- Email: [email protected]
Coordinating the Party: Assembly Factors and Ribogenesis
Molecular Cell
Previews Coordinating the Party: Assembly Factors and Ribogenesis Anna Marie Pyle1,2,3,* 1Howard
Hughes Medical Institute of Molecular, Cellular, and Developmental Biology 3Department of Chemistry Yale University, New Haven, CT 06520, USA *Correspondence: [email protected] http://dx.doi.org/10.1016/j.molcel.2013.11.007 2Department
Using hydroxyl-radical footprinting to map the structures of RNA molecules in whole cells, Soper et al. (2013) determine the specific role of assembly factors during the final stages of ribosomal subunit assembly and visualize structural features of intermediate states. Imagine you are hosting a huge party and nobody is assigned the task of ensuring that everything happens at the correct time. The food is delivered before the tables and tents, people arrive two hours early, and then a passing rainstorm makes a mess out of everything: very distressing. Ribosome assembly is a lot like an event with hundreds of guests, exotic food, and a full orchestra. Once the party starts upon transcription initiation, everything must go according to plan. In this issue, Soper and colleagues (2013) have found that ribosome assembly involves the participation of well-trained event planners who ensure that each stage of the party occurs properly, in the correct sequence, and with the appropriate timing and duration. By removing these planners (assembly factors) from the process and then monitoring the mishaps that subsequently occur, Soper and colleagues (2013) have pinpointed the specific roles of protein factors RimM and RbfA during assembly of the 30S ribosomal subunit, and they have structurally characterized a latestage assembly intermediate that is controlled by these factors. This analysis was made possible by two key tools. The first is hydroxyl-radical mapping of RNA structure in whole cells, using synchrotron radiation (Adilakshmi et al., 2009). By growing wild-type and mutant strains of bacteria under specific conditions and then bombarding them with hydroxyl radicals, the authors have obtained detailed comparative structural information about cellular RNAs at single nucleotide resolution. And unlike in-vitro RNA folding experiments (where the
whole RNA molecule has already been transcribed), this experiment enables Soper and colleagues to monitor RNAs that have folded under natural conditions, in the context of active transcription. The second key tool is the availability of mutant bacterial strains in which key assembly factors have been rendered inactive (DrimM and DrbfA) (Guo et al., 2013). It had already been established through genetic and biochemical studies that RimM (Guo et al., 2013) and RbfA (Datta et al., 2007) were important during late stages of 30S assembly. The DrimM and DrbfA strains were therefore ideal for examining the structural and mechanistic consequences of poor coordination during late stages of assembly. Using these powerful tools, Soper and colleagues make some intriguing observations. In the DrimM and DrbfA strains, 30S assembly stalls at a very late stage, resulting in abundant pre-30S particles (also referred to as 45 S particles) that have specific compositions and structures (Soper et al., 2013; Guo et al., 2013). Hydroxyl-radical footprinting of DrimM and DrbfA cells reveals that downstream portions of the ribosomal RNA are not properly folded: for example, the long H44 helix at the 30 -RNA terminus is undocked from the core and long-range tertiary interactions are absent (Soper et al., 2013). In addition, mass spectrometry on the stalled complexes was used to determine whether the protein composition had also changed (Soper et al., 2013). Indeed, pre-30S particles from the DrbfA strain lack ribosomal proteins S2, S3, and S21, which are known to associate during specific stages of
