Cilia and Flagella

Richard W Linck, University of Minnesota, Minneapolis, Minnesota, USA

Published online: December 2009

DOI: 10.1002/9780470015902.a0001258.pub2

Eukaryotic cilia and flagella are hair-like, cellular appendages composed of specialized microtubules and covered by a specialized extension of the cellular membrane. Their structure, genes, proteins and functions are highly conserved throughout evolution from protists to humans. Ciliary defects lead to physiological dysfunctions, developmental disorders and disease. Cilia and flagella have three, often interrelated functions: (1) As motile organelles beating like whips or oars, they propel cells through their environment or transport fluids along the surfaces of ciliated epithelia. (2) Both motile and nonmotile cilia act as antennae, sensing environmental cues and metabolic compounds, and initiating specific cellular responses. (3) Their microtubules act as railroad tracks, along which molecular motors transport other molecules out to the ciliary tip and back to the cell body – a process called intraflagellar transport. Given these functions, cilia and flagella are micromachines and they act as cybernetic devices to receive, process and communicate information.

Key Concepts

  • Structural concepts in ciliary/flagellar axoneme assembly and function include: the template function of the basal body, the polarity of the microtubules, the enantiomorphic asymmetry (handedness) of the axoneme and possibly the role of tektin filaments in positioning the effector molecules.
  • The assembly of the axoneme is tightly regulated by the expression of specific genes, by the limited amount of axonemal precursor proteins and by kinase enzymes.
  • The mechanochemical force for motility is provided by dynein arms (large multisubunit ATPase enzymes) that cause the doublet microtubules to slide past each other.
  • The beating cilia and flagella depends on many biochemical factors including: the different effects of outer versus inner dynein arm motors, the DRC (dynein regulatory complex) and DRC–radial spoke interactions mediated by kinases.
  • The waveform of beating cilia/flagella also depends on the precise geometric assemblage of the axoneme structures, the mechanical properties of those structures and principles of the Geometric Clutch hypothesis.
  • Associated with ciliary/flagellar membranes are numerous ion channels and signalling molecules.
  • Intraflagellar transport (IFT) involves anterograde and retrograde transport of specific molecules along the axoneme (via kinesin and dynein motors respectively), and it is an essential process for ciliary/flagellar assembly and their signalling functions.
  • The ciliary pore is formed by the membrane collar at the base of the cilium and by the stellate fibres of the basal body; it functions to sort, modify and permit entry of only membrane and protein constituents destined for transport and incorporation into the developing cilium.
  • Mutations in genes encoding structural and functional proteins of cilia and flagella lead to innumerable diseases and disorders called ciliopathies.
  • Eukaryotic cilia and flagella are estimated to have evolved roughly 850 million years ago following the appearance of the genes for tubulin (from bacteria), possibly for tektin and for proteins that establish the 9-fold symmetry.

Keywords: centriole; dynein; intraflagellar transport; kinesin; microtubule; motility; sensory reception; sperm; tektin; tubulin

