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ubiquitous coralline-algae surfaces, leaving tiny feeding scars like the adults. As it grows, the juvenile starfish adds arms to reach its final number and begins feeding on hard corals. How do they reproduce? Unlike many other starfish, the general body surface of crown-of-thorns starfish is soft and flexible, which enables it to swell as it develops huge gonads. Fecundity is obviously related to size, but, as an example, a 40 cm diameter crown-of-thorns starfish may commit about 45% of its total body energy to reproduction and shed an astonishing 50 million eggs. Like many marine invertebrates, crown-of-thorns starfish shed their gametes freely into the ocean and the gametes are wasted if there is no synchrony or proximity in spawning. Crown-of-thorns starfish don’t use precise cues for spawning: they may spawn at any stage of the lunar cycle and even join with other reef invertebrates in multi-species spawnings. They do, however, tend to spawn when the water temperature is about 28°C and often aggregate, apparently due to a spawning pheromone. Proximate spawnings of male and female crown-of-thorns starfish achieve almost 100% fertilisation. Even two crown-of-thorns starfish spawning 60 meters apart can achieve 23% fertilisation, resulting from the vast numbers of sperm released. Does this colossal reproductive capacity explain the ‘plagues’, then? Partially, yes. One must indeed go back through the lifecycle and consider the survival levels of the >108 eggs released by some crown-of-thorns starfish populations. It was observed that crown-of-thorns starfish plagues tended to occur three years after heavy rainfall and terrestrial runoff. Three years is about what it takes the crown-of-thorns starfish to grow to a point where they and their feeding traces become conspicuous. This suggests that the input of run-off nutrients influenced the survival and development of starfish larvae, by promoting higher levels of phytoplankton. So, ultimately, humans do influence the plagues: bad land-use practices in
areas adjacent to coral reefs lead to greater terrestrial run-off. In fact, much of the general deterioration of coral reefs internationally is due to these bad land-use practices. How can we get rid of them? Once large populations of crown-of-thorns starfish are observed on a reef it is extremely difficult to eliminate them and, even more, to eliminate them before they eat themselves out of coral (Figure 1). Between 1970 and 1983, almost 13 million crown-ofthorns starfish were removed from the reefs of the Ryukyu Islands, southern Japan, via a bounty for fishers, who changed from fishing to a more reliable income. Despite this huge effort, there are still large crown-of-thorns starfish populations in the Ryukyus. Complete removal is needed because cutting them up in situ isn’t the end: the pieces regenerate to make even more starfish! Successful control programs have only been achieved where there was a relatively small discrete population which was tackled quickly. Comprehensive control of crown-of-thorns starfish must eliminate the sources of nutrient input that promote survival of the larvae. Like all profound environmental problems, it is difficult and expensive to solve. The Australian government invested hundreds of millions of dollars for a Reef Rescue program over the past five years, and the program is now extended for another five years with further funding. Where can I find out more?
Birkeland, C., and Lucas, J.S. (1990). Acanthaster planci: Major Management Problem of Coral Reefs (Florida: CRC Press). Brodie, J., Brodie, Fabricius, K., De’ath, G., and Okaji, K. (2005). Are increased nutrient inputs responsible for more outbreaks of crown-ofthorns starfish? An appraisal of the evidence. Mar. Poll. Bull. 51, 266–278. De’ath, G., Fabricius, K.E., Sweatman, H., and Puotinen, M. (2012). The 27-year decline of coral cover on the Great Barrier Reef and its causes. Proc. Acad. Natl. Sci. USA 109, 17995–17999. Moran, P.J. (1986). The Acanthaster phenomenon. Oceanogr. Mar. Biol. 24, 379–480. Yasuda, N., Nagai, S., Hamaguchi, M., Okaji, K., Gerard, K., and Nadaoka, K. (2009). Gene flow of Acanthaster planci (L.) in relation to ocean currents revealed by microsatellite analysis. Mol. Ecol. 18, 1574–1590.