assembly (Talkington et al., 2005; Held et al., 1974. Perhaps most striking, Soper and colleagues show that posttranslational modifications on the bound ribosomal proteins are also different in the mutant strains. Specifically, the S18 and S5 proteins are acetylated in 30S particles from the wild-type strain, but these same proteins are only partially acetylated or lack acetylation altogether in the mutant strains where their respective binding sites are not formed correctly. For example, S18 is not acetylated in the DrimM strain where its binding site is perturbed, but it is acetylated in the DrbfA strain, where its binding site is formed properly. This intriguing finding is reminiscent of the role played by histone acetylation in nucleosomes, suggesting that just as in chromatin, acetylation can serve as a ‘‘go,’’ or ‘‘no go’’ signal for regulated function during ribosome assembly and perhaps in other ribonucleoprotein complexes. Given these findings, what are the actual roles of RimM and RbfA, and what does this mean for the mechanism of late 30S assembly? The requirement of factors to shepherd passage through the pre-30S intermediate could mean at least two things: either these proteins participate directly by sequentially recruiting ribosomal proteins, or they may act as traffic cops, ensuring that critical slow steps occur before high-affinity participants jump in and complete the last stage of assembly. Soper and colleagues think that the latter is most consistent with available data, citing a number of clues. For example, they do not think that RimM serves to recruit ribosomal
Molecular Cell 52, November 21, 2013 ª2013 Elsevier Inc. 469
Molecular Cell
Previews protein S19 to the complex, facilitating sequential stages of assembly through ordered addition of ribosomal proteins. Rather, they propose that RimM interacts with S19 and RNA helix 32 in a manner that holds the 30 -domain of 16S rRNA open, thereby delaying RNA folding at the downstream terminus and enabling helices 31 and 43 to form correctly. Consistent with previous studies (Dammel and Noller, 1995), RbfA facilitates the ordered refolding of 16 s RNA motifs. But, intriguingly, neither RimM nor RbfA are DEAD-box proteins or NTPases, which are the typical suspects in RNAremodeling events, although RbfA may act in a complex with a GTPase. Thus, RimM might facilitate folding through simple binding interactions, much like protein CBP2 bound to the near-native state of bI5 group I intron (Garcia and Weeks, 2004). It is also notable that late-stage assembly also involves the remodeling of protein structures, since the conformation of S5 appears to change, and protein acetylation makes a clear contribution to the assembly
pathway (Soper et al., 2013). Thus, the common notion that 16S rRNA and other large RNA molecules always fold incorrectly without oversight by mechanical protein chaperones (presumably because proteins are smarter than RNA) is outdated, or at least incomplete. Rather, a combination of coordinated RNA and protein structural reorganization and modification events provide checkpoints that ensure proper assembly of active 30S subunits. This study (Soper et al., 2013), together with complementary studies using cryo-electron microscopy (Guo et al., 2013) and mass spectrometry (Chen and Williamson, 2013), and other work on assembly factors (Strunk et al., 2011), sheds new light on the diverse and complex processes that go on behind the scenes during the successful process of ribosomal subunit assembly.
Chen, S.S., and Williamson, J.R. (2013). Mol. Biol. 425, 767–779. Dammel, C.S., and Noller, H.F. (1995). Genes Dev. 9, 626–637. Datta, P.P., Wilson, D.N., Kawazoe, M., Swami, N.K., Kaminishi, T., Sharma, M.R., Booth, T.M., Takemoto, C., Fucini, P., Yokoyama, S., and Agrawal, R.K. (2007). Mol. Cell 28, 434–445. Garcia, I., and Weeks, K.M. (2004). Biochemistry 43, 15179–15186. Guo, Q., Goto, S., Chen, Y., Feng, B., Xu, Y., Muto, A., Himeno, H., Deng, H., Lei, J., and Gao, N. (2013). Nucleic Acids Res. 41, 2609–2620. Held, W.A., Ballou, B., Mizushima, S., and Nomura, M. (1974). J. Biol. Chem. 249, 3103–3111. Soper, S.F.C., Dator, R.P., Limbach, P.A., and Woodson, S.A. (2013). Mol. Cell 52, this issue, 506–516.
REFERENCES
Strunk, B.S., Loucks, C.R., Su, M., Vashisth, H., Cheng, S., Schilling, J., Brooks, C.L., 3rd, Karbstein, K., and Skiniotis, G. (2011). Science 333, 1449–1453.