Figure 1. (a) High-voltage electron micrograph (EM) of sea urchin sperm with flagellar bends. Courtesy of IR Gibbons. (b) Scanning EM of protozoa fixed to preserve metachronal waves of beating cilia. Courtesy of SL Tamm. (c) and (d) Cross-sectional EMs of basal body and flagellar 9+2 axoneme of protozoa, typical of most simple cilia and flagella (membrane removed and viewed from the proximal end looking towards the distal tip). Fixed with tannic acid to show the protofilament substructure of the microtubules. The triplet ABC and doublet AB microtubules provide support for associated elements. Bar, 5 m for (a), 10 m for (b) and 0.94 m for (c) and (d). Reproduced form Linck and Stephens (2007) with permission from Wiley-Liss Inc.
Figure 2. (a) Cross-sectional diagram of a 9+2 axoneme (viewed from the proximal end towards the distal tip). All doublet microtubules may not be identical, for example in some species doublets 3 and 8 possess specialized attachments (Figure 3). Similarly, all dynein arms are not identical, for example in some species the arms form a specialized bridge between doublets 5 and 6 (see also Figure 10). Actual position of inner dynein arms and nexin links are not precisely drawn. The central pair singlet microtubules (1–2) possess a complex array of elements that are highly species-specific. (b) Model of an A-microtubule with associated dynein arms and radial spokes (modified from Porter and Sale, 2000). Outer arms are arranged with a 24-nm axial spacing. Inner arms have a more complex arrangement, but an overall 96-nm axial repeat: the tripartite I1 complex is shown in blue, the DRC in green and other components in pink. Radial spoke triplets (S1/S2/S3), with evolutionarily conserved spacings of 32+24+40=96 nm, are present in most species from protists (Tetrahymena) to humans, whereas in the protist Chlamydomonas there are spoke pairs (S1/S2) with spacings of 32+64=96 nm. The spokes are attached relative to the inner dynein arm components as shown, and relative to the polarity (+ end) of the microtubule.
Figure 3. Morphology of a mammalian spermatozoon. All cross-sections are viewed from the proximal end towards the distal tip. A simple 9+2 axoneme extends the full length. Attached to each doublet microtubule is a long, tapering outer dense fibre (numbered). In the middle piece region a mitochondrial sheath surrounds the outer dense fibres. In the principal piece a fibrous sheath replaces the mitochondrial sheath; longitudinal columns of the fibrous sheath replace outer dense fibres 3 and 8, and are firmly anchored to outer doublet microtubules 3 and 8, preventing them from sliding. The head and tail of human spermatozoa measure approximately 5 and 55 m in length, respectively. Courtesy of DW Fawcett.
Figure 4. Two-dimensional separation of polypeptides of Chlamydomonas flagellar axonemes by isoelectric focusing (IEF) and sodium dodecyl sulfate polyacrylamide electrophoresis (NaDodSO4/EP). Over 250 polypeptides can be resolved by their isoelectric points (pH) and molecular weights (Mr×10–3). The 12 polypeptides (arrows) were shown to form the radial spokes by comparing axonemes from wild-type versus mutant cells lacking spokes. Courtesy of G Piperno.
Figure 5. Top: Doublet tubules from Chlamydomonas and sea urchin (e.g. S. purpuratus) can be fractionated into Sarkosyl-stable ribbons of 3-protofilaments, shown by SDS-PAGE to be composed of -tubulin and a specific subset of other proteins (named, or given as Mr×10–3). Ribbons can be further fractionated into filaments composed of tektins A (approximately 53 kDA), B (approximately 50 kDa) and C (47 kDa). Bottom: Diagram of the conserved structure of a ciliary doublet microtubule and a centriolar triplet microtubule (C-tubule shaded). Standard protofilament (PF) numbering scheme from Linck and Stephens (2007). All of the PFs have been presumed to be composed of tubulin; however, evidence indicates that tektins may form at least one of them (Setter et al., 2006). The B-11th and C-11th components are novel proteins connecting B10 to A1 and C10 to B8, respectively. A postulated position of the Sarkosyl-stable ribbon of three PFs is shown in black, with a tektin filament possibly forming one of the PFs. The locations of the other ribbon proteins (e.g. efhc1/rib72, efhc2 and rib43a) are not known. Approximate positions of radial spokes (RS), inner dynein arms (IDA) and dynein regulatory complex (DRC) are from Porter and Sale (2000) and Nicastro et al. (2006). Reproduced from Linck and Stephens (2007) with permission from Wiley-Liss Inc.
Figure 6. Flagellar versus ciliary beat patterns and waveforms. (a) Model showing a sea urchin sperm flagellum consisting of arcs, straight segments and accompanying rotational twists (+/– degrees) along the axoneme. Courtesy of I. R. Gibbons. (b) Left, model of three-dimensional pattern of Paramecium ciliary beat: the rapid effective stroke (51) is within the plane of beat, and the slower recovery stroke (15) twists out of the plane. Right, diagram of a ciliary metachronal wave. Cilia within a line of synchrony are all in the same phase of beat. In forward-swimming cells the firing order of cilia is towards the left, whereas the direction of metachronal wave propagation is from left to right; in backward-swimming cells (e.g. during a chemotactic signalling response and influx of Ca2+), the direction of effective strokes (still in a 51 order) and their lines of synchrony change by 120°–150°. From Omoto and Kung (1980).
