- Email: [email protected]
Quality Control: Linking Retrotranslocation and Degradation
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Arabidopsis and rice genomes [18] suggests that lineage-specific tinkering with the composition of the plastid proteome can occur even over short evolutionary timescales. In the mean-time, multiple algal genome sequences are now publicly available — for example, those of the green algae Chlamydomonas and Ostreococcus (http://genome. jgi-psf.org/euk_cur1.html) and the red alga Cyanidioschyzon (http:// merolae.biol.s.u-tokyo.ac.jp) — and it is likely that ten or more plant and algal nuclear genome sequences will be available for even more thorough and systematic analyses within the next few years. An increasing number of genomes from ‘secondary’ plastid-containing algae such as diatoms, cryptophytes and chlorarachniophytes will also be completely sequenced (http:// www.jgi.doe.gov/). These organisms acquired their plastids through the engulfment of red or green algal endosymbionts [19] and the molecular dynamics accompanying the process of secondary endosymbiosis, in which gene transfers between evolutionarily distinct nuclear genomes are also a possibility, adds another layer of complexity to an already complicated picture [20]. Overall, it is sobering to consider how little we know about the nuts and bolts of endosymbiosis and the full scope of its role in the diversification of eukaryotic cells. References 1. Yoon, H.S., Hackett, J.D., Ciniglia, C., Pinto, G., and Bhattacharya, D. (2004). A molecular timeline for the origin of photosynthetic eukaryotes. Mol. Biol. Evol. 21, 809–818. 2. Reyes-Prieto, A., Hackett, J.D., Soares, M.B., Bonaldo, M.F., and Bhattacharya, D. (2006). Cyanobacterial contribution to algal nuclear genomes is primarily limited to plastid functions. Curr. Biol. 16, 2320–2325. 3. Martin, W., Rujan, T., Richly, E., Hansen, A., Cornelsen, S., Lins, T., Leister, D., Stoebe, B., Hasegawa, M., and Penny, D. (2002). Evolutionary analysis of Arabidopsis, cyanobacterial, and chloroplast genomes reveals plastid phylogeny and thousands of cyanobacterial genes in the nucleus. Proc. Natl. Acad. Sci. USA 99, 12246–12251. 4. Weeden, N.F. (1981). Genetic and biochemical implications of the endosymbiotic origin of the chloroplast. J. Mol. Evol. 17, 133–139.
5. Timmis, J.N., Ayliffe, M.A., Huang, C.Y., and Martin, W. (2004). Endosymbiotic gene transfer: organelle genomes forge eukaryotic chromosomes. Nat. Rev. Genet. 5, 123–135. 6. Bruce, B.D. (2000). Chloroplast transit peptides: structure, function and evolution. Trends Cell Biol. 10, 440–447. 7. McFadden, G.I. (1999). Plastids and protein targeting. J. Eukaryot. Microbiol. 46, 339–346. 8. Soll, J., and Schleiff, E. (2004). Protein import into chloroplasts. Nat. Rev. Mol. Cell Biol. 5, 198–208. 9. Martin, W., and Schnarrenberger, C. (1997). The evolution of the Calvin cycle from prokaryotic to eukaryotic chromosomes: a case study of functional redundancy in ancient pathways through endosymbiosis. Curr. Genet. 32, 1–18. 10. The Arabidopsis Genome Initiative (2000). Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature 408, 796–815. 11. Karol, K.G., McCourt, R.M., Cimino, M.T., and Delwiche, C.F. (2001). The closest living relatives of land plants. Science 294, 2351–2353. 12. Rodriguez-Ezpeleta, N., Brinkmann, H., Burey, S.C., Roure, B., Burger, G., Lo¨ffelhardt, W., Bohnert, H.J., Philippe, H., and Lang, B.F. (2005). Monophyly of primary photosynthetic eukaryotes: green plants, red algae, and glaucophytes. Curr. Biol. 15, 1325–1330. 13. Hall, W.T., and Claus, G. (1963). Ultrastructural studies on the blue-green algal symbiont in Cyanophora Paradoxa Korschikoff. J. Cell Biol. 19, 551–563. 14. Steiner, J.M., Bergho¨fer, J., Yusa, F., Pompe, J.A., Klo¨sgen, R.B., and Lo¨ffelhardt, W. (2005). Conservative sorting in a primitive plastid. The cyanelle
15.
16.
17.
18.
19.
