1
|
Myronidi I, Ring A, Wu F, Ljungdahl PO. ER-localized Shr3 is a selective co-translational folding chaperone necessary for amino acid permease biogenesis. J Cell Biol 2023; 222:e202208060. [PMID: 37477900 PMCID: PMC10359922 DOI: 10.1083/jcb.202208060] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/11/2022] [Revised: 04/16/2023] [Accepted: 06/26/2023] [Indexed: 07/22/2023] Open
Abstract
Proteins with multiple membrane-spanning segments (MS) co-translationally insert into the endoplasmic reticulum (ER) membrane of eukaryotic cells. Shr3, an ER membrane-localized chaperone in Saccharomyces cerevisiae, is required for the functional expression of a family of 18 amino acid permeases (AAP) comprised of 12 MS. We have used comprehensive scanning mutagenesis and deletion analysis of Shr3 combined with a modified split-ubiquitin approach to probe chaperone-substrate interactions in vivo. Shr3 selectively interacts with nested C-terminal AAP truncations in marked contrast to similar truncations of non-Shr3 substrate sugar transporters. Shr3-AAP interactions initiate with the first four MS of AAP and successively strengthen but weaken abruptly when all 12 MS are present. Shr3-AAP interactions are based on structural rather than sequence-specific interactions involving membrane and luminal domains of Shr3. The data align with Shr3 engaging nascent N-terminal chains of AAP, functioning as a scaffold to facilitate folding as translation completes.
Collapse
Affiliation(s)
- Ioanna Myronidi
- Department of Molecular Biosciences, The Wenner-Gren Institute, SciLifeLab, Stockholm University, Stockholm, Sweden
| | - Andreas Ring
- Department of Molecular Biosciences, The Wenner-Gren Institute, SciLifeLab, Stockholm University, Stockholm, Sweden
| | - Fei Wu
- Department of Biochemistry and Biophysics, SciLifeLab, Stockholm University, Stockholm, Sweden
| | - Per O. Ljungdahl
- Department of Molecular Biosciences, The Wenner-Gren Institute, SciLifeLab, Stockholm University, Stockholm, Sweden
| |
Collapse
|
2
|
Atomic Force Microscopy Reveals Complexity Underlying General Secretory System Activity. Int J Mol Sci 2022; 24:ijms24010055. [PMID: 36613499 PMCID: PMC9820662 DOI: 10.3390/ijms24010055] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/11/2022] [Revised: 12/08/2022] [Accepted: 12/09/2022] [Indexed: 12/24/2022] Open
Abstract
The translocation of specific polypeptide chains across membranes is an essential activity for all life forms. The main components of the general secretory (Sec) system of E. coli include integral membrane translocon SecYEG, peripheral ATPase SecA, and SecDF, an ancillary complex that enhances polypeptide secretion by coupling translocation to proton motive force. Atomic force microscopy (AFM), a single-molecule imaging technique, is well suited to unmask complex, asynchronous molecular activities of membrane-associated proteins including those comprising the Sec apparatus. Using AFM, the dynamic structure of membrane-external protein topography of Sec system components can be directly visualized with high spatial-temporal precision. This mini-review is focused on AFM imaging of the Sec system in near-native fluid conditions where activity can be maintained and biochemically verified. Angstrom-scale conformational changes of SecYEG are reported on 100 ms timescales in fluid lipid bilayers. The association of SecA with SecYEG, forming membrane-bound SecYEG/SecA translocases, is directly visualized. Recent work showing topographical aspects of the translocation process that vary with precursor species is also discussed. The data suggests that the Sec system does not employ a single translocation mechanism. We posit that differences in the spatial frequency distribution of hydrophobic content within precursor sequences may be a determining factor in mechanism selection. Precise AFM investigations of active translocases are poised to advance our currently vague understanding of the complicated macromolecular movements underlying protein export across membranes.
Collapse
|
3
|
Metelev M, Lundin E, Volkov IL, Gynnå AH, Elf J, Johansson M. Direct measurements of mRNA translation kinetics in living cells. Nat Commun 2022; 13:1852. [PMID: 35388013 PMCID: PMC8986856 DOI: 10.1038/s41467-022-29515-x] [Citation(s) in RCA: 11] [Impact Index Per Article: 5.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/01/2021] [Accepted: 03/17/2022] [Indexed: 01/09/2023] Open
Abstract
Ribosome mediated mRNA translation is central to life. The cycle of translation, however, has been characterized mostly using reconstituted systems, with only few techniques applicable for studies in the living cell. Here we describe a live-cell ribosome-labeling method, which allows us to characterize the whole processes of finding and translating an mRNA, using single-molecule tracking techniques. We find that more than 90% of both bacterial ribosomal subunits are engaged in translation at any particular time, and that the 30S and 50S ribosomal subunits spend the same average time bound to an mRNA, revealing that 30S re-initiation on poly-cistronic mRNAs is not prevalent in E. coli. Instead, our results are best explained by substantial 70S re-initiation of translation of poly-cistronic mRNAs, which is further corroborated by experiments with translation initiation inhibitors. Finally, we find that a variety of previously described orthogonal ribosomes, with altered anti-Shine-Dalgarno sequences, show significant binding to endogenous mRNAs.
