Ireton, K., Gunther, N.W., 4th, and Grossman, A.D. (1994). J. Bacteriol. 176, 5320–5329. Hirano, T. (2006). Nat. Rev. Mol. Cell Biol. 7, 311–322. Mascarenhas, J., Soppa, J., Strunnikov, A.V., and
Graumann, P.L. (2002). EMBO J. 21, 3108–3118. Murray, H., and Errington, J. (2008). Cell 135, 74–84. Sullivan, N.L., Marquis, K.A., and Rudner, D.Z. (2009). Cell, this issue.
Thanbichler, M., and Shapiro, L. (2006). J. Struct. Biol. 156, 292–303. Viollier, P.H., Thanbichler, M., McGrath, P.T., West, L., Meewan, M., McAdams, H.H., and Shapiro, L. (2004). Proc. Natl. Acad. Sci. USA 101, 9257–9262.
Shedding UV Light on Alternative Splicing Matthew S. Marengo1,2,* and Mariano A. Garcia-Blanco1,2,3,*
Department of Molecular Genetics and Microbiology Center for RNA Biology 3 Department of Medicine Duke University Medical Center, Durham, NC 27710, USA *Correspondence:
[email protected] (M.S.M),
[email protected] (M.A.G.-B.) DOI 10.1016/j.cell.2009.04.054 1 2
After DNA damage, cells modulate pre-messenger RNA (pre-mRNA) splicing to induce an anti- or proapoptotic response. In this issue, Muñoz et al. (2009) uncover a cotranscriptional mechanism for activating alternative pre-mRNA splicing after ultraviolet irradiation that depends unexpectedly on hyperphosphorylation of the RNA polymerase II C-terminal domain and decreased rates of transcription elongation. In response to a genotoxic insult, normal eukaryotic cells activate the DNA-damage response, which includes programs that mediate DNA repair and apoptosis. For decades, research on the DNA-damage response has focused on signaling kinases, the targets of transcription factors, and transcriptional regulation. More recently, however, it has been shown that alternative pre-messenger RNA (premRNA) splicing is also a target of the DNA-damage response (Katzenberger et al., 2006; Matsuoka et al., 2007). In this issue of Cell, Muñoz et al. (2009) describe a new mechanism for genotoxicity-induced alternative splicing that takes a shortcut around the DNA-damage response to target RNA polymerase II (RNAPII), the enzyme that synthesizes pre-mRNA. Muñoz et al. show that ultraviolet (UV) radiation changes the phosphorylation state of the carboxy-terminal repeat domain (CTD) of RNAPII (Figure 1). Using single-locus imaging by fluorescence recovery after photobleaching, the authors conclude that transcription elongation is slower in irradiated cultured
human cells. This change in the RNAPII elongation rate seems to affect the RNA available for cotranscriptional splicing (Goldstrohm et al., 2001). This results in alternative splicing of the BCL-X and CASPASE 9 pre-mRNAs, leading to a proapoptotic response. Alternative splicing of transcripts from these genes appears to occur independently of DNAdamage response signals that are known to play cotranscriptional or posttranscriptional regulatory roles. For example, BRCA1, originally identified as a protooncogene in breast cancer, encodes a protein that acts as a critical scaffold for DNA lesion detection and repair. At a DNA lesion, BRCA1 binds directly to the BRCA1-associated RING domain protein (BARD1), a ubiquitin ligase. BARD1 in turn can block RNAPII from interacting with the mRNA polyadenylation factor CstF and also marks RNAPII for degradation (Kim et al., 2006 and references therein) (Figure 1). This could affect steady-state mRNA levels of genes undergoing transcription. However, Muñoz et al. find that neither depleting BARD proteins nor changing the polyadenylation signals on
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RNA transcripts modulates a UV-induced splicing event. A broadly specific pharmacological block of the DNA-damage response signaling kinases, ATM and ATR, which are known to phosphorylate splicing factors (Matsuoka et al., 2007), also had no effect. Importantly, Muñoz et al. show that the effect of UV irradiation on alternative splicing is independent of p53, a central regulator of DNA-damage response-induced gene expression. Together, these results suggest the existence of a previously unidentified signaling pathway that modulates alternative splicing in response to DNA damage. The pleiotropic and incomplete effects of the pharmacological inhibition and the small-interfering RNA-mediated depletion of DNA-damage response proteins leave room for alternative explanations. Using a clever chemical genetics approach, however, the investigators were able to demonstrate a role for the RNAPII CTD in the modulation of alternative splicing. They poisoned the activity of endogenous RNAPII in cultured human cells with the toxin α-amanitin and introduced into the same cells α-amanitin-