Adilakshmi, T., Soper, S.F., and Woodson, S.A. (2009). Methods Enzymol. 468, 239–258.
Talkington, M.W., Siuzdak, G., and Williamson, J.R. (2005). Nature 438, 628–632.
Histone 3 S10 Phosphorylation: ‘‘Caught in the R Loop!’’ Konstantina Skourti-Stathaki1 and Nicholas J. Proudfoot1,* 1Sir William Dunn School of Pathology, South Parks Road, University of Oxford, Oxford OX1 3RE, UK *Correspondence: [email protected] http://dx.doi.org/10.1016/j.molcel.2013.11.006
In this issue of Molecular Cell, Castellano-Pozo et al. (2013) describe a connection between R loop structures and histone 3 S10 phosphorylation (H3S10P), a mark of chromatin compaction. Their results constitute a significant advance in our understanding of the role of R loops in genomic instability. R loops are three stranded structures where an RNA transcript hybridizes with the DNA template, leaving the nontemplate DNA single stranded (ss). R loops form naturally as intermediates during bacterial and mitochondrial replication and immunoglobulin class-switching in activated B lymphocytes (see Aguilera and Garcı´a-Muse, 2012 for review). They also form during transcription, especially
in human G-rich termination pause elements and over mammalian CpG island promoters (Skourti-Stathaki et al., 2011; Ginno et al., 2013). Persistent formation of R loops, however, can be a risky outcome of the transcription process, with deleterious effects on genome integrity (Aguilera and Garcı´a-Muse, 2012). Even though the R loop field is an expanding area of research, the mecha-
470 Molecular Cell 52, November 21, 2013 ª2013 Elsevier Inc.
nism leading from an R loop to DNA damage still remains largely unknown. Castellano-Pozo et al. (2013) now suggest an additional player in this process, the chromatin structure. They initially observed, in S. cerevisiae, that the R-loopaccumulating strain hpr1D shows high levels of the histone 3 S10 phosphorylation (H3S10P) chromatin mark. This histone modification is cell cycle regulated
Previews Coordinating the Party: Assembly Factors and Ribogenesis Anna Marie Pyle1,2,3,* 1Howard
Hughes Medical Institute of Molecular, Cellular, and Developmental Biology 3Department of Chemistry Yale University, New Haven, CT 06520, USA *Correspondence: [email protected] http://dx.doi.org/10.1016/j.molcel.2013.11.007 2Department
Using hydroxyl-radical footprinting to map the structures of RNA molecules in whole cells, Soper et al. (2013) determine the specific role of assembly factors during the final stages of ribosomal subunit assembly and visualize structural features of intermediate states. Imagine you are hosting a huge party and nobody is assigned the task of ensuring that everything happens at the correct time. The food is delivered before the tables and tents, people arrive two hours early, and then a passing rainstorm makes a mess out of everything: very distressing. Ribosome assembly is a lot like an event with hundreds of guests, exotic food, and a full orchestra. Once the party starts upon transcription initiation, everything must go according to plan. In this issue, Soper and colleagues (2013) have found that ribosome assembly involves the participation of well-trained event planners who ensure that each stage of the party occurs properly, in the correct sequence, and with the appropriate timing and duration. By removing these planners (assembly factors) from the process and then monitoring the mishaps that subsequently occur, Soper and colleagues (2013) have pinpointed the specific roles of protein factors RimM and RbfA during assembly of the 30S ribosomal subunit, and they have structurally characterized a latestage assembly intermediate that is controlled by these factors. This analysis was made possible by two key tools. The first is hydroxyl-radical mapping of RNA structure in whole cells, using synchrotron radiation (Adilakshmi et al., 2009). By growing wild-type and mutant strains of bacteria under specific conditions and then bombarding them with hydroxyl radicals, the authors have obtained detailed comparative structural information about cellular RNAs at single nucleotide resolution. And unlike in-vitro RNA folding experiments (where the
