hydin mutants
To determine how these CP defects affected the motility of cilia in hydin mutant mice, we observed and recorded side, front, and top views of ependymal cilia (Fig. S2, available at http://www.jcb.org/cgi/content/full/jcb.200710162/DC1). The ependymal cilia of wild-type animals (n = 10) exhibited a highly asymmetrical ciliary beat (Fig. 3 a and Video 1).
The beat began with the formation of a large bend at the base of the cilium that swept the cilium forward in a effective stroke (Fig. 3 c) followed by propagation of the bend during the recovery stroke to return the cilium to its starting position (Fig. 3 d). In contrast, the cilia of hy3/hy3 (n = 6) and OVE459 (n = 3) animals appeared to vibrate stiffly without forming distinct effective or recovery strokes (Fig. 3 b and Video 2). Instead, they formed a bend that was smaller than that of the wild type and spread over the proximal 1/2–2/3 of the cilium (Fig. 3 f); the bend then appeared to relax with little propagation, returning the cilium to an unbent position (Fig. 3 e and see Fig. 4 h). Stiff vibrating cilia were similarly observed in trachea of mutant animals, whereas the characteristic ciliary bending pattern was observed in wild-type trachea (Videos 3 and 4). Therefore, Hydin deficiency severely impairs the beat pattern of ependymal and tracheal cilia.
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19% paused for longer periods averaging 17 ms; n = 73 from five animals). In contrast, 79% of Hydin-deficient cilia showed prolonged pauses (mean of 30.1 ± 21.8 ms) and only 5% of the cilia did not stall (n = 64 from four animals). Therefore, the Hydin deficiency delayed the switch between the forward and backward motion of the cilia.
Cilia in hydin mutants beat at a lower frequency
To determine the ciliary beat frequency (CBF) of wild-type and hydin mutant cilia, line scans (kymograms) were obtained from videos showing cilia in top or side views (Fig. 5, a–d).
The changes in grayscale over time in these kymograms were plotted (Fig. 5, e and f) and the CBF was calculated (Fig. 5 g). At ambient temperature, ependymal cilia of wild-type animals had a mean CBF of 10.7 ± 3.7 Hz (n = 16 based on samples from four wild-type plus one heterozygous animal) compared with 6.8 ± 2.8 Hz for mutant animals (n = 18 based on samples from four hy3/hy3 animals); maximum CBF values were 18 and 12 Hz for wild-type and mutant animals, respectively. Similarly, the CBF of tracheal cilia was reduced in mutants (10.4 ± 1.2 Hz) in comparison with the wild type (15.7 ± 3.1 Hz) based on measurements on five and six samples, respectively, from one OVE459 mutant animal and one wild-type littermate (Fig. 5 h).
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Hydin deficiency reduces coordination between cilia
We further noted that the polarity of beat of the ependymal cilia in hydin mutants was defective. In wild-type ependymal cells, the plane of beat is similar for most of the cilia (Fig. 6 a).
Accordingly, lines used to visualize these planes are mostly parallel (standard deviation of 16.8°, n = 54 cilia; Fig. 6 b). In contrast, the beat planes of cilia from mutants are much less well-ordered (standard deviation of 36°, n = 42 cilia; Fig. 6, c and d). This impaired their ability to move in an organized manner. Wild-type cilia exhibited metachronal beating, which was revealed as regular diagonal lines on kymograms (Fig. 6, e and f). In contrast, Hydin-deficient cilia did not display metachrony; line scans of cilia from mutant ependymal epithelia consisted mostly of curves representing individual cilia (Fig. 6, g and h). Thus, Hydin deficiency reduces the coordination between ependymal cilia.
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Hydin deficiency impairs cilia-generated flow
To determine how the mutant cilia affected fluid flow, polystyrene beads (0.5 µm in diameter) were added to brain slices of two mutant OVE459 animals and two wild-type littermates. Particles moved fast and directionally along the ependyma of wild-type mice (Fig. 7 a and Video 5, available at http://www.jcb.org/cgi/content/full/jcb.200710162/DC1).