resistant RNAPII that harbors mutations in the repeating CTD heptad protein sequence (YSPTSPS: Y, tyrosine; S, serine; P, proline; T, threonine). Expression of mutant RNAPII in which every serine at positions 2 and 5 in the repeating CTD heptad sequence is replaced by a nonphosphorylatable alanine prevents UVinduced alternative splicing. In contrast, expression of RNAPII mutant proteins in which these serines are replaced by glutamates (which resemble phosphorylated serines) recapitulates UV-induced alternative splicing. These observations indicate that CTD phosphorylation is an important event downstream of UVinduced signaling and that CTD modifications are sufficient to change splicing patterns in living cells. The authors go on to explore the cause and effect of UV-induced CTD phosphorylation. Alternative splicing of the BCL-X and CASPASE 9 transcripts could be perturbed by inhibition of a cyclin-dependent kinase (CDK). CDK dependence would fit well with the observations of Muñoz et al. as CDK9 is known to phosphorylate the CTD of RNAPII upon UV irradiation (Nguyen et al., 2001). The consequence of the observed CTD phosphorylation after UV irradiation is, however, new and unexpected. Previous studies indicated that phosphorylation of the CTD heptad repeat serines at positions 2 and 5 is a mark of an elongation-competent RNAPII. Yet, the glutamate phosphomimetic RNAPII mutant proteins exhibit a slower rate of transcription elongation, resulting in local mRNA accumulation similar to that observed for endogenous RNAPII after UV-induced phosphorylation. One possible explanation for this is that CTD phosphorylation is much more than a bulk change in electrostatic charge—rather, it is a context-sensitive code for the differential recruitment of binding partners (Phatnani and Greenleaf, 2006). Mutation or UV-induced phosphorylation of the CTD may disrupt the context necessary for the proper recruitment of transcription elongation factors, resulting in an unexpectedly slow polymerase. It should be noted, however, that the findings in the experiments using fluorescence recovery after photobleaching to determine transcription elongation rates are open to alternative interpretations. Indeed, these experiments show
Figure 1. DNA Damage Alters Patterns of mRNA Splice Variants Ultraviolet (UV) irradiation causes a DNA lesion that can affect the progress of RNA polymerase II (RNAPII) as it moves along the damaged DNA molecule. It may do this by causing the enzyme to stall during transcription elongation or by activating the BARD1 ubiquitin ligase through the DNA-damage response (DDR) to block messenger RNA (mRNA) polyadenylation and induce RNAPII degradation (Brueckner et al., 2007). Such cis effects of UV irradiation may exert selective pressure to keep UV radiation-responsive genes small in size, thus reducing the likelihood of a lesion within them (McKay et al., 2004). UV irradiation can also cause effects in trans. Spliceosome subunits in the cell may be phosphorylated by DNA-damage response kinases or dephosphorylated by the ceramide-regulated protein phosphatase 1 (PP1) resulting in regulation of alternative splicing. Additionally, as demonstrated by Muñoz et al. (2009), a response to UV radiation that is independent of the DNA-damage response can activate a cyclin-dependent kinase (CDK), which phosphorylates and, paradoxically, slows RNAPII thereby changing cotranscriptional alternative splicing.
intriguing differences between mutant RNAPII proteins harboring modified CTD sequences and wild-type RNAPII molecules after UV irradiation. Future experiments could explore the role of the lipid messenger ceramide in UV-induced alternative splicing, as UV treatment increases ceramide levels. This lipid can induce dephosphorylation of splicing factors by protein phosphatase 1 (PP1) to regulate the alternative splicing of BCL-X and CASPASE 9 pre-mRNAs (Chalfant et al., 2002). Thus, CTD phosphorylation, ceramide, DNA-damage response kinases, and BARD may regulate independent or overlapping pre-mRNA processing programs (Figure 1). The authors also use a splicing-sensitive microarray to identify more UV-induced alternative splicing events, many of which appear to be linked to changes in overall mRNA expression levels. Nonetheless, the direct contribution of CTD phosphorylation to
the regulation of global splicing patterns remains unclear. Examining genome-wide splicing changes upon the expression of the glutamate phosphomimetic RNAPII mutant proteins could provide insight into possible global splicing effects. It may be that CTD phosphorylation-induced splicing upregulates a proapoptotic program, whereas other DNA-damage response signals upregulate a pro-repair program. The results of the Muñoz et al. study may have important implications for cancer therapy because the observed splicing effects can be recapitulated by treating cells with the anticancer genotoxin cisplatin. These splicing effects are also independent of the function of p53, which is commonly lost in many human cancers. Thus, understanding how genotoxicity-induced splicing is regulated in the absence of p53 may be critical for designing effective and less toxic cancer therapies.