Collapse
Affiliation(s)
- Mikhail Metelev
- Department of Cell and Molecular Biology, Uppsala University, Uppsala, Sweden
| | - Erik Lundin
- Department of Cell and Molecular Biology, Uppsala University, Uppsala, Sweden
| | - Ivan L Volkov
- Department of Cell and Molecular Biology, Uppsala University, Uppsala, Sweden
| | - Arvid H Gynnå
- Department of Cell and Molecular Biology, Uppsala University, Uppsala, Sweden
| | - Johan Elf
- Department of Cell and Molecular Biology, Uppsala University, Uppsala, Sweden
| | - Magnus Johansson
- Department of Cell and Molecular Biology, Uppsala University, Uppsala, Sweden.
| |
Collapse
|
4
|
Lyu Z, Genereux JC. Methodologies for Measuring Protein Trafficking across Cellular Membranes. Chempluschem 2021; 86:1397-1415. [PMID: 34636167 DOI: 10.1002/cplu.202100304] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/08/2021] [Revised: 08/19/2021] [Indexed: 12/11/2022]
Abstract
Nearly all proteins are synthesized in the cytosol. The majority of this proteome must be trafficked elsewhere, such as to membranes, to subcellular compartments, or outside of the cell. Proper trafficking of nascent protein is necessary for protein folding, maturation, quality control and cellular and organismal health. To better understand cellular biology, molecular and chemical technologies to properly characterize protein trafficking (and mistrafficking) have been developed and applied. Herein, we take a biochemical perspective to review technologies that enable spatial and temporal measurement of protein distribution, focusing on both the most widely adopted methodologies and exciting emerging approaches.
Collapse
Affiliation(s)
- Ziqi Lyu
- Department of Chemistry, University of California, Riverside, 501 Big Springs Road, 92521, Riverside, CA, USA
| | - Joseph C Genereux
- Department of Chemistry, University of California, Riverside, 501 Big Springs Road, 92521, Riverside, CA, USA
| |
Collapse
|
5
|
Hao W, Cui W, Cheng Z, Han L, Suo F, Liu Z, Zhou L, Zhou Z. Development of a base editor for protein evolution via in situ mutation in vivo. Nucleic Acids Res 2021; 49:9594-9605. [PMID: 34390349 PMCID: PMC8450078 DOI: 10.1093/nar/gkab673] [Citation(s) in RCA: 6] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/16/2021] [Revised: 07/21/2021] [Accepted: 07/24/2021] [Indexed: 12/16/2022] Open
Abstract
Protein evolution has significantly enhanced the development of life science. However, it is difficult to achieve in vitro evolution of some special proteins because of difficulties with heterologous expression, purification, and function detection. To achieve protein evolution via in situ mutation in vivo, we developed a base editor by fusing nCas with a cytidine deaminase in Bacillus subtilis through genome integration. The base editor introduced a cytidine-to-thymidine mutation of approximately 100% across a 5 nt editable window, which was much higher than those of other base editors. The editable window was expanded to 8 nt by extending the length of sgRNA, and conversion efficiency could be regulated by changing culture conditions, which was suitable for constructing a mutant protein library efficiently in vivo. As proof-of-concept, the Sec-translocase complex and bacitracin-resistance-related protein BceB were successfully evolved in vivo using the base editor. A Sec mutant with 3.6-fold translocation efficiency and the BceB mutants with different sensitivity to bacitracin were obtained. As the construction of the base editor does not rely on any additional or host-dependent factors, such base editors (BEs) may be readily constructed and applicable to a wide range of bacteria for protein evolution via in situ mutation.