Centre for Marine Science, School of Biological Sciences, The University of Queensland, St Lucia, Queensland, 4072, Australia. E-mail:
[email protected]
Microtubuleregulating kinesins Emma G. Sturgill and Ryoma Ohi* Kinesins can regulate microtubule dynamics? The conventional function of kinesins, much like a molecular freight train, is to transport cargo by motoring along a microtubule (MT) track. But there exists another class of kinesins whose job relates more to track maintenance than transportation. These ‘regulatory kinesins’ modify the track on which they walk in order to shape the MT cytoskeleton. Regulatory kinesins control MT assembly and/or disassembly in order to influence the organization and dynamics of MT-based cellular machines. In other words, regulatory kinesins reconfigure the layout of the ‘rail map’. How many kinesins do this? Of the ~45 kinesins encoded by the human genome, 9 are known to regulate microtubule dynamics. Microtubule-regulating kinesins stratify into three basic classes: elongases, pause factors, and depolymerases (Figure 1). The kinesin7 CENP-E has been shown to promote microtubule elongation, suggesting that it may function as an elongase. kinesin-4s and -8s function as pause, or assembly-attenuating, factors and this class includes the mammalian motors Kif4/Xklp1 (a kinesin-4) and Kif18A (a kinesin-8). The largest and most studied class of regulatory kinesins, the depolymerases, is made up of members of the kinesin-8, -13, and -14 families. Specific examples include MCAK/Kif2C (a kinesin-13), yeast Kip3 (a kinesin-8) and Kar3 (a kinesin-14). It is worth noting that some kinesins indirectly impact MT dynamics (e.g., kinesin-1s promote MT elongation by activating JNK and delivering MT assembly factors to plus-ends). However, this Quick Guide focuses on kinesins whose motor activity directly alters MT dynamics. When did they start doing that? Evolutionarily, regulatory kinesins are conserved throughout the eukaryotic kingdom. Two out of six kinesins (Kar3
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and Kip3) in budding yeast directly regulate MT dynamics. Interestingly, one eukaryote, Theileria annulata, encodes just two kinesins — a kinesin-8 and a kinesin-13 — suggesting that microtubule regulation is an ancient and critical function of kinesin-like motors (Claire Walczak, personal communication). Historically, kinesins were first discovered to have MT-regulating capabilities in the mid 1990s. It is reasonable to assume that the list of regulatory kinesins is incomplete, as regulatory functions are still being uncovered for specific kinesins. For example, the ability of Kif19 to depolymerize MTs was discovered last year. What cellular processes do these motors control? As evidenced by their conservation from yeast to human, regulatory kinesins contribute to essential cellular processes. You can bet that regulatory kinesins are involved any time a cellular activity capitalizes on MT dynamics. One job of regulatory kinesins is to build MT-based cellular machines, such as the mitotic spindle. Compared to the interphase array, the spindle is composed of a copious amount of short dynamic MTs. Upon mitotic entry, depolymerases like Kif2A and MCAK must ramp up the MT catastrophe frequency in order to shift the distribution of MT number and length. In this manner, regulatory kinesins contribute to the massive rearrangement of the MT cytoskeleton necessary for spindle assembly. In addition to assembling MT-based structures, regulatory kinesins finetune the dynamics of these cellular machines for optimized performance. In the mitotic spindle, for example, Kif18A dampens kinetochore-MT dynamics to prevent excessive chromosome movement and MCAK dismantles flawed kinetochore-MT attachments. In mouse epithelial cells, the kinesin-8 Kif19 prevents excessive elongation of motile cilia. And kinesin-13s in Giardia and Leishmania coordinate with intraflagellar transport machinery to control cilia length. Each of these examples demonstrate how cells utilize regulatory kinesins to modulate the dynamic instability inherent to MTs in order to fine-tune cellular processes involving the cytoskeleton.
Subunit loss enhanced by: Kinesin-8s (Kip3, Kif19) Kinesin-13s (MCAK/Kif2C) Kinesin-14 (Kar3)
Suppression of dynamics: Kinesin-4 (Kif4/Xklp1) Kinesin-8s (Kif18A)
Subunit addition enhanced by: Kinesin-7 (CENP-E)
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Figure 1. Microtubule-regulating kinesins. Regulatory kinesins partition into three classes. Depolymerases (red) promote microtubule disassembly. Elongases (green) promote microtubule assembly. Pause factors (blue) suppress the inherent dynamicity of microtubules.