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Acknowledgments The authors thank C. Webster for improving the figure and A. Greenleaf, D. Wassarman, J. Pearson, C. Bennett, and T. Robinson for comments on the figure and manuscript. References Brueckner, F., Hennecke, U., Carell, T., and Cramer, P. (2007). Science 315, 859–862. Chalfant, C.E., Rathman, K., Pinkerman, R.L., Wood, R.E., Obeid, L.M., Ogretmen, B.,
and Hannun, Y.A. (2002). J. Biol. Chem. 277, 12587–12595. Goldstrohm, A.C., Greenleaf, A.L., and GarciaBlanco, M.A. (2001). Gene 277, 31–47. Katzenberger, R.J., Marengo, M.S., and Wassarman, D.A. (2006). Mol. Cell. Biol. 26, 9256–9267. Kim, H.S., Li, H., Cevher, M., Parmelee, A., Fonseca, D., Kleiman, F.E., and Lee, S.B. (2006). Cancer Res. 66, 4561–4565. Matsuoka, S., Ballif, B.A., Smogorzewska, A., McDonald, E.R., 3rd, Hurov, K.E., Luo, J., Bakalarski, C.E., Zhao, Z., Solimini, N., Lerenthal, Y., et al.
(2007). Science 316, 1160–1166. McKay, B.C., Stubbert, L.J., Fowler, C.C., Smith, J.M., Cardamore, R.A., and Spronck, J.C. (2004). Proc. Natl. Acad. Sci. USA 101, 6582–6586. Muñoz, M.J., Santangelo, M.S.P., Paronetto, M.P., de la Mata, M., Pelisch, F., Boireau, S., Glover-Cutter, K., Ben-Dov, C., Blaustein, M., Lozano, J.J., et al. (2009). Cell, this issue. Nguyen, V.T., Kiss, T., Michels, A.A., and Bensaude, O. (2001). Nature 414, 322–325. Phatnani, H.P., and Greenleaf, A.L. (2006). Genes Dev. 20, 2922–2936.
Keeping the Beat in the Rising Heat David M. Virshup1,* and Daniel B. Forger2
Program in Cancer and Stem Cell Biology, Duke-NUS Graduate Medical School, Singapore 169857 Department of Mathematics and Center for Computational Medicine and Biology, University of Michigan, Ann Arbor, MI 48109, USA *Correspondence:
[email protected] DOI 10.1016/j.cell.2009.04.051
1 2
Circadian clocks use temperature compensation to keep accurate time over a range of temperatures, thus allowing reliable timekeeping under diverse environmental conditions. Mehra et al. (2009) and Baker et al. (2009) now show that phosphorylation-regulated protein degradation plays a key role in circadian temperature compensation. Rising temperatures result in rising reaction rates for most chemical and biochemical processes. Fish in icy Michigan lakes have slower metabolic rates than their brethren in Southeast Asia, and hibernating animals slow their basal metabolic rates by lowering their core temperature. Although for some biological systems there is an adaptive advantage in having reaction rates that change with temperature, other processes must maintain the same reaction rate regardless of temperature (Figure 1A). In particular, the 24 hr circadian clock accurately keeps time regardless of temperature. This phenomenon, known as temperature compensation, has fascinated circadian biologists for years (Hastings and Sweeney, 1957; Pittendrigh, 1954). Two new studies from Jay Dunlap’s group, published in Cell (Mehra et al., 2009) and Molecular Cell (Baker et al., 2009), shed light on the molecular basis of temperature compensation.
Temperature compensation is especially important for poikilotherms (organisms whose internal temperature depends on the environment) but is also preserved in homeotherms like us and can be detected in hibernating mammals, laboratory animals, and cultured mammalian cells (for example see Izumo et al., 2003). The preservation of temperature compensation even in nonhibernating mammals suggests that it is an intrinsic and universally conserved feature of the circadian clock mechanism. The most likely basis for this mechanism, proposed long before the biochemical details of circadian timekeeping were known, is that a series of counterbalanced biochemical reactions in the core of the clock undergo equal and opposing changes with alterations in temperature (Hastings and Sweeney, 1957) (Figures 1B and 1C). However, despite more than 50 years of theories and experimentation, as well as enor-
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mous gains in the molecular understanding of the circadian clock, the mechanism of temperature compensation has remained a black box. The basic mechanism of all known circadian timekeepers involves the rhythmic accumulation of key transcriptional repressors, such as the PERIOD protein in the fly Drosophila melanogaster, the CRYPTOCHROME proteins in mammals, and the FREQUENCY (FRQ) protein in the fungus Neurospora crassa. The abundance of these proteins is regulated largely by a balance, which changes over the course of the day, between phosphorylation and dephosphorylation events that regulate protein stability. One might suspect, then, that temperature compensation could occur if the rising temperature simultaneously increased the opposing activities of kinases, phosphatases, and the protein degradation machinery. In their new study, Mehra et al. (2009) show using a combined genetic and biochemical approach that