20.
of Cyanophora paradoxa. FEBS J. 272, 987–998. Lo¨ffelhardt, W., Bohnert, H.J., and Bryant, D.A. (1997). The complete sequence of the Cyanophora paradoxa cyanelle genome. In Origins of Algae and Their Plastids, D. Bhattacharya, ed. (Wien: Springer), pp. 149–162. Martin, W., Stoebe, B., Goremykin, V., Hansmann, S., Hasegawa, M., and Kowallik, K.V. (1998). Gene transfer to the nucleus and the evolution of chloroplasts. Nature 393, 162–165. De Bodt, S., Maere, S., and Van de Peer, Y. (2005). Genome duplication and the origin of angiosperms. Trends Ecol. Evol. 20, 591–597. Richly, E., and Leister, D. (2004). An improved prediction of chloroplast proteins reveals diversities and commonalities in the chloroplast proteomes of Arabidopsis and rice. Gene 329, 11–16. Bhattacharya, D., Yoon, H.S., and Hackett, J.D. (2003). Photosynthetic eukaryotes unite: endosymbiosis connects the dots. Bioessays 26, 50–60. Archibald, J.M. (2005). Jumping genes and shrinking genomes—probing the evolution of eukaryotic photosynthesis with genomics. IUBMB Life 57, 539–547.
The Canadian Institute for Advanced Research, Program in Evolutionary Biology, Department of Biochemistry and Molecular Biology, Dalhousie University, Sir Charles Tupper Medical Building, 5850 College Street, Halifax, Nova Scotia, B3H 1X5, Canada. E-mail: [email protected] DOI: 10.1016/j.cub.2006.11.008
Quality Control: Linking Retrotranslocation and Degradation Misfolded proteins in the ER require the p97 AAA ATPase for dislocation across the membrane prior to degradation by the cytosolic proteasome. The mechanism by which dislocated proteins are delivered to the proteasome from p97 is unclear, but recent studies suggest an important regulatory role for the protein ataxin-3. Colin J. Stirling1 and J. Michael Lord2 The fatal neurodegenerative Machado-Joseph disease, also known as spinocerebellar ataxia type 3, is caused by mutations of the polyglutamine-containing protein ataxin-3 [1]. The biochemical properties of ataxin-3 are well known, but its physiological role has been elusive. Two recent studies [2,3] now implicate ataxin-3 in the process known as endoplasmicreticulum-associated degradation
(ERAD), suggesting a link between endoplasmic reticulum (ER) stress and the neuropathology associated with disease. After targeting to the ER, proteins are screened by a quality control system to prevent misfolded forms from progressing through the secretory pathway. Rather than accumulating within the cell, these aberrant proteins are disposed of by ERAD. This requires the dislocation of such proteins across the ER membrane and their subsequent degradation
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by the cytosolic proteasome system. The precise nature of the membrane channel through which dislocation occurs is not yet known, but a number of proteins have been implicated, including Sec61, Derlin-1 and VIMP [4]. However, evidence suggests that the hexameric AAA ATPase p97/VCP (known as Cdc48 in yeast) plays an important role in driving the dislocation of a range of proteins [5–8]. A complex comprising p97, and the adaptor proteins Ufd1 and Npl4, is recruited to the cytosolic face of the ER membrane, where it is believed to ‘pull’ dislocating polypeptides through the channel and into the cytosol. During their dislocation, ERAD substrates are normally polyubiquitinated, and the p97 complex has been shown to bind directly to these polyubiquitin chains. A further set of polyubiquitin-binding proteins, including Rad23p and Dsk2p, are associated with the proteasome, where they function in the final delivery of the ERAD substrate to the degradation machinery [9]. The mechanism by which an ERAD substrate is transferred from the p97 complex to proteasomal polyubiquitin-binding proteins is not known. Previous studies have shown that ataxin-3 can interact with both p97/VCP and Rad23p [10,11], and two recent papers [2,3] suggest that ataxin-3 is a critical link in this chain. Wang et al. [2] and Zhong and Pittman [3] have studied the effects of the expression of wild-type and mutant forms of ataxin-3 on a range of ERAD substrates. Both studies confirm that full-length ataxin-3 binds directly to p97, but ataxin-3 was also found in a membrane-associated form that could be co-immunoprecipitated together with several other ERAD components, including Derlin-1, VIMP and the ER-specific ubiquitin ligase Hrd1p [2]. Previous studies have shown that ataxin-3 has three functional ubiquitin-interacting modules plus a deubiquitinating activity [12–15]. These new studies [2,3] provide evidence that ataxin-3, and specifically its deubiquitinating activity, are required in ERAD. The
ataxin-3 C14A mutant lacks the catalytic deubiquitinating activity but is still able to bind ubiquitinated substrates with high affinity. Ectopic expression of ataxin-3 C14A in transfected cells led to the accumulation of typical ERAD substrates, including TCRa, a membrane component of the T-cell receptor which is degraded in the absence of other T-cell receptor subunits. The accumulated forms of TCRa were found to be extensively polyubiquitinated in atx C14A cells, suggesting a role for ataxin-3 in the deubiquitination of substrate during the normal ERAD process. This was further confirmed by the demonstration that p97-bound substrate could be deubiquitinated in vitro in the presence of wild-type ataxin-3 but not the C14A mutant. Consistent with this, Wang et al. [2] further demonstrated a dramatic increase in the amount of ubiquitinated proteins that could be co-immunoprecipitated with p97in ataxin-3 C14A cells compared to controls. Zhong and Pittman [3] further examined the effect of pathological mutations in the polyglutamine tract of ataxin-3. Their studies show that mutant ataxin-3 binds p97/VCP more avidly than the wild-type protein, and this correlates with decreased degradation of the ERAD substrate CD3d [3]. These studies suggest that the pathology of ataxin-3 mutants relates to defective ERAD. Taken together, the new data [2,3] indicate that ataxin-3 plays a critical role in the p97-associated degradation of ERAD substrates, and that this is necessary for the efficient release of substrate from the p97 dislocation machinery. Complete deubiquitination of substrate would prevent subsequent interaction with proteasomal polyubiquitin-binding proteins and so it is proposed that ataxin-3 might function as an ‘editing enzyme’ that merely shortens polyubiquitin chains. In this model the normal deubiquitinating activity of ataxin-3 would trim polyubiquitin chains to a level that allows their release from the p97 complex, but still permits binding to crucial proteasomal components.