The velocity of individual particles varied with the distance from the cilia; near the epidermis, velocities of 80–120 µm/s were observed. In contrast, the particles vibrated in place or moved only slowly over the ependyma of mutant mice (10 µm/s; Fig. 7 b and Video 6). Similar results were observed for hy3/hy3 animals and their wild-type littermates (unpublished data). Rapid directional movement also was observed for polystyrene beads added to slices of trachea from wild-type animals (Fig. 7 c), whereas a directional movement of particles was not observed with samples from mutant trachea (Fig. 7 d). In conclusion, the cilia-generated fluid flow in the ventricles and trachea is greatly impaired by the Hydin deficiency.
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Mammalian Hydin is required for proper control of the dynein arms
The lack of Hydin in mice restricted the ability of the cilia to bend. Wild-type cilia generate a strong bend near the base of the cilium during the effective stroke and propagate this bend to the tip of the cilium during the recovery stroke. Cilia of hydin mutants lack the ability to focus the bending to a restricted part of the cilium and then propagate it along its length; instead they alternate between an almost uniformly curved shape and an almost straight shape. This stiff forward-backward movement resembles the motion of cilia with defects in the inner dynein arms or radial spokes in human PCD patients (Chilvers et al., 2003), which suggests that the absence of Hydin specifically affects the CP–radial spoke–inner dynein arm control pathway. This movement is in contrast to the circular beat pattern observed in cilia from PCD patients lacking the complete CP or the almost complete paralysis observed for cilia from PCD patients lacking the outer arms (Chilvers et al., 2003; Stannard et al., 2004; Carlen and Stenram, 2005). The fact that the Hydin-deficient mice cilia still beat in a plane shows that Hydin and the C2b projection of the CP are not required to maintain a planar beat. We further observed that the mutant cilia move with a reduced velocity, which suggests a role for the CP in controlling the speed of dynein-driven interdoublet sliding. This notion is supported by observations on the C. reinhardtii CP complex 1 mutant (cpc1), which lacks the C1b projection and displays normal flagellar bending patterns but has a reduced CBF (Mitchell and Sale, 1999).
Mice cilia lacking Hydin frequently stall, usually at the positions where the direction of beat changes. This defect closely resembles aspects of the phenotype caused by hydin knockdown in C. reinhardtii (Lechtreck and Witman, 2007). The flagella of C. reinhardtii hydin RNAi cells were arrested randomly at the beginning or end of the effective and recovery strokes, and flagella with residual motility often paused in these positions. At these points of reversal of beat direction, the activity of the dynein arms needs to be switched from one side of the axoneme to the other (Satir and Matsuoka, 1989; Nakano et al., 2003; Wargo et al., 2004). Because the hydin-deficient C. reinhardtii flagella were arrested at these switch points, it was hypothesized that hydin is involved in turning the arms on or off in opposite halves of the axoneme. The stalling of the Hydin-deficient mouse cilia suggests that mammalian Hydin is similarly involved in regulating the dynein arms during the transitions between effective and recovery strokes.
The apparent conservation of both the structural and functional roles of Hydin in C. reinhardtii and mammals is remarkable given major differences in the operation of the axonemal machinery. In C. reinhardtii, the CP is twisted and rotates within the axonemal shaft (Mitchell and Nakatsugawa, 2004). Therefore, the CP continuously changes its position relative to the outer doublets, and an individual projection could influence the activity of dyneins on different doublets. In mammals, the CP is in a fixed position and does not rotate, so that a given projection always faces the same group of doublets. Because the basal foot points in the direction of the effective stroke (Gibbons, 1961; Mitchell et al., 2007), the C2b projection faces a subset of those doublets that would be active during the effective stroke (Fig. S3, b and c). It is possible that in the Hydin-deficient mice cilia, these arms fail to be activated, leading to an effective stroke with an attenuated bend. Moreover, if the normal sequential activation of the arms were interrupted at this point, the arms in the opposite half of the axoneme might fail to be activated so that the cilium relaxes and returns passively to its unbent position. In this case, the cilium would appear to bend in only one direction, as observed for the Hydin-deficient mice cilia. Consistent with such a hypothesis, it should be noted that because both bending and beat frequency are much reduced in the Hydin-deficient mice cilia compared to wild-type cilia, the total amount of dynein-driven interdoublet sliding must be greatly reduced in the mutant axonemes.