Collapse
Affiliation(s)
- Wenliang Hao
- The Key Laboratory of Industrial Biotechnology (Ministry of Education), School of Biotechnology, Jiangnan University, 1800 Lihu Avenue, Wuxi 214122, China
| | - Wenjing Cui
- The Key Laboratory of Industrial Biotechnology (Ministry of Education), School of Biotechnology, Jiangnan University, 1800 Lihu Avenue, Wuxi 214122, China
| | - Zhongyi Cheng
- The Key Laboratory of Industrial Biotechnology (Ministry of Education), School of Biotechnology, Jiangnan University, 1800 Lihu Avenue, Wuxi 214122, China
| | - Laichuang Han
- The Key Laboratory of Industrial Biotechnology (Ministry of Education), School of Biotechnology, Jiangnan University, 1800 Lihu Avenue, Wuxi 214122, China
| | - Feiya Suo
- The Key Laboratory of Industrial Biotechnology (Ministry of Education), School of Biotechnology, Jiangnan University, 1800 Lihu Avenue, Wuxi 214122, China
| | - Zhongmei Liu
- The Key Laboratory of Industrial Biotechnology (Ministry of Education), School of Biotechnology, Jiangnan University, 1800 Lihu Avenue, Wuxi 214122, China
| | - Li Zhou
- The Key Laboratory of Industrial Biotechnology (Ministry of Education), School of Biotechnology, Jiangnan University, 1800 Lihu Avenue, Wuxi 214122, China
| | - Zhemin Zhou
- The Key Laboratory of Industrial Biotechnology (Ministry of Education), School of Biotechnology, Jiangnan University, 1800 Lihu Avenue, Wuxi 214122, China
| |
Collapse
|
6
|
Davis MM, Lamichhane R, Bruce BD. Elucidating Protein Translocon Dynamics with Single-Molecule Precision. Trends Cell Biol 2021; 31:569-583. [PMID: 33865650 DOI: 10.1016/j.tcb.2021.03.009] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/11/2020] [Revised: 03/17/2021] [Accepted: 03/18/2021] [Indexed: 01/28/2023]
Abstract
Translocons are protein assemblies that facilitate the targeting and transport of proteins into and across biological membranes. Our understanding of these systems has been advanced using genetics, biochemistry, and structural biology. Despite these classic advances, until recently we have still largely lacked a detailed understanding of how translocons recognize and facilitate protein translocation. With the advent and improvements of cryogenic electron microscopy (cryo-EM) single-particle analysis and single-molecule fluorescence microscopy, the details of how translocons function are finally emerging. Here, we introduce these methods and evaluate their importance in understanding translocon structure, function, and dynamics.
Collapse
Affiliation(s)
- Madeline M Davis
- Department of Biochemistry and Cellular and Molecular Biology, University of Tennessee at Knoxville, Knoxville, TN 37996, USA
| | - Rajan Lamichhane
- Department of Biochemistry and Cellular and Molecular Biology, University of Tennessee at Knoxville, Knoxville, TN 37996, USA
| | - Barry D Bruce
- Department of Biochemistry and Cellular and Molecular Biology, University of Tennessee at Knoxville, Knoxville, TN 37996, USA; Department of Microbiology, University of Tennessee at Knoxville, Knoxville, TN 37996, USA; Graduate Program in Genome Science and Technology, University of Tennessee at Knoxville, Knoxville, TN 37996, USA; Chemical and Biomolecular Engineering, University of Tennessee at Knoxville, Knoxville, TN 37996, USA.
| |
Collapse
|
7
|
Itskanov S, Kuo KM, Gumbart JC, Park E. Stepwise gating of the Sec61 protein-conducting channel by Sec63 and Sec62. Nat Struct Mol Biol 2021; 28:162-172. [PMID: 33398175 PMCID: PMC8236211 DOI: 10.1038/s41594-020-00541-x] [Citation(s) in RCA: 34] [Impact Index Per Article: 11.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/28/2020] [Accepted: 11/12/2020] [Indexed: 12/12/2022]
Abstract
Many proteins are transported into the endoplasmic reticulum by the universally conserved Sec61 channel. Post-translational transport requires two additional proteins, Sec62 and Sec63, but their functions are poorly defined. Here, we determined cryo-EM structures of several variants of Sec61–Sec62–Sec63 complexes from Saccharomyces cerevisiae and Thermomyces lanuginosus and show that Sec62 and Sec63 induce opening of the Sec61 channel. Without Sec62, the translocation pore of Sec61 remains closed by the plug domain, rendering the channel inactive. We further show that the lateral gate of Sec61 must first be partially opened by interactions between Sec61 and Sec63 in cytosolic and lumenal domains, a simultaneous disruption of which completely closes the channel. The structures and molecular dynamics simulations suggest that Sec62 may also prevent lipids from invading the channel through the open lateral gate. Our study shows how Sec63 and Sec62 work together in a hierarchical manner to activate Sec61 for post-translational protein translocation.