Interestingly, the ubiquitin chains of ERAD substrates in Saccharomyces cerevisiae are progressively shortened as they transit to the proteasome [16]. Such a role for mammalian ataxin-3 would require tight regulation since any change in the level of deubiquitinating activity might be expected to perturb ubiquitin trimming and thus prevent efficient transfer of substrates to the proteasome. Indeed, ectopic over-expression of even wild-type ataxin-3 resulted in the accumulation of TCRa and CD3d [2,3], suggesting that this molecule is an important regulator of mammalian ERAD. References 1. Kawaguchi, Y., Okamoto, T., Taniwaki, M., Aizawa, M., Inoue, M., Katayama, S., Kawakami, H., Nakamura, S., Akiguchi, L., et al. (1994). CAG expansions in a novel gene for the Machado-Joseph disease at chromosome 14q32.1. Nat. Genet. 8, 221–229. 2. Wang, Q., Li, L., and Ye, Y. (2006). Regulation of retrotranslocation by p97-associated deubiquitinating enzyme ataxin-3. J. Cell Biol. 174, 963–971. 3. Zhong, X., and Pittman, R.N. (2006). Ataxin-3 binds VCP/p97 and regulates retrotranslocation of ERAD substrates. Human Mol. Genet. 15, 2409–2420. 4. Meusser, B., Hirsch, C., Jarosch, E., and Sommer, T. (2005). ERAD: the long road to destruction. Nat. Cell Biol. 7, 766–772. 5. Bays, N.W., Wilhovsky, S.K., Goradia, A., Hodgkiss-Harlow, K., and Hampton, R.Y. (2001). HRD4/NPL4 is required for the proteasomal processing of ubiquitinated ER proteins. Mol. Biol. Cell 12, 4114–4128. 6. Jarosch, E., Taxis, C., Volkwein, C., Bordallo, J., Finley, D., Wolf, D.H., and Sommer, T. (2002). Protein dislocation from the ER requires polyubiquitination and the AAA ATPase Cdc48. Nat. Cell Biol. 4, 134–139. 7. Rabinovich, E., Kerem, A., Frohlich, K.U., Diamant, N., and Bar-Nun, S. (2002). AAA ATPase p97/Cdc48, a cytosolic chaperone required for endoplasmic reticulum-associated protein degradation. Mol. Cell. Biol. 22, 626–634. 8. Ye, Y., Meyer, H.H., and Rapoport, T.A. (2001). The AAA ATPase Cdc48/p97 and its partners transport proteins from the ER to the cytosol. Nature 414, 652–656. 9. Medicherla, B., Kostova, Z., Schaefer, A., and Wolf, D.H. (2004). A genomic screen identifies Dsk2p and Rad23p as essential components of ER-associated degradation. EMBO Rep. 5, 692–697. 10. Doss-Pepe, E.W., Stenroos, E.S., Johnson, W.G., and Madura, K. (2003). Ataxin-3 interactions with Rad23 and valosin-containing protein and its associations with ubiquitin chains and the proteasome are consistent with a role in ubiquitin-mediated proteolysis. Mol. Cell. Biol. 23, 6469–6483. 11. Wang, G., Sawai, N., Kotliarova, S., Kanazawa, I., and Nukina, N. (2000). Ataxin-3, the MJD1 gene product, interacts with the two human homologues of yeast DNA repair protein RAD23, HHR23A and HHR23B. Human Mol. Genet. 9, 1795–1803.