Impaired ciliary motility causes hydrocephalus in hydin mutants
The abnormal motility of the mutant cilia greatly reduced or eliminated the flow generated by the ependymal cilia. This lack of flow is likely to be the underlying cause of the development of hydrocephalus in these mutants. Impaired ciliary or flagellar motility also has been reported in mice lacking the axonemal dynein heavy chain Mdnah5 or the CP protein sperm-associated antigen 6 (Spag6; Ibanez-Tallon et al., 2002; Sapiro et al., 2002; Zhang et al., 2007). Mdnah5 mutants have severely abnormal motility of the ependymal cilia and reduced ependymal flow (Ibanez-Tallon et al., 2004). Mice deficient in Spag6 display significantly reduced sperm motility (Sapiro et al., 2002). Both mutations produce hydrocephalus. This, collectively with our findings, indicates that impaired ciliary motility alone is sufficient to cause hydrocephalus in mice. In the mdnah5 homozygous mice, stenosis of the cerebral aqueduct between the third and fourth ventricles was observed on P6; this resulted in triventricular hydrocephalus with massive enlargement of the third ventricle, whereas the fourth ventricle did not enlarge (Ibanez-Tallon et al., 2004). Therefore, it was proposed that ciliary motility is required to keep the narrow cerebral aqueduct open.
Hydrocephalus also occurs in tg737orpk mutant mice (Banizs et al., 2005). These mice are homozygous for a hypomorphic allele of IFT88/polaris, a component of the intraflagellar transport machinery required for the assembly of motile and immotile cilia (Pazour et al., 2000). As a consequence, the mice develop cilia-related defects, including polycystic kidney disease and retinal degeneration, and the ependymal cilia are malformed and fail to generate ependymal flow. The cerebral aqueduct becomes blocked by P6, but the first signs of disease are evident before blockage is apparent. Therefore, it has been suggested that defects in the primary cilia on the choroid plexus of tg737orpk mutant mice result in an overproduction of cerebrospinal fluid leading to hydrocephalus (Banizs et al., 2005, 2007). This precise scenario is unlikely for hydin and spag6 mutants because the defective gene products are located in the CP, which is absent in primary cilia. Furthermore, these mutants lack the pleiotropic disorders characteristic of mutants having defective primary cilia (Badano et al., 2006). However, one cannot rule out the possibility that defects in the motile cilia of the choroid plexus or the ependyma cause cerebrospinal fluid overproduction by impairing the proper distribution of factors required to control its production.
Loss of Hydin does not cause situs inversus
The nodal cilia of mice embryos undergo an unusual whirling movement that generates a leftward flow of extraembryonic fluid that is required for the establishment of left–right asymmetry; an inability to assemble these cilia or impairment of their motility results in the randomization of left–right asymmetry (Nonaka et al., 1998; Ibanez-Tallon et al., 2002). Although nodal cilia have generally been thought to have a 9 + 0 axoneme lacking a CP, central microtubules recently were observed in some cilia of the mouse node and in up to 62% of the cilia in certain regions of the notochordal plate (equivalent to the mouse node) in rabbits, which raises questions about the function of the CP in these cilia (Feistel and Blum, 2006; Caspary et al., 2007). Our observation that hydin mutant mice lack situs abnormalities indicates that Hydin, at least, has no important role in generating the whirling motility of nodal cilia. Similarly, mutation of the CP protein Spag6 in mice does not cause situs abnormalities (Sapiro et al., 2002). Neither have situs abnormalities been observed in PCD patients with CP defects (Sturgess et al., 1979; Tamalet et al., 2001; Chilvers et al., 2003; Stannard et al., 2004; Carlen and Stenram, 2005). These observations strongly suggest that the CP itself is not required for the whirling motion characteristic of nodal cilia (Okada et al., 2005) and, consequently, CP defects do not result in situs abnormalities. Consistent with this, all posterior notochordal cilia in the rabbit exhibit a whirling movement (Okada et al., 2005; Feistel and Blum, 2006) even though some of these cilia have central microtubules and others do not.
