How do cilia beat?

How do cilia beat?

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From what I have learnt, the mechanism of the beating of cilia is: movement of dynein towards the (-)-end i.e. towards the microtubule-organizing center (MTOC), causing the 2 pairs of microtubules to (supposedly) slide. This is prevented by linker proteins anchored in the adjacent pairs of microtubules. Therefore, when dyneins move, the linker proteins become skewed hence and bending of cilia.

My question is: As far as I know, dynein can only move in one direction (towards (-)-end) i.e. the cilia can only move in one direction, which is forward beating action. I would like to ask how can the cilia move back to the original position (backward movement) and initiate the next beating?

Edit: I have checked Wikipedia, it says there is a process called intraflagellar transport (IFT), which is bi-directional and allows the retrograde transport of cilia (backward movement?) by dyneins, which seems contradictory to what I have learnt, so how can this process actually happen?

Celebrate Cytochemistry

Pituitary gonadotropes are immunolabeled fluorescent green for LH (see above banner) and nuclei stain blue with DAPI. However, in the above view, they are dual labeled for Cre-recombinase with dylight 594 (red) in the nuclei and cytoplasm. This makes the nuclei purple and the cytoplasm yellow.

5 nM Gold markers detect LH in the Golgi complex and in a secretory granule.

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How Cilia Do the Wave

Thin, hair-like biological structures called cilia are tiny but mighty. Each one, made up of more than 600 different proteins, works together with hundreds of others in a tightly-packed layer to move like a crowd at a ball game doing "the wave." Their synchronized motion helps sweep mucus from the lungs and usher eggs from the ovaries into the uterus. By controlling how fluid flows around an embryo, cilia also help ensure that organs like the heart develop on the correct side of your body.

But despite cilia's importance, scientists don't have a good understanding of the mechanism that controls how cilia beat in unison to perform their many essential functions. To investigate this, a group of federally-funded researchers at Brandeis University, led by Zvonimir Dogic and Daniela Nicastro, created the first-ever models of artificial cilia.

The main ingredient was microtubules, or hollow protein tubes that give plant and animal cells structure and help organize and move their components for cell division. Motor proteins and a compound that assembles microtubules into bundles also went into the mix.

Inside a machine called a flow chamber, the artificial cilia moved like the real thing: They beat together in a series of synchronized, self-organized waves. In some cases, as you see here, the lab-made cilia could even push debris along the surface of a bubble, mimicking transport along a cell's surface.

As the first example of a system that beats like cilia, the new models could have applications in fields ranging from cell biology to physics and nanoscience. The models will also open new doors for studying ciliopathies, rare but serious genetic disorders that result when cilia don't move normally.

The researchers anticipate that artificial cilia could even shed light on other self-organizing systems, such as bacterial colonies, flocks of migrating birds and traffic patterns.

This research was supported by the National Institutes of Health (NIH) and the National Science Foundation (NSF). To see more cool images and videos of basic biomedical research in action, visit the NIH's Biomedical Beat Cool Image Gallery.

Any opinions, findings, and conclusions or recommendations expressed in this material are those of the author and do not necessarily reflect the views of the National Science Foundation. See the Research in Action archive.

Cilium Function

Cilia can help to remove contaminants from organs or tissue by helping to move fluids over the cell. The lining of the nasopharynx and the trachea are covered in cilia. These ciliated epithelial cells remove mucus, bacteria, and other debris from the lungs. Another example is the lining of the fallopian tubes. The cilia here are responsible for helping in fertilization by movement of the egg towards the uterus.

Kinocilia are a specialized type of cilia found on the apical ends of vertebrate hair cells. Along with stereocilia, non-motile collections of actin filaments related to cilia, they are involved in hearing and balance (mechanoreception).

This figure depicts the internal structure of a cilium, showing the nine pairs of outer microtubules and the two central microtubules, connected by protein linkers and dynein molecules.

This figure depicts epithelial cells covered in small hair-like cilia.

1. What are cilia composed of?
A. Microfilaments
B. Microtubules
C. Keratin
D. Actin

2. What type of organism does not have cilia?
A. Bacteria
B. Protists
C. Plants
D. Animals

3. Which of the following is not a function of cilia?
A. Locomotion
B. Feeding
C. Reproduction
D. Fighting infection
E. None of the above

Structure and Function of Cilia

Structurally, each cilium comprises a microtubular backbone - the ciliary axoneme - surrounded by plasma membrane (see figure below).