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12. Chow, M.K., Mmackay, J.P., Whisstock, J.C., Scanlon, M.J., and Bottomley, S.P. (2004). Structural and functional analysis of the Josephin domain of the polyglutamine protein ataxin-3. Biochem. Biophys. Res. Commun 322, 387–394. 13. Berke, S.J., Chai, Y., Marrs, G.L., Wen, H., and Paulson, H.L. (2005). Defining the role of ubiquitin-interacting motifs in the polyglutamine diseases protein, ataxin-3. J. Biol. Chem. 280, 32026–32034. 14. Mao, Y., Senic-Matuglia, F., Di Flore, P.P., Polo, S., Hodsdon, M.E., and De
Camilli, P. (2005). Deubiquitinating function of ataxin-3: insights from the solution structure of the Josephin domain. Proc. Natl. Acad. Sci. USA 102, 12700–12705. 15. Burnett, B., Li, F., and Pittman, R.N. (2003). The polyglutamine neurodegenerative protein ataxin-3 binds polyubiquitinated proteins and has ubiquitin protease activity. Human Mol. Genet. 12, 3195–3205. 16. Richly, H., Rape, M., Braun, S., Rumpf, S., Hoege, C., and Jensch, S. (2005). A series of ubiquitin binding factors connects CDC48/p97 to substrate
Growth Regulation: A Beginning for the Hippo Pathway A signaling pathway involving two protein kinases, Hippo and Warts, restricts the growth of imaginal discs in Drosophila. Four recent studies taken together show that the protocadherin Fat can regulate Warts in two different ways. Iswar K. Hariharan Genetic studies in Drosophila have led to the identification of a signaling pathway, sometimes referred to as ‘the Hippo pathway’, that appears to be important in regulating tissue growth (reviewed in [1,2]). Inactivating mutations in several components of this pathway result in a dramatic overgrowth of mutant tissue in the imaginal discs — sacs of epithelial tissue found in the larva that eventually become adult structures such as the eye, wing and leg. Each component of the Hippo pathway identified to date has one or more mammalian orthologs that probably function in an analogous manner to their Drosophila counterparts. Thus, this pathway is likely to have a role in diverse species in the determination of the overall size of individual organs. Evidence is also accumulating to show that altered signaling via this pathway can contribute to the development of cancer in mammals [2–4]. At the core of this pathway (reviewed in [1,2]), is a module composed of two protein kinases, Hippo and Warts. Hippo, a member of the Ste20 superfamily, acts upstream of Warts, a member of the nuclear Dbf2 related (NDR) family of protein kinases, and activates it by direct
phosphorylation. Hippo-mediated Warts activation is facilitated by Salvador, a WW domain-containing protein that probably functions as a scaffold, and also by Mats, a protein that binds directly to Warts. When activated, Warts can phosphorylate a transcriptional co-activator, Yorkie [5] and reduce its activity, possibly by excluding it from the nucleus. Yorkie promotes, either directly or indirectly, the transcription of genes that promote growth and cell-cycle progression as well as genes that inhibit apoptosis. Thus, increased activity of Hippo and Warts correlates with a reduction in tissue growth and a reduction of Hippo/Warts activity allows growth to occur. Given the obvious importance of the Hippo pathway in regulating tissue growth, a gaping hole in our understanding of its role in organismal development has been the inability to correlate the level of activation of this pathway with any known physiological parameter. In contrast, the activity of many other growth regulators can be stimulated by extracellular growth factors (e.g. phosphatidylinositol 3-kinase) or the availability of nutrients (e.g. Tor). A possible role for the Hippo pathway was suggested by the finding that two proteins with four point one, ezrin, radixin, moesin (FERM) domains, Merlin and
multiubiquitination and proteasomal targeting. Cell 120, 73–84. 1
Faculty of Life Sciences, University of Manchester, Manchester M13 9PT, UK. 2 Department of Biological Sciences, University of Warwick, Coventry CV4 7AL, UK. E-mail: [email protected]
DOI: 10.1016/j.cub.2006.11.013
Expanded, may function redundantly as activators of Hippo [6]. Proteins with FERM domains are thought to link the cortical cytoskeleton with integral membrane proteins [7], leading to speculation that they, and hence the Hippo pathway, may function to inhibit tissue growth in response to increased cell density or possibly even mechanical stresses. Four recent studies [8–11], three published in Current Biology and one in Nature Genetics all implicate the protocadherin Fat [12,13] as an activator of the Hippo pathway. Linking Fat to the Hippo pathway represents an important development, since it is the first evidence that the activity of the Hippo pathway can potentially be regulated by an extracellular signal that is most likely a ligand expressed on the surface of an adjacent cell. While it has been known for many years that inactivating mutations in fat result in increased tissue growth, the reason for the increase in growth was not known. By analyzing the expression of transcriptional targets of the Hippo pathway, the authors of all four studies conclude that cells that are mutant for fat resemble, in several ways, cells that have inactivating mutations in hippo or warts. Importantly, however, the studies differ in their explanations for how Fat can regulate signaling via the Hippo pathway and propose two distinct models of Fat function. As will be discussed below, it is likely that Fat can regulate the Hippo pathway in both of these ways, but their relative importance under physiological conditions has yet to be determined.
