YidC: Evaluating the Importance of the Native Environment

Structure

Previews absence of Pol 3 might assist Ctf18-RFC in clamp loading away from the fork. Beyond its activity as clamp loader, interesting questions also remain concerning Ctf18-RFC’s role during checkpoint activation: for instance, how does it cooperate with Pol 3 and the checkpoint mediator Mrc1 to recruit the Rad53 kinase? The goal of future mechanistic studies will be to combine our knowledge of the dual biochemical activities of Ctf18-RFC, clamp loading by its RFC subunits and polymerase binding by the Ctf18-1-8 module, into one integrated model able to account for Ctf18-RFC’s diverse roles in DNA replication, S-phase checkpoint, and sister chromatid cohesion. REFERENCES Clark, K.L., Halay, E.D., Lai, E., and Burley, S.K. (1993). Co-crystal structure of the HNF-3/fork

head DNA-recognition motif resembles histone H5. Nature 364, 412–420. Crabbe´, L., Thomas, A., Pantesco, V., De Vos, J., Pasero, P., and Lengronne, A. (2010). Analysis of replication profiles reveals key role of RFC-Ctf18 in yeast replication stress response. Nat. Struct. Mol. Biol. 17, 1391–1397. Garcı´a-Rodrı´guez, L.J., De Piccoli, G., Marchesi, V., Jones, R.C., Edmondson, R.D., and Labib, K. (2015). A conserved Pol3 binding module in Ctf18-RFC is required for S-phase checkpoint activation downstream of Mec1. Nucleic Acids Res. 43, 8830–8838. Grabarczyk, D.B., Silkenat, S., and Kisker, C. (2018). Structural basis for the recruitment of Ctf18-RFC to the replisome. Structure 26, this issue, 137–144. Mayer, M.L., Gygi, S.P., Aebersold, R., and Hieter, P. (2001). Identification of RFC(Ctf18p, Ctf8p, Dcc1p): an alternative RFC complex required for sister chromatid cohesion in S. cerevisiae. Mol. Cell 7, 959–970. Murakami, T., Takano, R., Takeo, S., Taniguchi, R., Ogawa, K., Ohashi, E., and Tsurimoto, T. (2010).

Stable interaction between the human proliferating cell nuclear antigen loader complex Ctf18-replication factor C (RFC) and DNA polymerase epsivlon is mediated by the cohesion-specific subunits, Ctf18, Dcc1, and Ctf8. J. Biol. Chem. 285, 34608– 34615. Naiki, T., Kondo, T., Nakada, D., Matsumoto, K., and Sugimoto, K. (2001). Chl12 (Ctf18) forms a novel replication factor C-related complex and functions redundantly with Rad24 in the DNA replication checkpoint pathway. Mol. Cell. Biol. 21, 5838–5845. Shiomi, Y., and Nishitani, H. (2017). Control of genome integrity by RFC complexes; conductors of PCNA loading onto and unloading from chromatin during DNA replication. Genes (Basel) 8, 52. Wade, B.O., Liu, H.W., Samora, C.P., Uhlmann, F., and Singleton, M.R. (2017). Structural studies of RFCCtf18 reveal a novel chromatin recruitment role for Dcc1. EMBO Rep. 18, 558–568. Yao, N.Y., and O’Donnell, M. (2012). The RFC clamp loader: structure and function. Subcell. Biochem. 62, 259–279.

YidC: Evaluating the Importance of the Native Environment Timothy A. Cross1,* 1Department of Chemistry and Biochemistry, Institute of Molecular Biophysics and National High Magnetic Field Lab, Florida State University, Tallahassee, FL, USA *Correspondence: [email protected] https://doi.org/10.1016/j.str.2017.12.008

In this issue of Structure, Baker et al. (2018) take advantage of recent technological breakthroughs in solid-state NMR spectroscopy and electron cryogenic tomography to characterize structural and functional differences between reconstituted YidC in E. coli lipids and YidC as overexpressed in the E. coli inner cellular membrane. The vast heterogeneity and complexity of membrane protein environments influence their structure. As Christian Anfinsen commented, protein structure is dictated ‘‘by the amino acid sequence in a given environment’’ (Anfinsen, 1973). The last four words of Anfinsen’s quote are often dropped when crediting Anfinsen with recognizing that the amino acid sequence dictates a protein’s structure, thus implying that protein structure is dictated solely by the amino acid sequence. This, however, is not true, because compounds such as denaturants in the protein’s environment cause structural changes. An

article by Baker et al. (2018) in this issue of Structure examines how the native environment affects the structure of YidC, a membrane protein that facilitates membrane insertion and folding of other proteins expressed via the ribosome. The authors employ technologies of solid-state NMR spectroscopy (ssNMR) and electron cryogenic tomography (CryoET) to characterize the protein in the very complex native environment of the E. coli inner cellular membrane. ssNMR has a long history of characterizing high-resolution transmembrane peptides and proteins in liquid crystalline