Motile cilia are characterized by a typical ’9+2’ architecture with nine outer microtubule doublets and a central pair of microtubules (e.g bronchi).

Primary cilia appear typically as single appendages microtubules on the apical surface of cells and lack the central pair of microtubules (e.g. in kidney tubules).

Ciliary proteins are synthesized in the cell body and must be transported to the tip of the axoneme. This is achieved by Intraflagellar Transport (IFT), an ordered and highly regulated anterograde and retrograde translocation of polypeptide complexes (IFT particles) along the length of the ciliary axoneme.

Dysfunction or defects in motile and primary cilia are now understood to underlie a number of devastating genetic conditions - termed ciliopathies - which carry a heavy economic and health burden on individuals, families and society.

Much is still unknown about the structure and function of motile and primary cilia, but we believe that more research into these critically important cellular organelles will eventually bring about better ways to treat and help people whose lives are impacted by defective cilia.


Important to kidney function, cilia monitor the flow of urine in this pair of organs. For nearly a decade, scientists have looked for a link between kidney disease and cilia. Although the exact mechanism isn’t clear, researchers believe that kidney cilia become damaged and unable to monitor urine flow, causing the kidneys to become scarred and diseased, leading to kidney failure.

Chlamydomonas mutants and structural analysis reveal functional specialization of many conserved axonemal dynein motors and regulators of motility

Chlamydomonas mutants have been particularly informative, yielding insight into the composition and structure of the axoneme, the regulation of motility, the role of different dynein motors, and the assembly and length regulation of the cilium (Avasthi and Marshall 2012). Here, we focus on how Chlamydomonas has furthered our understanding of the mechanism and regulation of ciliary bending and, particularly, how it has contributed to knowledge of the general roles of the outer and inner dynein arms for the control of beat frequency and waveform.

Initially and most generally, the axonemal dyneins are referred to as the outer and inner dynein arms, but, in reality, there are many different, conserved dynein motors that are highly localized in the axoneme and that serve special purposes for the control of movement (King and Kamiya 2009). The best understood axonemal dynein is the outer dynein arm. The outer dynein arms are biochemically very complex: each is composed of two or three distinct dynein ATPases and at least 16 different subunits. They are assembled individually in rows on doublet numbers 2–9 in Chlamydomonas axonemes and repeat at a regular 24-nm period along each outer doublet (see figure 4 Goodenough and Heuser 1982). Along most of the axoneme, each outer dynein arm is structurally—and, usually, biochemically—identical to its neighbor. Mutations in the genes that encode the structural proteins of the outer dynein arm or associated factors involved in outer dynein arm targeting and assembly can result in a failure to assemble all or a part of the outer dynein arm and, consequent, in defective motility (King and Kamiya 2009).

The Chlamydomonas axoneme ultrastructure. Cryoelectron tomography slices show (a) a longitudinal, (b) a three-dimensional view, and (c) a cross-sectional view of a Chlamydomonas axoneme. The red boxes highlight one 96-nanometer (nm) axonemal repeat unit in each view. (d, e) Isosurface renderings and (f, g) a simplified schematic show an averaged 96-nm axonemal repeat in (d, f) longitudinal and (e, g) cross-sectional orientation. The cross-sectional slice is taken close to radial spoke 2, viewing from the proximal to the distal end. Key axonemal structures are highlighted: the A- and B-tubules (At, Bt), the nexin-dynein regulatory complex (N-DRC), radial spokes (RS1, RS2), the calmodulin and spoke associated complex (CSC), and the inner and outer dynein arms (IA, OA). The inner arm dyneins include the I1 complex (dynein f α and β) and dyneins a–g. Source: Adapted with permission from Heuser and colleagues ( 2012a, 2012b).

The Chlamydomonas axoneme ultrastructure. Cryoelectron tomography slices show (a) a longitudinal, (b) a three-dimensional view, and (c) a cross-sectional view of a Chlamydomonas axoneme. The red boxes highlight one 96-nanometer (nm) axonemal repeat unit in each view. (d, e) Isosurface renderings and (f, g) a simplified schematic show an averaged 96-nm axonemal repeat in (d, f) longitudinal and (e, g) cross-sectional orientation. The cross-sectional slice is taken close to radial spoke 2, viewing from the proximal to the distal end. Key axonemal structures are highlighted: the A- and B-tubules (At, Bt), the nexin-dynein regulatory complex (N-DRC), radial spokes (RS1, RS2), the calmodulin and spoke associated complex (CSC), and the inner and outer dynein arms (IA, OA). The inner arm dyneins include the I1 complex (dynein f α and β) and dyneins a–g. Source: Adapted with permission from Heuser and colleagues ( 2012a, 2012b).