Why use motors when the cell contains so many other regulators of microtubule dynamics? Actually, some scientists speculate that kinesins were originally selected for their ability to regulate MT dynamics, and that motility evolved later. This is because dynamic polymers preceded motor proteins evolutionarily, so proteins that bound these polymers might have coupled to polymer dynamics initially. Nonetheless, there are advantages to having a motor domain on a regulatory factor. For example, the processivity of Kif18A is exquisitely tuned to get the motor to the plusends of kinetochore fibers, which are built from spindle microtubules that attach to kinetochores. This enables Kif18A to ‘measure’ MT length, so that long MTs are affected more than short MTs. How do kinesins regulate microtubule dynamics? In general, depolymerases can shrink MTs by removing tubulin from MT ends or, for dynamic MTs, by suppressing subunit addition. The latter mechanism probably works because it promotes loss of the GTP-tubulin cap. Similarly, elongases
can elongate MTs by adding tubulin to MT ends or by preventing catastrophes. The mechanistic details of these activities are poorly understood, but it is appreciated that regulatory kinesins can utilize their motor domains to alter the structure of MT protofilaments. MCAK, the best-described regulatory kinesin, stabilizes the bent conformation of protofilaments that predisposes MTs to depolymerization. In contrast, CENP-E has been proposed to stabilize the straight conformation of protofilaments that predisposes MTs to polymerization. But how does one kinesin use its motor domain to move while another uses it to manipulate MT dynamics? While all kinesins share a similar motor domain, structural differences can lead to variations of the ATPase cycle and create additional tubulin contact points. For example, the mechanochemical cycle of MCAK is limited by ATP hydrolysis rather than ADP release. This results in MCAK diffusing along the MT lattice instead of walking. The MT end stimulates MCAK to exchange nucleotides, transitioning the motor to a tightly
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bound state. Taken together, these unique features of the MCAK motor domain tune it to identify and stabilize curved protofilaments at MT ends. Notably, our current knowledge on how kinesins alter MT dynamics is heavily influenced by work on kinesin-13s, largely because mechanistic details of how other kinesins work do not exist. We await further studies to see if the kinesin13 paradigm is universal, or if other kinesins use unique biochemistries to shape the MT cytoskeleton.
Correspondence
Independent acquisition of sodium selectivity in bacterial and animal sodium channels
Where can I find out more?
Benjamin J. Liebeskind1,*, David M. Hillis1, and Harold H. Zakon1,2,3
Department of Cell and Developmental Biology, Vanderbilt University Medical Center, Nashville, TN 37232, USA. *E-mail:
[email protected]
Electrical signaling in animal nerves and muscles is largely carried out by proteins in the superfamily of voltagegated ion channels [1]. These proteins are based on a single homologous domain, but different types exist as single-domain tetramers, two-domain dimers, or four-domain proteins that comprise the whole pore-forming structure [1]. Four-domain channels are hypothesized to have evolved from a single-domain ancestor by two rounds of internal duplication [2]. The role that a channel plays in a cell’s physiology is largely determined by its selectivity for specific ion species and by the stimulus that opens the channel — its method of ‘gating’. The voltage-gated sodium (Nav) and calcium channels (Cav), which drive the upstroke of action potentials and transduce electrical signals into cellular signals, respectively, both have the four-domain architecture, whereas voltage-gated potassium channels (Kv) have only one domain. Crystallographic studies have led to important discoveries about ion permeation and gating in the single domain Kv channels, but structural studies of the four-domain Nav and Cav channels have not achieved the same level of precision [3], leaving the atomic details of these important proteins in the dark. The recent discovery of and subsequent crystallographic work on a voltage-gated, sodium-selective, singledomain channel in bacteria (BacNav) was therefore greeted with excitement as a potential model of four-domain Nav channels [4–6]. The selectivity filter of BacNav channels is very different from that of eukaryotic Nav channels, however, and these studies often lack clear statements of homology between the two channel types [4–6]. BacNav channels are often referred to as