whole RNA molecule has already been transcribed), this experiment enables Soper and colleagues to monitor RNAs that have folded under natural conditions, in the context of active transcription. The second key tool is the availability of mutant bacterial strains in which key assembly factors have been rendered inactive (DrimM and DrbfA) (Guo et al., 2013). It had already been established through genetic and biochemical studies that RimM (Guo et al., 2013) and RbfA (Datta et al., 2007) were important during late stages of 30S assembly. The DrimM and DrbfA strains were therefore ideal for examining the structural and mechanistic consequences of poor coordination during late stages of assembly. Using these powerful tools, Soper and colleagues make some intriguing observations. In the DrimM and DrbfA strains, 30S assembly stalls at a very late stage, resulting in abundant pre-30S particles (also referred to as 45 S particles) that have specific compositions and structures (Soper et al., 2013; Guo et al., 2013). Hydroxyl-radical footprinting of DrimM and DrbfA cells reveals that downstream portions of the ribosomal RNA are not properly folded: for example, the long H44 helix at the 30 -RNA terminus is undocked from the core and long-range tertiary interactions are absent (Soper et al., 2013). In addition, mass spectrometry on the stalled complexes was used to determine whether the protein composition had also changed (Soper et al., 2013). Indeed, pre-30S particles from the DrbfA strain lack ribosomal proteins S2, S3, and S21, which are known to associate during specific stages of
assembly (Talkington et al., 2005; Held et al., 1974. Perhaps most striking, Soper and colleagues show that posttranslational modifications on the bound ribosomal proteins are also different in the mutant strains. Specifically, the S18 and S5 proteins are acetylated in 30S particles from the wild-type strain, but these same proteins are only partially acetylated or lack acetylation altogether in the mutant strains where their respective binding sites are not formed correctly. For example, S18 is not acetylated in the DrimM strain where its binding site is perturbed, but it is acetylated in the DrbfA strain, where its binding site is formed properly. This intriguing finding is reminiscent of the role played by histone acetylation in nucleosomes, suggesting that just as in chromatin, acetylation can serve as a ‘‘go,’’ or ‘‘no go’’ signal for regulated function during ribosome assembly and perhaps in other ribonucleoprotein complexes. Given these findings, what are the actual roles of RimM and RbfA, and what does this mean for the mechanism of late 30S assembly? The requirement of factors to shepherd passage through the pre-30S intermediate could mean at least two things: either these proteins participate directly by sequentially recruiting ribosomal proteins, or they may act as traffic cops, ensuring that critical slow steps occur before high-affinity participants jump in and complete the last stage of assembly. Soper and colleagues think that the latter is most consistent with available data, citing a number of clues. For example, they do not think that RimM serves to recruit ribosomal
Molecular Cell 52, November 21, 2013 ª2013 Elsevier Inc. 469
Molecular Cell
Previews protein S19 to the complex, facilitating sequential stages of assembly through ordered addition of ribosomal proteins. Rather, they propose that RimM interacts with S19 and RNA helix 32 in a manner that holds the 30 -domain of 16S rRNA open, thereby delaying RNA folding at the downstream terminus and enabling helices 31 and 43 to form correctly. Consistent with previous studies (Dammel and Noller, 1995), RbfA facilitates the ordered refolding of 16 s RNA motifs. But, intriguingly, neither RimM nor RbfA are DEAD-box proteins or NTPases, which are the typical suspects in RNAremodeling events, although RbfA may act in a complex with a GTPase. Thus, RimM might facilitate folding through simple binding interactions, much like protein CBP2 bound to the near-native state of bI5 group I intron (Garcia and Weeks, 2004). It is also notable that late-stage assembly also involves the remodeling of protein structures, since the conformation of S5 appears to change, and protein acetylation makes a clear contribution to the assembly
pathway (Soper et al., 2013). Thus, the common notion that 16S rRNA and other large RNA molecules always fold incorrectly without oversight by mechanical protein chaperones (presumably because proteins are smarter than RNA) is outdated, or at least incomplete. Rather, a combination of coordinated RNA and protein structural reorganization and modification events provide checkpoints that ensure proper assembly of active 30S subunits. This study (Soper et al., 2013), together with complementary studies using cryo-electron microscopy (Guo et al., 2013) and mass spectrometry (Chen and Williamson, 2013), and other work on assembly factors (Strunk et al., 2011), sheds new light on the diverse and complex processes that go on behind the scenes during the successful process of ribosomal subunit assembly.