Consistent with the pioneering studies of Gibbons BH and Gibbons ( 1973), the most notable consequence of failure in assembly of the outer dynein arm is the reduced ciliary beat frequency (Brokaw and Kamiya 1987, Brokaw 1994, Kamiya 2002). The outer dynein arm and ciliary beat frequency can be regulated by phosphorylation and changes in calcium (Christensen et al. 2001, King and Kamiya 2009, King 2010). In the ciliates Paramecium and Tetrahymena, an increase in cAMP around the axoneme produces faster swimming by phosphorylating a small outer dynein arm–associated protein (Christensen et al. 2001). In vitro, the phosphorylated outer dynein arm causes faster sliding of microtubules, which implies that less time is taken to produce the same amount of sliding throughout a beat cycle—that is, that the beat frequency is faster. Diverse evidence also indicates the outer dynein arms respond to mechanical perturbation of the axoneme (Hayashibe et al. 1997). A mechanical feedback is probably important for switching between active and inactive states and required for forward and reverse bending. Consistent with a mechanical feedback control, as was discussed above (Shingyoji et al. 1977), bending of the axoneme can activate dynein-driven microtubule sliding (Morita and Shingyoji 2004, Hayashi and Shingyoji 2008). In addition, these data may also indirectly address an important model for the mechanical control of bending, called the geometric clutch hypothesis, which envisions changes in interdoublet distances corresponding to arm activity and bend production related to distortion of the axoneme during bending (for a full discussion, see Lindemann 2011). The predicted distortion in the axoneme during bending has been observed through electron microscopy (Lindemann and Mitchell 2007).

The inner dynein arms are much more complex than the outer dynein arms (for reviews, see Kamiya 2002, King and Kamiya 2009). Along an axonemal 96-nm repeat on a doublet microtubule (figure 4, which has four identical outer dynein arms) the inner dynein arms include at least seven different dyneins, which are distinct in composition and location (see Bui et al. 2012). The inner dynein arms were characterized through biochemical fractionation of dynein components and through electron microscopy of axonemes missing subsets of the inner dynein arms (King and Kamiya 2009). In Chlamydomonas axonemes, which lack a subset of inner dynein arms, the ciliary waveform is altered, phototaxis is disrupted, and the swimming speed of the cells is slowed. A direct analysis of ciliary beating through high-speed video has confirmed that a failure in assembly of any of the individual inner dynein arms results in an altered ciliary waveform, a parameter that is essential to effective ciliary beating and physiology (Brokaw and Kamiya 1987).

The precise role of each inner dynein arm is not yet understood. However, one of the two-headed inner dynein arms, called I1 dynein, is thought to be particularly important for the control of axonemal bending. I1 dynein activity is regulated by kinases and phosphatases located in the axoneme (for a review, see Wirschell et al. 2011). Changes in I1 activity can be measured through changes in microtubule sliding velocity, but it is not yet known how changes in velocity corresponds to changes in bending. One possibility is that an increase in sliding velocity produced by the inner dynein arms without a change in beat frequency would correspond to an increase in bend magnitude. In addition, I1 dynein may regulate bending through control of the activity of other dyneins, including the outer dynein arms and the single-headed inner dynein arms (Kotani et al. 2007, Yamamoto et al. 2013).

In general, we do not yet know the function of each of the axonemal single-headed dyneins. However, an analysis of a mutant called ida9 has revealed that one of the inner dynein arms, dynein c, is required for ciliary movement in a viscous fluid (Yagi et al. 2005). In addition, powerful screens have revealed new mutants that regulate inner dynein arm activity (Kamiya et al. 1991, Kamiya 2002). For example, screens have revealed the enzymes responsible for polyglutamylation and that this posttranslational modification of tubulin is crucial for the activity in a subset of the inner dynein arms (Kubo et al. 2010).