Arabidopsis and rice genomes [18] suggests that lineage-specific tinkering with the composition of the plastid proteome can occur even over short evolutionary timescales. In the mean-time, multiple algal genome sequences are now publicly available — for example, those of the green algae Chlamydomonas and Ostreococcus (http://genome. jgi-psf.org/euk_cur1.html) and the red alga Cyanidioschyzon (http:// merolae.biol.s.u-tokyo.ac.jp) — and it is likely that ten or more plant and algal nuclear genome sequences will be available for even more thorough and systematic analyses within the next few years. An increasing number of genomes from ‘secondary’ plastid-containing algae such as diatoms, cryptophytes and chlorarachniophytes will also be completely sequenced (http:// www.jgi.doe.gov/). These organisms acquired their plastids through the engulfment of red or green algal endosymbionts [19] and the molecular dynamics accompanying the process of secondary endosymbiosis, in which gene transfers between evolutionarily distinct nuclear genomes are also a possibility, adds another layer of complexity to an already complicated picture [20]. Overall, it is sobering to consider how little we know about the nuts and bolts of endosymbiosis and the full scope of its role in the diversification of eukaryotic cells. References 1. Yoon, H.S., Hackett, J.D., Ciniglia, C., Pinto, G., and Bhattacharya, D. (2004). A molecular timeline for the origin of photosynthetic eukaryotes. Mol. Biol. Evol. 21, 809–818. 2. Reyes-Prieto, A., Hackett, J.D., Soares, M.B., Bonaldo, M.F., and Bhattacharya, D. (2006). Cyanobacterial contribution to algal nuclear genomes is primarily limited to plastid functions. Curr. Biol. 16, 2320–2325. 3. Martin, W., Rujan, T., Richly, E., Hansen, A., Cornelsen, S., Lins, T., Leister, D., Stoebe, B., Hasegawa, M., and Penny, D. (2002). Evolutionary analysis of Arabidopsis, cyanobacterial, and chloroplast genomes reveals plastid phylogeny and thousands of cyanobacterial genes in the nucleus. Proc. Natl. Acad. Sci. USA 99, 12246–12251. 4. Weeden, N.F. (1981). Genetic and biochemical implications of the endosymbiotic origin of the chloroplast. J. Mol. Evol. 17, 133–139.
5. Timmis, J.N., Ayliffe, M.A., Huang, C.Y., and Martin, W. (2004). Endosymbiotic gene transfer: organelle genomes forge eukaryotic chromosomes. Nat. Rev. Genet. 5, 123–135. 6. Bruce, B.D. (2000). Chloroplast transit peptides: structure, function and evolution. Trends Cell Biol. 10, 440–447. 7. McFadden, G.I. (1999). Plastids and protein targeting. J. Eukaryot. Microbiol. 46, 339–346. 8. Soll, J., and Schleiff, E. (2004). Protein import into chloroplasts. Nat. Rev. Mol. Cell Biol. 5, 198–208. 9. Martin, W., and Schnarrenberger, C. (1997). The evolution of the Calvin cycle from prokaryotic to eukaryotic chromosomes: a case study of functional redundancy in ancient pathways through endosymbiosis. Curr. Genet. 32, 1–18. 10. The Arabidopsis Genome Initiative (2000). Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature 408, 796–815. 11. Karol, K.G., McCourt, R.M., Cimino, M.T., and Delwiche, C.F. (2001). The closest living relatives of land plants. Science 294, 2351–2353. 12. Rodriguez-Ezpeleta, N., Brinkmann, H., Burey, S.C., Roure, B., Burger, G., Lo¨ffelhardt, W., Bohnert, H.J., Philippe, H., and Lang, B.F. (2005). Monophyly of primary photosynthetic eukaryotes: green plants, red algae, and glaucophytes. Curr. Biol. 15, 1325–1330. 13. Hall, W.T., and Claus, G. (1963). Ultrastructural studies on the blue-green algal symbiont in Cyanophora Paradoxa Korschikoff. J. Cell Biol. 19, 551–563. 14. Steiner, J.M., Bergho¨fer, J., Yusa, F., Pompe, J.A., Klo¨sgen, R.B., and Lo¨ffelhardt, W. (2005). Conservative sorting in a primitive plastid. The cyanelle
15.
16.
17.
18.
19.