Figure 7. Highly simplified schematic of the dynein crossbridge cycle leading to microtubule sliding, showing four basic states (a)–(d), and steps 1–4. A+, plus end of the A-tubule of one doublet microtubule; B+, plus end of the B-tubule of the adjacent doublet tubule. Dynein arms are represented by squares and rectangles, permanently anchored to the A-tubule. Starting at step 4: ATP binds to the first P-loop of a DHC, dissociating the crossbridge from the B-tubule. Step 1: The binding of ATP causes a change in the state of dynein, causing it to bind to the B-tubule. Step 2: Dynein hydrolyses ATP to ADP+Pi; the energy from hydrolysis is stored somehow in the motor domain of the DHC, but the products are not yet released. Step 3: Closely coupled to the release of ADP and Pi, a conformational change is thought to occur in the arm (leading to state d), resulting in the translocation or shear of the A-tubule in the minus direction with the arm acting as a mechanical lever. Step 4: The crossbridge is now ready to begin a new cycle. How one dynein crossbridge affects the phase of next dynein arms (?) is not known. The average translocation distance () per crossbridge cycle has been measured to be 8 nm. See Carter et al. (2008) for greater detail.
Figure 8. Polypeptide composition and structure of the dynein outer arm of Chlamydomonas (Bernashski et al., 1999); compared with Table 1. The large C-terminal motor domains (and knob-like projections) of the heavy chains (-HCs) are oriented upward, with loosely associated 45 kDa polypeptides. The approximate positional arrangements of the intermediate chains (IC1 and IC2) and light chains (1–8), many of which are associated with the N-terminals of the HCs, are shown. The types of some LCs are indicated, that is, LRR (leucine-rich repeat) LC1, Tctex2 LC2, thioredoxin LC3 and 5, and calmodulin-like LC4.
Figure 9. Radial spoke–central pair interactions during ciliary bending. Left: EM thin section through a ciliary bend and basal body connection at the bottom. Radial spoke triplets (l–8) are attached along two A-tubules (d1 and d5) and project inward to the central pair microtubule (CPM) complex. Proximally, sliding is prevented by the basal body–axoneme connection. Distally, progressive sliding produces accumulated positional differences () between each doublet tubule and the CPM. Right: diagrams (a)–(e) illustrate a series of interactions that might occur between spoke heads (S1–S3) and the CPM projections (1–9). Spokes are permanently anchored to the A-tubules in triplets, spaced as in Figure 2b, and spoke heads are free to slide past the CPM, spaced axially at 16 nm. Open spoke heads are out of phase with, and not attached to, the projections. As sliding takes place, spoke heads (solid) come into phase with and attach to specific projections, and may tilt, as sliding continues. Spoke–CPM interactions are thought to function both as mechanical constraints to sliding and as signalling devices that interact with the DRC to regulate dynein arm activities. Modified from Warner and Satir (1974).
Figure 10. Basic principles of the ‘Geometric Clutch’ model for the generation of propagated flagellar bending waves (Lindemann, 2007). (a) Doublet microtubules are prevented from freely sliding by their anchorage in the basal body (black end) and by elastic, resistive elements (nexin links and central pair-radial spoke interactions). (b) The dynein-driven sliding force between doublets 6–9, coupled with the shear resistance, generates tension between doublet tubules, resulting in the formation of a bend (as in (a)). This tension develops as a transverse t-force that acts to pull the dynein arms on doublets 1–4 away from their interacting B-tubules (as in (a) and (b)), locally inhibiting dynein-driven sliding on one side of the axoneme only and allowing the bend to propagate. (c) Eventually, as the axoneme bends, the t-force acts across the axoneme to deform it, pulling apart and inactivating the dyneins on doublets 6–9, and allowing the dynein arms on doublets 1–4 to engage and generate a reverse bend (not shown). In computer models, if the tip of the axoneme is anchored instead, the same mechanism will generate and propagate a bend in the opposite direction. Such a result has been shown experimentally, and can explain the behaviour of Crithidia. Figure courtesy of Charles B. Lindemann. Reproduced from Lindemann and Mitchell (2007) with permission from Wiley-Liss Inc.
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Related Sub-Topics:

Cellular Transport | Intracellular and Extracellular Signalling | Cell Motility | Cell Motility and Transport | Information Flow | Molecular Transport | Biochemistry of Cell Motility
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Article Title: Cilia and Flagella
 
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Linck, Richard W(Dec 2009) Cilia and Flagella. In: eLS. John Wiley & Sons Ltd, Chichester. http://www.els.net [doi: 10.1002/9780470015902.a0001258.pub2]