Cilia and Flagella

Abstract

Eukaryotic cilia and flagella are hair‐like cellular appendages composed of specialised microtubules and covered by a specialised 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 and the enantiomorphic asymmetry (handedness) of the axoneme.
  • 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 of 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 gate 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 a billion years ago, following the appearance of the genes for tubulin (from bacteria) and proteins that establish the ninefold symmetry.

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

Figure 1. (a) High‐voltage electron micrograph (EM) of a sea urchin sperm with flagellar bends. Modified with permission from IR Gibbons, 1975 © Society of General Physiologists. (b) Scanning EM of a protozoan cell fixed to preserve metachronal waves of beating cilia. Courtesy of SL Tamm. (c) and (d) Cross‐sectional EMs of a basal body and a 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. Reproduced with permission from Linck and Stephens, 2007 © Wiley‐Liss, Inc. Bar, 5 µm for (a), 10 µm for (b) and 0.94 µm for (c) and (d).
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 specialised attachments (Figure). Similarly, all dynein arms are not identical, for example, in some species, the arms form a specialised bridge between doublets 5 and 6 (see also Figure). The actual positions and shapes of inner dynein arms and nexin links are not precisely drawn (for details see below). The central pair singlet microtubules (1–2) possess a complex array of elements that are highly species‐specific. For further description, see Figure. (b) Model of an A‐microtubule with associated dynein arms and radial spokes. Outer arms are arranged with a 24‐nm axial spacing. Inner arms and nexin links have a more complex arrangement, but an overall 96‐nm axial repeat: the tripartite I1 complex is shown in blue, the DRC‐nexin complex 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 ( ) to humans, whereas in the protist only the base of S3 is present (Lin ., ). The spokes are attached relative to the inner dynein arm components as shown, and relative to the polarity (+ end) of the microtubule. (Modified with permission from Heuser et al. J. Cell Biol. 2009;52:66–83. doi: 10.1083/jcb.52.1.66 © Rockefeller University Press.)
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 µm 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 sulphate 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. Reproduced with permission from Piperno G et al. 1977 © G Piperno.
Figure 5. (a) Flagellar doublet microtubules from sea urchin (e.g. ) sperm (and from flagella, not shown) can be fractionated into stable ribbons (R) of 3‐protofilmaents (PFs, three vertical lines), shown by negative stain EM. Partial extraction of ribbons with urea reveals a single, hyper‐stable tektin filament (F) that is colinear with the Ribbon. Bar, 100 nm. Reproduced from Linck . J. Biol. Chem. 2014;289:17427–17444. doi: 10.1074/jbc.M114.568949 © the American Society for Biochemistry and Molecular Biology. (b) By NaDodSO4/EP analysis, ribbons from and sea urchin flagella are composed of αβ‐tubulin and a specific subset of other proteins (named, or given as × 10−3). Ribbon (Rib) proteins include two Ca2+‐binding proteins that are named differently in different species (e.g. Rib72 in , Rib74 and Rib85.5 in , and efhc1 and efhc2 in mouse and human); here these are referred to as JME1 and JME2, given that their mutated, homologous genes cause Juvenile Myoclonic Epilepsy in humans (Linck ., ). Rib43a and Rib45 are also present, are evolutionarily conserved in humans, but have no known function. Ribbons from sea urchin and mammalian cilia and flagella can be further fractionated by extraction with 2 mol L−1 urea into filaments composed of equimolar tektins A, B and C (with molecular masses of approximately 50 kDa); however, tektin is solubilised from the ribbon by urea. (c) Diagram of the conserved structure of a ciliary AB‐doublet microtubule and a centriolar triplet microtubule (with C‐tubule shaded), based on data from Linck and Stephens (), Linck . () and Nicastro . (). Most or all of the numbered PFs are composed of tubulin. A single stable ribbon of 3‐PFs (black) from panel (a) corresponds to the partition PFs of the A‐tubule, that is, either A11‐12‐13 or A12‐13‐1. The partition‐associated material is shown in green; microtubule inner protein (MIP)‐1 is shown in blue; MIP‐2 in red and MIP‐3 in yellow. PF B10 is connected to A1 by a novel junctional protein shown in purple, and a presumably similar protein (light purple) connects PF C10 to B8 in centrioles. The tektin ABC filament is an integral part of the ribbon; JME1 and JME2 behave as a dimer and are also associated with the ribbon, but their precise location within the A‐tubule has not been determined. Approximate positions of the radial spokes (RS), inner dynein arms (IDA) and the dynein regulatory complex (DRC) are from Heuser . () and Nicastro . ().
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. Reproduced with permission from IR Gibbons, 1975 © Society of General Physiologists. (b) Left, model of three‐dimensional pattern of ciliary beat: the rapid effective stroke (5→1) is within the plane of beat, and the slower recovery stroke (1→5) 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 5→1 order) and their lines of synchrony change by 120°–150°. Reproduced with permission from Omoto and Kung. J. Cell Biol. 1980;87:33–46. doi:10.1083/jcb.87.1.33 © Rockefeller University Press.)
Figure 7. Highly simplified schematic of the dynein cross‐bridge 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 cross‐bridge 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 cross‐bridge is now ready to begin a new cycle. How one dynein cross‐bridge affects the phase of next dynein arms (?) is not known. The average translocation distance (δ) per cross‐bridge cycle has been measured to be 8 nm. See Carter . () for greater detail.
Figure 8. Polypeptide composition and structure of the dynein outer arm of . The large C‐terminal motor domains (and knob‐like projections) of the heavy chains (αβγ‐HCs) are oriented upward, with loosely associated 45‐kDa polypeptides. The approximate positional arrangements of the ICs (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. Reproduced with permission from Bernashski et al. (1999) © American Chemical Society.
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 (Δ) 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 Figureb, 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 with permission from Warner and Satir. J. Cell Biol. 1974;63:35–63. doi:10.1083/jcb.63.1.35 © Rockefeller University Press. See also Mitchell (), Wirschell . () and Yang and Smith ().
Figure 10. Basic principles of the ‘Geometric Clutch’ model for the generation of propagated flagellar bending waves (Lindemann, ). (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 ‐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 ‐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 . (Reproduced with permission from Lindemann and Mitchell 2007 © Wiley‐Liss, Inc.)
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Linck, Richard W(Jul 2015) Cilia and Flagella. In: eLS. John Wiley & Sons Ltd, Chichester. http://www.els.net [doi: 10.1002/9780470015902.a0001258.pub3]