The central pair and radial spokes are required for normal ciliary motility and the control the dynein motors

In Chlamydomonas, a failure in assembly of the central pair or radial spokes results either in ciliary paralysis (Witman et al. 1978, Smith EF and Yang 2004) or in greatly altered and unproductive bending movement. Together, the central pair apparatus and the radial spokes operate by both mechanical and chemical signaling to ultimately control axonemal dynein activity (Smith EF and Yang 2004). The mechanism of interaction between the central pair and the radial spoke was confirmed in recent experimental studies involving Chlamydomonas (Oda et al. 2014). They showed that the addition of nonspecific proteins to the radial spoke head could suppress paralysis of a paralyzed central pair mutant that was missing part of the central pair projections. The simplest interpretation of this result is that the added proteins permitted a physical interaction between the spoke head and the projections that is required to activate axonemal dyneins.

Transmission of signals from the central pair and the radial spokes to the dynein motors

The analysis of Chlamydomonas mutants has also revealed conserved axonemal components that transmit signals from the central pair and radial spoke structures to the dynein motors. Most notable are the calmodulin and spoke associated complex (CSC Dymek et al. 2011, Heuser et al. 2012a) and the DRC see the next section for insights from recent cryoelectron tomography (cryo-ET) analyses of the axoneme (for reviews, see Heuser et al. 2009, Porter 2012) and of the Mia–I1 dynein complex (Yamamoto et al. 2013). The CSC and the DRC play roles in the control of the dynein motors: They are both ideally located to link the radial spokes to the outer doublets and the dynein motors. An important current goal is to determine how calcium and the calmodulin complexes, located in the central apparatus and the CSC, operate to control axonemal dyneins. The I1 dynein and the associated Mia complex (Yamamoto et al. 2013) may also play a role similar to that of the DRC in the regulation of or resistance to dynein-driven microtubule sliding and in the control of axonemal bending.

Suppressor mutations in Chlamydomonas and the DRC

Major insight for understanding the control of axonemal dyneins resulted from the classic genetic studies of Huang and colleagues ( 1982). A genetic screen revealed new genes that suppressed paralysis (i.e., rescue motility) in radial spoke or central pair mutants so that the mutants once again became motile. The suppressor mutants included new mutations in the dynein motor heavy chains that restored movement without the recovery of the radial spoke and without central pair defects. These results indicate that, in the absence of the radial spokes and the central pair, dynein motor activity is inhibited throughout the axoneme, but it can be restored by any molecular change that permits the dynein to become active without input from the spoke–central pair system.

The same genetic suppressor screen (Huang et al. 1982) revealed another regulatory complex, which was named the DRC by Piperno in 1994. Subsequent studies further defined the DRC protein subunits (for a review, see Porter 2012). A major recent advance, made possible by the increased resolution of cryo-ET, is the discovery that the DRC is also the nexin interdoublet link. Therefore, the DRC structure is now called the nexin-dynein regulatory complex (N-DRC Heuser et al. 2009). Recent studies have also shown a biochemical–structural interaction between the N-DRC and the outer dynein arm intermediate chains, physically linking the outer dynein arm and the N-DRC (Oda et al. 2013). Therefore, the N-DRC appears to play roles in the regulation of the dynein motors and serves as an interdoublet link such that it likely aligns outer doublets to ensure efficient interactions between the dynein motors and the B-tubule of the adjacent doublet microtubule (Bower et al. 2013). To accommodate the sliding of doublet microtubules, all N-DRC–outer dynein arm interdoublet links on the sliding doublet must, at some point, break their connection with the adjacent B-microtubule (Holwill and Satir 1990). In a final understanding of the switch point model, it will be important to know the manner in which the N-DRC links are regulated and the timing of breakage and reformation for the active and passively sliding halves of the axoneme.

Insights from cryo-ET

A new level of resolution of the axoneme has been achieved through cryo-ET. These data have already had a major impact on the understanding of ciliary movement. The basic methodology has been reviewed (e.g., Nicastro 2009, Bui and Ishikawa 2013), and methods that now merge structural localization with cryo-ET have been described (Oda and Kikkawa 2013). Briefly, the process involves the rapid freezing of live cells or isolated axonemes, followed by low-dose electron microscopy of the structures in the frozen native state, data collection, computation-based tomography, and image averaging, which produce high-resolution images in three dimensions. Combined with very informative structural mutants in Chlamydomonas, a very-high-resolution picture of axonemal structure has emerged. The new structural information is summarized in figure 4, including examples of electron tomograms (figure 4a– 4c), averages and 3D isosurface renderings (figure 4d, 4e), and summary schematic diagrams of a single doublet microtubule in longitudinal and cross section (figure 4f, 4g).