2 Structure 26, January 2, 2018 ª 2017 Elsevier Ltd.

lipid bilayers (Ketchem et al., 1997), while CryoET has a history of imaging cells and, more recently, subcellular structures (Chang et al., 2016). Unlike the homogeneous environment for aqueous proteins with a uniform concentration of water, a relatively uniform dielectric constant, and uniform hydrogen-bonding capacity, the native environment for membrane proteins is exceptionally heterogeneous (Figure 1). The water concentration ranges downward from that of bulk water by maybe ten orders of magnitude at the center of the bilayer. The dielectric constant varies

Structure

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Figure 1. Dramatic Gradients across a Lipid Bilayer (A) A heterogeneous membrane environment influences membrane structure. (B) The dielectric constant (3; Nymeyer and Zhou, 2008). (C) Water concentration (M, estimated). (D) Lateral pressure (Atm; Cantor, 1999). (E) The fatty acyl chain order parameter (SCD; Seelig and Seelig, 1974).

by two orders of magnitude, from a high in the interfacial region to a low at the bilayer center (Nymeyer and Zhou, 2008). Lateral pressure models suggest that the pressure can range above 300 atmospheres at the boundary between the lipid interfacial region and the hydrophobic interior of the membrane environment (Cantor, 1999). There is also a fluidity gradient across the membrane, as described by an order parameter for the fatty acyl chain methylene groups (Seelig and Seelig, 1974). Such physical properties have a dramatic influence on the molecular interactions responsible for stabilizing the noncovalent secondary, tertiary, and quarternary structure of proteins. Since the force associated with the many different electrostatic interactions responsible for these higher levels of protein structure are inversely related to the dielectric constant, it is critical to carefully model the native environment in samples used for structural characterization (Cross et al., 2013). For instance, it is now well recognized that transmembrane helices occurring at the interface between the protein and the lipid fatty acyl environment are much more uniform in structure

than those in water soluble proteins, due to the strengthening of hydrogen bonds (Kim and Cross, 2002). In the study by Baker et al. (2018), CryoET is brought together with ssNMR, two technologies that are undergoing dramatic technological revolutions at this time. CryoET and Cryo electron microscopy (CryoEM) developments in the past few years, involving direct electron detectors, phase plates, and advanced computational tools, have resulted in near atomic resolution for CryoEM and greatly enhanced resolution on the nanometer scale in tomographic images derived from CryoET. These advances have been punctuated with a recent Chemistry Nobel Prize to Jacques Dubochet, Joachim Frank, and Richard Henderson. Less well known are the dramatic enhancements occurring in biological ssNMR, illustrated, in part, here with ssNMR proton detection. Thanks to fast magic-angle sample spinning coupled with deuteration of the protein (expressing the protein in the presence of D2O), the 1H-1H homonuclear couplings that normally lead to severe line-broadening are suppressed, resulting in narrow resonances. Further-

more, in biological systems, detection through nuclei, such as 13C or 15N, has much lower sensitivity than detection through the high gyromagnetic ratio of the 1Hs. A sparse distribution of protons can be inserted into the protein by backexchange of the amide deuterons with H2O leading to much greater sensitivity for the NMR spectra to the extent that spectra from small samples can be obtained in a couple of hours at 800 MHz. This is despite the fact that fast spinning implies a small rotor diameter (1.3 mm) and an approximately 3 mL sample volume, implying, at most, 1 mg of protein. Here, the authors are using a protein preparation that is the isolated E. coli inner membrane (e.g., Miao et al., 2012) with no purification of YidC from the other proteins and lipids of the inner membrane. In other words, all of the native proteins, in addition to the overexpressed YidC, are present. Because the YidC expression was induced at the time when the 15N and 13C isotopic labels were added, and because the expression of YidC took over the cellular machinery for expression, the vast majority of the 15N and 13C labels are in YidC and not the other membrane proteins. Consequently, the observed NMR signals are from YidC alone, but the other membrane proteins, lipids, etc. of the cellular membrane dilute the YidC, which is therefore bound to be present in the rotor at a quantity that is below 1 mg. Such sensitivity is very similar to that for solution NMR with the great added benefit that the protein is in its native environment, albeit overexpressed. The authors used CryoET to confirm that the rifampcin treatment, leading to overexpression of YidC, did not distort the native-like ultrastructure of the cell envelopes. They then obtained high-resolution proton-detected 1H-15N ssNMR spectroscopy to compare spectra of the purified and reconstituted preparation of YidC in E. coli lipids with that of the intact native-like membranes containing YidC. Although only a fraction of the resonances were resolved in these 2D spectra, it is very clear that many resonances are shifted between the native-like membrane preparation and the purified and reconstituted preparation. The native-like membranes have an asymmetric distribution of lipids between the bilayer leaflets and Structure 26, January 2, 2018 3