20.
of Cyanophora paradoxa. FEBS J. 272, 987–998. Lo¨ffelhardt, W., Bohnert, H.J., and Bryant, D.A. (1997). The complete sequence of the Cyanophora paradoxa cyanelle genome. In Origins of Algae and Their Plastids, D. Bhattacharya, ed. (Wien: Springer), pp. 149–162. Martin, W., Stoebe, B., Goremykin, V., Hansmann, S., Hasegawa, M., and Kowallik, K.V. (1998). Gene transfer to the nucleus and the evolution of chloroplasts. Nature 393, 162–165. De Bodt, S., Maere, S., and Van de Peer, Y. (2005). Genome duplication and the origin of angiosperms. Trends Ecol. Evol. 20, 591–597. Richly, E., and Leister, D. (2004). An improved prediction of chloroplast proteins reveals diversities and commonalities in the chloroplast proteomes of Arabidopsis and rice. Gene 329, 11–16. Bhattacharya, D., Yoon, H.S., and Hackett, J.D. (2003). Photosynthetic eukaryotes unite: endosymbiosis connects the dots. Bioessays 26, 50–60. Archibald, J.M. (2005). Jumping genes and shrinking genomes—probing the evolution of eukaryotic photosynthesis with genomics. IUBMB Life 57, 539–547.
The Canadian Institute for Advanced Research, Program in Evolutionary Biology, Department of Biochemistry and Molecular Biology, Dalhousie University, Sir Charles Tupper Medical Building, 5850 College Street, Halifax, Nova Scotia, B3H 1X5, Canada. E-mail: [email protected] DOI: 10.1016/j.cub.2006.11.008
Quality Control: Linking Retrotranslocation and Degradation Misfolded proteins in the ER require the p97 AAA ATPase for dislocation across the membrane prior to degradation by the cytosolic proteasome. The mechanism by which dislocated proteins are delivered to the proteasome from p97 is unclear, but recent studies suggest an important regulatory role for the protein ataxin-3. Colin J. Stirling1 and J. Michael Lord2 The fatal neurodegenerative Machado-Joseph disease, also known as spinocerebellar ataxia type 3, is caused by mutations of the polyglutamine-containing protein ataxin-3 [1]. The biochemical properties of ataxin-3 are well known, but its physiological role has been elusive. Two recent studies [2,3] now implicate ataxin-3 in the process known as endoplasmicreticulum-associated degradation
(ERAD), suggesting a link between endoplasmic reticulum (ER) stress and the neuropathology associated with disease. After targeting to the ER, proteins are screened by a quality control system to prevent misfolded forms from progressing through the secretory pathway. Rather than accumulating within the cell, these aberrant proteins are disposed of by ERAD. This requires the dislocation of such proteins across the ER membrane and their subsequent degradation
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by the cytosolic proteasome system. The precise nature of the membrane channel through which dislocation occurs is not yet known, but a number of proteins have been implicated, including Sec61, Derlin-1 and VIMP [4]. However, evidence suggests that the hexameric AAA ATPase p97/VCP (known as Cdc48 in yeast) plays an important role in driving the dislocation of a range of proteins [5–8]. A complex comprising p97, and the adaptor proteins Ufd1 and Npl4, is recruited to the cytosolic face of the ER membrane, where it is believed to ‘pull’ dislocating polypeptides through the channel and into the cytosol. During their dislocation, ERAD substrates are normally polyubiquitinated, and the p97 complex has been shown to bind directly to these polyubiquitin chains. A further set of polyubiquitin-binding proteins, including Rad23p and Dsk2p, are associated with the proteasome, where they function in the final delivery of the ERAD substrate to the degradation machinery [9]. The mechanism by which an ERAD substrate is transferred from the p97 complex to proteasomal polyubiquitin-binding proteins is not known. Previous studies have shown that ataxin-3 can interact with both p97/VCP and Rad23p [10,11], and two recent papers [2,3] suggest that ataxin-3 is a critical link in this chain. Wang et al. [2] and Zhong and Pittman [3] have studied the effects of the expression of wild-type and mutant forms of ataxin-3 on a range of ERAD substrates. Both studies confirm that full-length ataxin-3 binds directly to p97, but ataxin-3 was also found in a membrane-associated form that could be co-immunoprecipitated together with several other ERAD components, including Derlin-1, VIMP and the ER-specific ubiquitin ligase Hrd1p [2]. Previous studies have shown that ataxin-3 has three functional ubiquitin-interacting modules plus a deubiquitinating activity [12–15]. These new studies [2,3] provide evidence that ataxin-3, and specifically its deubiquitinating activity, are required in ERAD. The
ataxin-3 C14A mutant lacks the catalytic deubiquitinating activity but is still able to bind ubiquitinated substrates with high affinity. Ectopic expression of ataxin-3 C14A in transfected cells led to the accumulation of typical ERAD substrates, including TCRa, a membrane component of the T-cell receptor which is degraded in the absence of other T-cell receptor subunits. The accumulated forms of TCRa were found to be extensively polyubiquitinated in atx C14A cells, suggesting a role for ataxin-3 in the deubiquitination of substrate during the normal ERAD process. This was further confirmed by the demonstration that p97-bound substrate could be deubiquitinated in vitro in the presence of wild-type ataxin-3 but not the C14A mutant. Consistent with this, Wang et al. [2] further demonstrated a dramatic increase in the amount of ubiquitinated proteins that could be co-immunoprecipitated with p97in ataxin-3 C14A cells compared to controls. Zhong and Pittman [3] further examined the effect of pathological mutations in the polyglutamine tract of ataxin-3. Their studies show that mutant ataxin-3 binds p97/VCP more avidly than the wild-type protein, and this correlates with decreased degradation of the ERAD substrate CD3d [3]. These studies suggest that the pathology of ataxin-3 mutants relates to defective ERAD. Taken together, the new data [2,3] indicate that ataxin-3 plays a critical role in the p97-associated degradation of ERAD substrates, and that this is necessary for the efficient release of substrate from the p97 dislocation machinery. Complete deubiquitination of substrate would prevent subsequent interaction with proteasomal polyubiquitin-binding proteins and so it is proposed that ataxin-3 might function as an ‘editing enzyme’ that merely shortens polyubiquitin chains. In this model the normal deubiquitinating activity of ataxin-3 would trim polyubiquitin chains to a level that allows their release from the p97 complex, but still permits binding to crucial proteasomal components.