The cryo-ET approach offer numerous advantages: high resolution, approaching and exceeding 3 nm, revealing the detained substructure of each axonemal component pristine preservation by rapid freezing and visualization of structures in 3D in intact organelles. The notable features in figure 4 include a definition of the longitudinal 96-nm axonemal repeat (figure 4 D and E), which is probably the basic unit of activity along the axoneme and the substructure of the outer dynein arms, including the resolution of the globular motor domains (figure 4d– 4g). Also illustrated are the locations and substructure of each inner dynein arm (I1/f dynein and dyneins a–g Bui et al. 2012, Heuser et al. 2012b, Lin et al. 2014), the N-DRC (Heuser et al. 2009), the CSC (Dymek et al. 2011, Heuser et al. 2012a), and the radial spokes (Pigino et al. 2011, Barber et al. 2012, Oda et al. 2014). In addition, cryo-ET has revealed the substructure of the central pair apparatus (Carbajal-Gonzalez et al. 2013, Oda et al. 2014).

In a relatively short time, these structural advances have contributed new understanding of the structural basis of ciliary bending. For example, cryo-ET has also revealed physical links among the outer dynein arms, the inner dynein arms, and the N-DRC that could explain the coordination of activity among the structures (e.g., Oda et al. 2013). This result begins to address a major question—that of how the activity of the outer and inner dynein arms is coordinated (Kamiya 2002). Along with other biophysical and structural studies, cryo-ET has also revealed a structural basis for axonemal dyneins’ power stroke (Lin et al. 2014). The promise of these new studies is that these structural advances will define the structural changes associated with ciliary bending and will directly test models for cilairy bend formation and bend propagation. For example, the structural analysis of rapidly frozen live sea urchin sperm axonemes reveals that the structural state of the dynein motors on the doublet located on the inside of the bend differs from the state of the dyneins located on the doublets on the opposite side of the axoneme, on the outside of the bend (see supplemental figure 1 in Lin et al. 2014). These data appear to support a switch point model for axonemal bending, possibly revealing “on” and “off” states of the dyneins (see Brokaw, 2009). Predictably, taking advantage of cryo-ET and preservation through freezing, further analysis of the sea urchin sperm axonemes, Chlamydomonas mutants, and cilia preserved in metachronal beat stages will provide a comprehensive picture of axonemal and dynein structural changes associated with bend initiation and bend propagation.

Cilia beat to an unexpected rhythm in male reproductive tract, study in mice reveals

Waves of undulating cilia drive several processes essential to life. They clear debris and mucus from the respiratory tract, move spinal fluid through the brain and transport embryos from the ovaries to the uterus for implantation. According to a new study in mice, however, cilia perform somewhat differently in the male reproductive tract.

The study, reported in the Proceedings of the National Academy of Sciences, reveals that cilia in the efferent ductules, which carry sperm away from the testes, don't propel the sperm forward, as was once thought. Instead, the cilia agitate the sperm to keep them from aggregating and clogging the tubes so that they can reach their ultimate destination.

"Motile cilia are eyelashlike extensions of specific epithelial cells and have a beat that moves fluid over a surface," said Rex Hess, a University of Illinois professor emeritus of comparative biosciences and a major collaborator on the study led by Dr. Wei Yan, a foundation professor of physiology and cell biology at the University of Nevada, Reno School of Medicine.

"For more than 150 years, most papers and books on the subject stated that motile cilia of the efferent ductules move sperm cells in one direction, like they do in the female fallopian tubes," Hess said. "But we found that the cilia in this organ beat in an unusual manner that stirs and agitates the luminal fluid and sperm.

"The textbooks were wrong," Hess said. "Cilia in the male don't transport they keep the sperm dancing."

Contraction of smooth muscle lining the ductal walls moves the sperm into the epididymis.

"The research is also important because it focuses on microRNA, which is a product of a noncoding region of DNA," Yan said. "In our study, we found that just two clusters of five microRNAs control the proper formation of motile cilia. That is amazing."

When the Yan lab knocked out these microRNAs in mice, the researchers observed that the sperm aggregated into clumps that blocked the efferent ducts. This led to a backup of fluid in the testes, causing male infertility.

"Many infertile men may have clogged efferent ductules, but unfortunately this condition has been ignored due to a lack of knowledge on this important structure," Yan said. "Our discovery has significant translational potential and may one day lead to new methods that can help patients."