Structure

Previews a unique YidC orientation with respect to the bilayer leaflets, as well as a vast array of other proteins present. In contrast, the purified protein reconstituted with purified E. coli lipids forms bilayers with uniform composition for the two leaflets and hence no unique orientation for YidC with respect to this synthetic membrane environment. The comparison of the spectra from these two YidC samples shows that many resonances have the same frequency, while others are either shifted in frequency or differ in spectral intensity. The changes in spectral intensities could reflect changes in dynamics or N-H exchange rates, while the changes in frequency would be associated with structural differences in the two sample preparations. No effort was made here to make sequence-specific assignments; however, some amino acid type assignments were made. YidC interacts with some nascent protein sequences produced from ribosomes, such as subunit c of ATP synthase. Here, the authors show that the interaction of nascent subunit c is different when the YidC is in native-like membranes compared to that in previously published studies using purified and reconstituted YidC in nanodiscs of varying composition. The resulting

dissociation constants from the nanodisc fluorescent studies suggest much stronger binding than what was suggested here in the CryoET studies of the native membrane preparations. Baker et al. (2018) thus demonstrate, using both ssNMR and CryoET, the critical importance of performing structural and functional studies in preparations that maintain as closely as possible the protein’s native membrane environment. The purification and reconstitution of YidC into a symmetric bilayer—even using appropriate lipids for YidC—was not sufficient to achieve native YidC functionality. Furthermore, the revolutionary advances in technology for CryoET and ssNMR, coupled with their ability to characterize native-like sample preparations, suggest that the authors’ approach of combining these techniques will be applicable to many complex macromolecular systems in cellular membranes, leading to enhanced understanding of how these subcellular structures function. REFERENCES Anfinsen, C.B. (1973). Principles that govern the folding of protein chains. Science 181, 223–230. Baker, L.A., Sinnige, T., Schellenberger, P., deKeyzer, J., Siebert, C.A., Driessen, A.J.M., Baldus,

€newald, K. (2018). Combined 1H-deM., and Gru tected solid-state NMR spectroscopy and electron cryotomography to study membrane proteins across resolutions in native environments. Structure 26, this issue, 161–170. Cantor, R.S. (1999). Lipid composition and the lateral pressure profile in bilayers. Biophys. J. 76, 2625–2639. Chang, Y.-W., Rettberg, L.A., Treuner-Lange, A., Iwasa, J., Søgaard-Andersen, L., and Jensen, G.J. (2016). Architecture of the type IVa pilus machine. Science 351, aad2001. Cross, T.A., Murray, D.T., and Watts, A. (2013). Helical membrane protein conformations and their environment. Eur. Biophys. J. 42, 731–755. Ketchem, R., Roux, B., and Cross, T. (1997). Highresolution polypeptide structure in a lamellar phase lipid environment from solid state NMR derived orientational constraints. Structure 5, 1655–1669. Kim, S., and Cross, T.A. (2002). Uniformity, ideality, and hydrogen bonds in transmembrane a-helices. Biophys. J. 83, 2084–2095. Miao, Y., Qin, H., Fu, R., Sharma, M., Can, T.V., Hung, I., Luca, S., Gor’kov, P.L., Brey, W.W., and Cross, T.A. (2012). M2 proton channel structural validation from full-length protein samples in synthetic bilayers and E. coli membranes. Angew. Chem. Int. Ed. 51, 8383–8386. Nymeyer, H., and Zhou, H.X. (2008). A method to determine dielectric constants in nonhomogeneous systems: application to biological membranes. Biophys. J. 94, 1185–1193. Seelig, A., and Seelig, J. (1974). The dynamic structure of fatty acyl chains in a phospholipid bilayer measured by deuterium magnetic resonance. Biochemistry 13, 4839–4845.

Order within a Disordered Structure Amy H. Tien1 and Marianne D. Sadar1,* 1Genome Sciences Centre, BC Cancer, Vancouver, BC V5Z 1L3, Canada *Correspondence: [email protected] https://doi.org/10.1016/j.str.2017.12.007

The transcriptional activity of the androgen receptor is tightly regulated by an intrinsically disordered N-terminal transactivation domain. In this issue of Structure, De Mol et al. (2018) identify a motif in the disordered transactivation domain that can be induced to adopt a helical conformation essential for interaction with the transcriptional machinery. Intrinsically disordered proteins (IDPs) have the plasticity to exhibit different structures. This flexibility permits multiple interaction motifs to form numerous complexes with proteins, DNA, or RNA, to regulate various cellular functions such as transcriptional activation, translation,

and cell signaling pathways (Wright and Dyson, 2015). Such proteins can therefore act as hubs of interaction networks. A partially folded conformation is favored in many cellular processes for a variety of reasons. First, its flexibility allows different molecular interactions under

4 Structure 26, January 2, 2018 ª 2017 Published by Elsevier Ltd.

different conditions. Second, it is able to weakly bind to targets with high specificity, resulting in transient complex formation and dynamic signaling regulation. Third, conformational change is induced upon binding to targets, leading to stabilization. Finally, it allows easy