HYDIN, CP defects, and human disease
In the hydin mutant mice, the motility of the tracheal cilia also was severely impaired. Therefore, we would expect that humans with defects in HYDIN would have ciliary dyskinesia and develop PCD, although without accompanying situs inversus (see previous section). Definitive diagnosis of PCD in patients without situs inversus traditionally requires the demonstration of loss of an axonemal structure (in airway cilia or sperm flagella) by EM and/or the demonstration of defective ciliary movement on airway cells obtained by nasal brushing or biopsy (Meeks and Bush, 2000; van's Gravesande and Omran, 2005). Based on our results, defects in human HYDIN would cause a subtle ultrastructural defect that might easily escape EM analysis. In fact, in at least 3% of PCD patients, no ultrastructural defect is observed. In such patients, we suggest that particular care should be given to examination of the CP. Similarly, defects in human HYDIN would be expected to cause a relatively small reduction in beat frequency that might not be noticed by methods that examine CBF but not waveform or the ability to generate fluid flow. Therefore, if nearly normal CBF is found in suspected PCD patients, the waveform and coordination of the cilia should be examined for abnormalities, ideally by high-speed video microscopy (Chilvers et al., 2003; Noone et al., 2004).
Are defects in HYDIN likely to cause hydrocephalus in humans? As noted in the Introduction, a mutation causing hydrocephalus has been mapped to within 1.2 Mb of the HYDIN locus at 16q22.2-q22.3 (Callen et al., 1990). Although not all patients with defects in motile cilia develop hydrocephalus, there are numerous cases where an association between PCD and hydrocephalus has been reported (De Santi et al., 1990; al-Shroof et al., 2001; Kosaki et al., 2004), and the incidence of hydrocephalus caused by aqueduct stenosis in PCD patients is estimated to be 83x higher than in the general population (Ibanez-Tallon et al., 2004). It is likely that morphological differences such as the diameter of the cerebral aqueduct in the developing brain are responsible for the different effects of dysmotile cilia on humans, where immotile cilia most commonly cause PCD, and mice, where defects in motile cilia almost always cause hydrocephalus. In any case, we predict that dysmotile cilia caused by defects in HYDIN will increase the risk of hydrocephalus in humans.
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Video microscopy of ependymal and tracheal cilia
Animals were killed by intraperitoneal injection of pentobarbital sodium. Brains were removed, washed in HBSS (Invitrogen) supplemented with 25 mM Hepes (sHBSS, pH 7.4), trimmed for sagital or coronal sectioning of the third and lateral ventricles, and sectioned into 130-µm slices using a vibratome (OTS-4000; Electron Microscopy Sciences). Sections in sHBSS were observed with differential interference contrast microscopy using an inverted microscope (IX71; Olympus) equipped with a 60x water immersion objective (numerical aperture 1.20) and a zoom adaptor (Nikon). Images (640 pixels x 480 lines) of ciliary activity were recorded at 200 frames per second with a high-speed, progressive scan charge-coupled device camera (TM-6740; Pulnix) and image acquisition software (Video Savant; IO Industries) as described previously (Zhang and Sanderson, 2003; Delmotte and Sanderson, 2006). Samples were analyzed at room temperature typically within 25–60 min after euthanasia; importantly, cilia continued to beat rapidly for 24 h in slices incubated in Dulbecco's Modified Eagle's Medium supplemented with 10% FBS, penicillin, and streptomycin at 37°C in 10% CO2. The CBF of ependymal cilia was calculated as described in Results. To visualize cilia-generated fluid flow, sections with an exposed third ventricle were placed into chambers and polystyrene beads (0.5 µm in diameter; Sigma-Aldrich) were added.
For study of airway cilia, trachea were removed, cleaned under a dissecting microscope, cut into rings or strips to record side and top views, respectively, and observed as described in the preceding paragraph. The CBF of tracheal cilia was measured as described previously (Delmotte and Sanderson, 2006).