II. Method of locomotion

The above describe organ beat in a different way causing different types of movement in protozoans, so protozoans have several types of movement such as amoeboid, flagellar, ciliary, and metabolic movement. Some of the protozoans movements are described here –

1- Amoeboid movement

Sarcodina, certain Mastigophora, and Sporozoa have characteristic amoeboid movement. The process of amoeboid movement is done by pseudopodia formations, pseudopodia are formed by streaming flow of cytoplasm in the direction of movement.

2- Flagellar movement

Flagellar movement is present in Mastigophora, which bears one or more flagellum. There are three types of flagellar movement that are recognized.

A- Paddle stroke

This type of flagellar movement is first described by Ulehla and Krijsman in 1925. They describe that in this flagellar movement of the flagellum is sideway consist of effective stroke or down-stroke in the opposite direction of movement and relaxed recovery stroke, during recovery stroke flagellum brought forward again and ready for next effective stroke. As flagella give effective stroke in water in backward direction then water propels organism in the forward direction.

B- Undulating motion

In this type of movement wave-like undulation takes place from base to tip or from tip to base. If wave-like undulation takes place from tip to base, the animal is pulled in the forward direction, and if wave-like undulation takes place from base to tip animal is pulled in the backward direction. And when undulation is spiral animals rotate.

C- Simple conical gyration

It is described in Butschli’s screw theory, this theory postulates spiral turning like a screw. This screw-like motion causes the pulling of the animal in the forward direction with spiral rotation as well as gyration of the animal. Although the exact mechanism for this type of flagellar beat is unknown, it is believed that axonemal fibers are involved in this process. Sliding tubules theory describe, doublet slide past each other, which is the cause of movement in flagella, and energy for this process is mitochondrial ATP.

3- Ciliary movement

In the case of ciliary movement, the cilia oscillate in a pendulum-like manner. In each oscillation, there is a fast effective stroke followed by the recovery stroke, like flagellar movement. During effective stroke cilia expel the water in the backward direction like an oar of the boat, and in response if this effective stroke water propels the animal in the forward direction. During recovery stroke, cilia come in forward direction ready for next effective stroke. Cilia neither beat simultaneously nor independently, cilia beat progressively in a characterized wave-like manner.

Mode of swimming by cilia

By ciliary movement animal directly does not follow the straight movement, they rotate spirally like a bullet of rifle in left-handed helix manner. It might be because cilia do not beat directly straight, beating is somehow obliquely toward the right and might be cilia at oral groove beat more obliquely and vigorously away from the mouth. This combined effect causes swimming movement in the animal.

4- Metabolic movement

This is due to the pellicular contractile structure. In this type of movement, organisms show gliding or wriggling, or peristalsis. Microtubules present in their pellicle is responsible for this type of movement.

When cilia lose the beat

Normal cilia beat smoothly (top), but hydin-lacking cilia (bottom) are stiff.

Beating cilia circulate cerebrospinal fluid through the brain. If they falter, fluid can accumulate, causing the brain-damaging condition hydrocephalus. Mice carrying mutations in the gene for the protein hydin develop hydrocephalus, but it wasn't known why. Last year, the researchers discovered a clue by finding that the absence of hydin caused flagella in the protist Chlamydomonas to freeze up. Now the team has observed a similar effect in the cilia of mice.

Cilia from mice with Hydin mutations beat abnormally. Instead of showing a smooth back-and-forth movement, the cilia merely vibrated. They also beat more slowly and often stopped. Though the precise function of the hydrin protein is unclear, it is known to be part of the cilia's axoneme. Indeed, the Hydin mice also had a subtle structural defect in their cilia—a knob on one of the central microtubules was absent.

The authors believe that this defect might cause a lack of coordination among the dynein motors that move cilia. Normally, motors on one side of the shaft kick into gear to produce the forward stroke. Then they flip off and motors on the opposite side turn on, producing the return stoke. But the researchers suspect that this switch doesn't occur in hydin-lacking cilia, causing the shaft to relax prematurely. The results predict that humans with defects in hydin will develop hydrocephalus and primary ciliary dyskinesia, an inherited disorder caused by defective airway cilia.

When Cilia Go Wrong

Fully functional cilia are important for healthy respiratory function. When cilia do not function properly, a medical condition called bronchitis can develop. Bronchitis is the inflammation of the respiratory lining. Mucus and debris accumulate, resulting in constriction of the airways. Breathing becomes difficult, phlegm accumulates and coughing occurs. Heavy smokers have a hard time recovering from bronchitis, because the chemicals in cigarette smoke damage the cilia that line the airways. Eventually the number of cilia is reduced and those that remain may stop functioning altogether.