Interestingly, the ubiquitin chains of ERAD substrates in Saccharomyces cerevisiae are progressively shortened as they transit to the proteasome [16]. Such a role for mammalian ataxin-3 would require tight regulation since any change in the level of deubiquitinating activity might be expected to perturb ubiquitin trimming and thus prevent efficient transfer of substrates to the proteasome. Indeed, ectopic over-expression of even wild-type ataxin-3 resulted in the accumulation of TCRa and CD3d [2,3], suggesting that this molecule is an important regulator of mammalian ERAD. References 1. Kawaguchi, Y., Okamoto, T., Taniwaki, M., Aizawa, M., Inoue, M., Katayama, S., Kawakami, H., Nakamura, S., Akiguchi, L., et al. (1994). CAG expansions in a novel gene for the Machado-Joseph disease at chromosome 14q32.1. Nat. Genet. 8, 221–229. 2. Wang, Q., Li, L., and Ye, Y. (2006). Regulation of retrotranslocation by p97-associated deubiquitinating enzyme ataxin-3. J. Cell Biol. 174, 963–971. 3. Zhong, X., and Pittman, R.N. (2006). Ataxin-3 binds VCP/p97 and regulates retrotranslocation of ERAD substrates. Human Mol. Genet. 15, 2409–2420. 4. Meusser, B., Hirsch, C., Jarosch, E., and Sommer, T. (2005). ERAD: the long road to destruction. Nat. Cell Biol. 7, 766–772. 5. Bays, N.W., Wilhovsky, S.K., Goradia, A., Hodgkiss-Harlow, K., and Hampton, R.Y. (2001). HRD4/NPL4 is required for the proteasomal processing of ubiquitinated ER proteins. Mol. Biol. Cell 12, 4114–4128. 6. Jarosch, E., Taxis, C., Volkwein, C., Bordallo, J., Finley, D., Wolf, D.H., and Sommer, T. (2002). Protein dislocation from the ER requires polyubiquitination and the AAA ATPase Cdc48. Nat. Cell Biol. 4, 134–139. 7. Rabinovich, E., Kerem, A., Frohlich, K.U., Diamant, N., and Bar-Nun, S. (2002). AAA ATPase p97/Cdc48, a cytosolic chaperone required for endoplasmic reticulum-associated protein degradation. Mol. Cell. Biol. 22, 626–634. 8. Ye, Y., Meyer, H.H., and Rapoport, T.A. (2001). The AAA ATPase Cdc48/p97 and its partners transport proteins from the ER to the cytosol. Nature 414, 652–656. 9. Medicherla, B., Kostova, Z., Schaefer, A., and Wolf, D.H. (2004). A genomic screen identifies Dsk2p and Rad23p as essential components of ER-associated degradation. EMBO Rep. 5, 692–697. 10. Doss-Pepe, E.W., Stenroos, E.S., Johnson, W.G., and Madura, K. (2003). Ataxin-3 interactions with Rad23 and valosin-containing protein and its associations with ubiquitin chains and the proteasome are consistent with a role in ubiquitin-mediated proteolysis. Mol. Cell. Biol. 23, 6469–6483. 11. Wang, G., Sawai, N., Kotliarova, S., Kanazawa, I., and Nukina, N. (2000). Ataxin-3, the MJD1 gene product, interacts with the two human homologues of yeast DNA repair protein RAD23, HHR23A and HHR23B. Human Mol. Genet. 9, 1795–1803.