For still images and slow-motion videos, individual frames were cropped and adjusted for brightness and contrast in Photoshop (Adobe). Figures were assembled using Illustrator (Adobe). Line scans (kymograms) were prepared by extracting a row of pixels from each image of a series and placing them sequentially in time to create a single image using Scion Image (Scion Corporation). Videos were made using QuickTime 7.2 (Apple). To analyze the movement of individual cilia, frames were copied from QuickTime movies into Illustrator, and cilia were traced using the paintbrush tool. When the target cilium was not entirely visible, missing parts were filled in based on observations made on other cilia.
Ultrastructural analysis of cilia
Brain slices and trachea ring sections were made as described in the previous section or thicker slices were made by hand and fixed for 2 h with 2% glutaraldehyde and 2.5% formaldehyde in 75–100 mM cacodylate buffer. After several washes in buffer, the tissues were treated with 1% OsO4 for 1 h. For TEM, fixed specimens were washed twice with buffer and twice with water and incubated overnight in aqueous 1% uranyl acetate at 4°C. After several washes in water, the slices were dehydrated, embedded, and sectioned using standard procedures. Brain slices were flat embedded and then mounted onto epon blocks to ensure proper orientation of the cilia. Thin sections were analyzed using CM10 and CM12 electron microscopes (Philips). For SEM, samples were washed and dehydrated after OsO4 fixation (see TEM fixation protocol), critically point dried, coated with a 4-nm-thick layer of iridium, and examined using a field emission SEM (Quanta 200F; FEI). Averaged images were prepared with Photoshop.
Immunofluorescence microscopy
For immunofluorescence microscopy, freshly prepared trachea from a 6-d-old mutant and a 5-d-old wild-type animal were brushed with a wooden stick, washed into HBSS supplemented with 25 mM Hepes, pH 7.4, centrifuged onto poly-L-lysine–coated coverslips, fixed with methanol at –20°C for 8 min, dried, and then blocked with PBS containing 0.05% Tween 20, 3% fish gelatin, and 1% BSA for 30 min. Anti-acetylated tubulin (1:800; Sigma-Aldrich) was applied overnight at 4°C and goat anti–mouse F(ab)2 of IgG Alexa Fluor 488 (1:400; Invitrogen) was applied for 90 min. Images were acquired using Axiovision software and a camera (AxioCam MRm) on a microscope (Axioskop 2 Plus) equipped with a 100x 1.4 numerical aperture oil differential interference contrast Plan Apochromat objective (all from Carl Zeiss, Inc.) and epifluorescence. Image brightness and contrast were adjusted using Photoshop 5.0. Figures for publication were assembled using Illustrator 8.0. Capture times and adjustments were similar for images mounted together.
Online supplemental material
Fig. S1 shows SEM images of brain and trachea from wild-type and hy3/hy3 animals demonstrating that the assembly of cilia is not affected by the loss of Hydin. Epithelial cells from trachea were also stained with anti-acetylated tubulin for immunofluorescence to show the presence of full-length cilia. Fig. S2 shows two video frames each of side, front, and top views of wild-type ependymal cilia. Fig. S3 shows TEM images of a wild-type epithelial cell from the trachea in overview and detail revealing that the CP projection C2b points in the same direction as the basal foot, a marker pointing in the direction of the power stroke. Videos 1 and 2 show the motility of ependymal cilia from wild-type and hydin mutant animals, respectively. Videos 3 and 4 show the motility of tracheal cilia from wild-type and hydin mutant animals, respectively. Videos 5 and 6 show the flow generated by wild-type and mutant ependymal cilia. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200710162/DC1.
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Acknowledgments |
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This study was supported by National Institutes of Health grants to M.J. Sanderson (HL071930) and G.B. Witman (GM030626) and by support from the Robert W. Booth Fund at the Greater Worcester Community Foundation to G.B. Witman. Core facilities used in this research were supported by Diabetes Endocrinology Research Center (grant DK32520).
Submitted: 23 October 2007
Accepted: 9 January 2008
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