Bringmann, H., Skiniotis, G., Spilker, A., Kandels-Lewis, S., Vernos, I., and Surrey, T. (2004). A kinesin-like motor inhibits microtubule dynamic instability. Science 303, 1519–1522. Desai, A., Verma, S., Mitchison, T.J., and Walczak, C.E. (1999). Kin I kinesins are microtubule-destabilizing enzymes. Cell 96, 69–78. Endow, S.A., Kang, S.J., Satterwhite, L.L., Rose, M.D., Skeen, V.P., and Salmon, E.D. (1994). Yeast Kar3 is a minus-end microtubule motor protein that destabilizes microtubules preferentially at the minus ends. EMBO J. 13, 2708–2713. Friel, C.T., and Howard, J. (2012). Coupling of kinesin ATP turnover to translocation and microtubule regulation: one engine, many machines. J. Muscle Res. Cell Motil. 33, 377–383. Gardner, M.K., Zanic, M., Gell, C., Bormuth, V., and Howard, J. (2011). Depolymerizing kinesins Kip3 and MCAK shape cellular microtubule architecture by differential control of catastrophe. Cell 147, 1092–1103. Gupta, M.L., Jr., Carvalho, P., Roof, D.M., and Pellman, D. (2006). Plus end-specific depolymerase activity of Kip3, a kinesin8 protein, explains its role in positioning the yeast mitotic spindle. Nat. Cell Biol. 8, 913–923. Howard, J., and Hyman, A.A. (2009). Growth, fluctuation and switching at microtubule plus ends. Nat. Rev. Mol. Cell Biol. 10, 569–574. Sardar, H.S., Luczak, V.G., Lopez, M.M., Lister, B.C., and Gilbert, S.P. (2010). Mitotic kinesin CENP-E promotes microtubule plus-end elongation. Curr. Biol. 20, 1648–1653. Stumpff, J., Du, Y., English, C.A., Maliga, Z., Wagenbach, M., Asbury, C.L., Wordeman, L., and Ohi, R. (2011). A tethering mechanism controls the processivity and kinetochoremicrotubule plus-end enrichment of the kinesin-8 Kif18A. Mol. Cell 43, 764–775. Su, X., Ohi, R., and Pellman, D. (2012). Move in for the kill: motile microtubule regulators. Trends Cell Biol. 22, 567–575. Varga, V., Helenius, J., Tanaka, K., Hyman, A.A., Tanaka, T.U., and Howard, J. (2006). Yeast kinesin-8 depolymerizes microtubules in a length-dependent manner. Nat. Cell Biol. 8, 957–962. Walczak, C.E., Cai, S., and Khodjakov, A. (2010). Mechanisms of chromosome behaviour during mitosis. Nat. Rev. Mol. Cell Biol. 11, 91–102. Wordeman, L. (2010). How kinesin motor proteins drive mitotic spindle function: Lessons from molecular assays. Semin. Cell Dev. Biol. 21, 260–268.
‘ancestors’ of Nav channels [5], a claim whose evolutionary meaning is difficult to interpret. Basic research on the organismal function of BacNav channels, moreover, has lagged behind the sophisticated structural studies. This situation leaves it unclear whether the molecular correlates of function are truly comparable between eukaryotic Nav and BacNav channels. We help address this by grounding the relationship of BacNav channels to other major channel groups in an evolutionary framework. The constituent domains of fourdomain channels have what may be called molecular serial homology, where all four domains are equally related to the single-domain precursor [2]. We therefore followed the procedure of Strong et al. [2] and broke the fourdomain channels into their constituent domains, making the smallest homologous unit (the domain) into the operational taxonomic units in the phylogeny. Figure 1 shows strong support for the traditional view of ion channel evolution [2], with a single origin of the four-domain structure in Nav and Cav channels. DI and DIII form a clade, as do DII and DIV, in keeping with the hypothesis of two sequential rounds of internal gene duplication [2]. BacNav channels fell outside the four-domain group with strong support, rejecting the notion that BacNav channels can be considered Nav channels [4] in an evolutionary sense. Instead, they grouped near CatSper channels, consistent with earlier studies showing that both BacNav and CatSper channels are used as pH sensors in the bacterial and sperm cells in which they are respectively expressed [7,8]. We therefore propose that the BacNav, CatSper, and the novel single-domain protist types be viewed provisionally as a pH-gated group, based both on evolutionary relatedness and conservation of function. This tree rejects the possibility of BacNav channels being placed within Nav channels, but it is still possible that BacNav are functionally similar to the precursors of animal Nav channels. There are two mutually exclusive hypotheses about the evolution of ion selectivity in voltage-gated ion channels (Figure S1 in Supplemental Information, published with this article online). In one scenario (Figure S1A), sodium selectivity is independently acquired in BacNav and animal Nav channels. In the other, BacNav channels are similar in function to the common ancestor