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12. Chow, M.K., Mmackay, J.P., Whisstock, J.C., Scanlon, M.J., and Bottomley, S.P. (2004). Structural and functional analysis of the Josephin domain of the polyglutamine protein ataxin-3. Biochem. Biophys. Res. Commun 322, 387–394. 13. Berke, S.J., Chai, Y., Marrs, G.L., Wen, H., and Paulson, H.L. (2005). Defining the role of ubiquitin-interacting motifs in the polyglutamine diseases protein, ataxin-3. J. Biol. Chem. 280, 32026–32034. 14. Mao, Y., Senic-Matuglia, F., Di Flore, P.P., Polo, S., Hodsdon, M.E., and De
Camilli, P. (2005). Deubiquitinating function of ataxin-3: insights from the solution structure of the Josephin domain. Proc. Natl. Acad. Sci. USA 102, 12700–12705. 15. Burnett, B., Li, F., and Pittman, R.N. (2003). The polyglutamine neurodegenerative protein ataxin-3 binds polyubiquitinated proteins and has ubiquitin protease activity. Human Mol. Genet. 12, 3195–3205. 16. Richly, H., Rape, M., Braun, S., Rumpf, S., Hoege, C., and Jensch, S. (2005). A series of ubiquitin binding factors connects CDC48/p97 to substrate
Growth Regulation: A Beginning for the Hippo Pathway A signaling pathway involving two protein kinases, Hippo and Warts, restricts the growth of imaginal discs in Drosophila. Four recent studies taken together show that the protocadherin Fat can regulate Warts in two different ways. Iswar K. Hariharan Genetic studies in Drosophila have led to the identification of a signaling pathway, sometimes referred to as ‘the Hippo pathway’, that appears to be important in regulating tissue growth (reviewed in [1,2]). Inactivating mutations in several components of this pathway result in a dramatic overgrowth of mutant tissue in the imaginal discs — sacs of epithelial tissue found in the larva that eventually become adult structures such as the eye, wing and leg. Each component of the Hippo pathway identified to date has one or more mammalian orthologs that probably function in an analogous manner to their Drosophila counterparts. Thus, this pathway is likely to have a role in diverse species in the determination of the overall size of individual organs. Evidence is also accumulating to show that altered signaling via this pathway can contribute to the development of cancer in mammals [2–4]. At the core of this pathway (reviewed in [1,2]), is a module composed of two protein kinases, Hippo and Warts. Hippo, a member of the Ste20 superfamily, acts upstream of Warts, a member of the nuclear Dbf2 related (NDR) family of protein kinases, and activates it by direct
phosphorylation. Hippo-mediated Warts activation is facilitated by Salvador, a WW domain-containing protein that probably functions as a scaffold, and also by Mats, a protein that binds directly to Warts. When activated, Warts can phosphorylate a transcriptional co-activator, Yorkie [5] and reduce its activity, possibly by excluding it from the nucleus. Yorkie promotes, either directly or indirectly, the transcription of genes that promote growth and cell-cycle progression as well as genes that inhibit apoptosis. Thus, increased activity of Hippo and Warts correlates with a reduction in tissue growth and a reduction of Hippo/Warts activity allows growth to occur. Given the obvious importance of the Hippo pathway in regulating tissue growth, a gaping hole in our understanding of its role in organismal development has been the inability to correlate the level of activation of this pathway with any known physiological parameter. In contrast, the activity of many other growth regulators can be stimulated by extracellular growth factors (e.g. phosphatidylinositol 3-kinase) or the availability of nutrients (e.g. Tor). A possible role for the Hippo pathway was suggested by the finding that two proteins with four point one, ezrin, radixin, moesin (FERM) domains, Merlin and
multiubiquitination and proteasomal targeting. Cell 120, 73–84. 1
Faculty of Life Sciences, University of Manchester, Manchester M13 9PT, UK. 2 Department of Biological Sciences, University of Warwick, Coventry CV4 7AL, UK. E-mail: [email protected]
DOI: 10.1016/j.cub.2006.11.013
Expanded, may function redundantly as activators of Hippo [6]. Proteins with FERM domains are thought to link the cortical cytoskeleton with integral membrane proteins [7], leading to speculation that they, and hence the Hippo pathway, may function to inhibit tissue growth in response to increased cell density or possibly even mechanical stresses. Four recent studies [8–11], three published in Current Biology and one in Nature Genetics all implicate the protocadherin Fat [12,13] as an activator of the Hippo pathway. Linking Fat to the Hippo pathway represents an important development, since it is the first evidence that the activity of the Hippo pathway can potentially be regulated by an extracellular signal that is most likely a ligand expressed on the surface of an adjacent cell. While it has been known for many years that inactivating mutations in fat result in increased tissue growth, the reason for the increase in growth was not known. By analyzing the expression of transcriptional targets of the Hippo pathway, the authors of all four studies conclude that cells that are mutant for fat resemble, in several ways, cells that have inactivating mutations in hippo or warts. Importantly, however, the studies differ in their explanations for how Fat can regulate signaling via the Hippo pathway and propose two distinct models of Fat function. As will be discussed below, it is likely that Fat can regulate the Hippo pathway in both of these ways, but their relative importance under physiological conditions has yet to be determined.