Chen, S.S., and Williamson, J.R. (2013). Mol. Biol. 425, 767–779. Dammel, C.S., and Noller, H.F. (1995). Genes Dev. 9, 626–637. Datta, P.P., Wilson, D.N., Kawazoe, M., Swami, N.K., Kaminishi, T., Sharma, M.R., Booth, T.M., Takemoto, C., Fucini, P., Yokoyama, S., and Agrawal, R.K. (2007). Mol. Cell 28, 434–445. Garcia, I., and Weeks, K.M. (2004). Biochemistry 43, 15179–15186. Guo, Q., Goto, S., Chen, Y., Feng, B., Xu, Y., Muto, A., Himeno, H., Deng, H., Lei, J., and Gao, N. (2013). Nucleic Acids Res. 41, 2609–2620. Held, W.A., Ballou, B., Mizushima, S., and Nomura, M. (1974). J. Biol. Chem. 249, 3103–3111. Soper, S.F.C., Dator, R.P., Limbach, P.A., and Woodson, S.A. (2013). Mol. Cell 52, this issue, 506–516.
REFERENCES
Strunk, B.S., Loucks, C.R., Su, M., Vashisth, H., Cheng, S., Schilling, J., Brooks, C.L., 3rd, Karbstein, K., and Skiniotis, G. (2011). Science 333, 1449–1453.
Adilakshmi, T., Soper, S.F., and Woodson, S.A. (2009). Methods Enzymol. 468, 239–258.
Talkington, M.W., Siuzdak, G., and Williamson, J.R. (2005). Nature 438, 628–632.
Histone 3 S10 Phosphorylation: ‘‘Caught in the R Loop!’’ Konstantina Skourti-Stathaki1 and Nicholas J. Proudfoot1,* 1Sir William Dunn School of Pathology, South Parks Road, University of Oxford, Oxford OX1 3RE, UK *Correspondence: [email protected] http://dx.doi.org/10.1016/j.molcel.2013.11.006
In this issue of Molecular Cell, Castellano-Pozo et al. (2013) describe a connection between R loop structures and histone 3 S10 phosphorylation (H3S10P), a mark of chromatin compaction. Their results constitute a significant advance in our understanding of the role of R loops in genomic instability. R loops are three stranded structures where an RNA transcript hybridizes with the DNA template, leaving the nontemplate DNA single stranded (ss). R loops form naturally as intermediates during bacterial and mitochondrial replication and immunoglobulin class-switching in activated B lymphocytes (see Aguilera and Garcı´a-Muse, 2012 for review). They also form during transcription, especially
in human G-rich termination pause elements and over mammalian CpG island promoters (Skourti-Stathaki et al., 2011; Ginno et al., 2013). Persistent formation of R loops, however, can be a risky outcome of the transcription process, with deleterious effects on genome integrity (Aguilera and Garcı´a-Muse, 2012). Even though the R loop field is an expanding area of research, the mecha-
470 Molecular Cell 52, November 21, 2013 ª2013 Elsevier Inc.
nism leading from an R loop to DNA damage still remains largely unknown. Castellano-Pozo et al. (2013) now suggest an additional player in this process, the chromatin structure. They initially observed, in S. cerevisiae, that the R-loopaccumulating strain hpr1D shows high levels of the histone 3 S10 phosphorylation (H3S10P) chromatin mark. This histone modification is cell cycle regulated