Collapse
Affiliation(s)
- Samuel Itskanov
- Biophysics Graduate Program, University of California, Berkeley, CA, USA
| | - Katie M Kuo
- School of Chemistry and Biochemistry, Georgia Institute of Technology, Atlanta, GA, USA
| | - James C Gumbart
- School of Chemistry and Biochemistry, Georgia Institute of Technology, Atlanta, GA, USA.,School of Physics, Georgia Institute of Technology, Atlanta, GA, USA
| | - Eunyong Park
- Department of Molecular and Cell Biology, University of California, Berkeley, CA, USA. .,California Institute for Quantitative Biosciences, University of California, Berkeley, CA, USA.
| |
Collapse
|
8
|
Catipovic MA, Rapoport TA. Protease protection assays show polypeptide movement into the SecY channel by power strokes of the SecA ATPase. EMBO Rep 2020; 21:e50905. [PMID: 32969592 DOI: 10.15252/embr.202050905] [Citation(s) in RCA: 10] [Impact Index Per Article: 2.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/14/2020] [Revised: 08/31/2020] [Accepted: 09/01/2020] [Indexed: 12/18/2022] Open
Abstract
Bacterial secretory proteins are translocated post-translationally by the SecA ATPase through the protein-conducting SecY channel in the plasma membrane. During the ATP hydrolysis cycle, SecA undergoes large conformational changes of its two-helix finger and clamp domains, but how these changes result in polypeptide movement is unclear. Here, we use a reconstituted purified system and protease protection assays to show that ATP binding to SecA results in a segment of the translocation substrate being pushed into the channel. This motion is prevented by mutation of conserved residues at the finger's tip. Mutation of SecA's clamp causes backsliding of the substrate in the ATP-bound state. Together, these data support a power stroke model of translocation in which, upon ATP binding, the two-helix finger pushes the substrate into the channel, where it is held by the clamp until nucleotide hydrolysis has occurred.
Collapse
Affiliation(s)
- Marco A Catipovic
- Howard Hughes Medical Institute, Harvard Medical School, Boston, MA, USA.,Department of Cell Biology, Harvard Medical School, Boston, MA, USA
| | - Tom A Rapoport
- Howard Hughes Medical Institute, Harvard Medical School, Boston, MA, USA.,Department of Cell Biology, Harvard Medical School, Boston, MA, USA
| |
Collapse
|
9
|
Chattrakun K, Hoogerheide DP, Mao C, Randall LL, King GM. Protein Translocation Activity in Surface-Supported Lipid Bilayers. LANGMUIR : THE ACS JOURNAL OF SURFACES AND COLLOIDS 2019; 35:12246-12256. [PMID: 31448613 PMCID: PMC10906442 DOI: 10.1021/acs.langmuir.9b01928] [Citation(s) in RCA: 9] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/10/2023]
Abstract
Surface-supported lipid bilayers are used widely throughout the nanoscience community as cellular membrane mimics. For example, they are frequently employed in single-molecule atomic force microscopy (AFM) studies to shed light on membrane protein conformational dynamics and folding. However, in AFM as well as in other surface-sensing techniques, the close proximity of the supporting surface raises questions about preservation of the biochemical activity. Employing the model translocase from the general secretory (Sec) system of Escherichia coli, here we quantify the activity via two biochemical assays in surface-supported bilayers. The first assesses ATP hydrolysis and the second assesses polypeptide translocation across the membrane via protection from added protease. Hydrolysis assays revealed distinct levels of activation ranging from medium (translocase-activated) to high (translocation-associated) that were similar to traditional solution experiments and further identified an adenosine triphosphatase population exhibiting characteristics of conformational hysteresis. Translocation assays revealed turn over numbers that were comparable to solution but with a 10-fold reduction in apparent rate constant. Despite differences in kinetics, the chemomechanical coupling (ATP hydrolyzed per residue translocated) only varied twofold on glass compared to solution. The activity changed with the topographic complexity of the underlying surface. Rough glass coverslips were favored over atomically flat mica, likely due to differences in frictional coupling between the translocating polypeptide and surface. Neutron reflectometry and AFM corroborated the biochemical measurements and provided structural characterization of the submembrane space and upper surface of the bilayer. Overall, the translocation activity was maintained for the surface-adsorbed Sec system, albeit with a slower rate-limiting step. More generally, polypeptide translocation activity measurements yield valuable quantitative metrics to assess the local environment about surface-supported lipid bilayers.
Collapse
Affiliation(s)
- Kanokporn Chattrakun
- Department of Physics and Astronomy, University of Missouri-Columbia, Columbia, Missouri 65211, United States
| | - David P. Hoogerheide
- Center for Neutron Research, National Institute of Standards and Technology, Gaithersburg, Maryland 20899, United States
| | - Chunfeng Mao
- Department of Biochemistry, University of Missouri-Columbia, Columbia, Missouri 65211, United States
| | - Linda L. Randall
- Department of Biochemistry, University of Missouri-Columbia, Columbia, Missouri 65211, United States
| | - Gavin M. King
- Department of Physics and Astronomy, University of Missouri-Columbia, Columbia, Missouri 65211, United States
- Department of Biochemistry, University of Missouri-Columbia, Columbia, Missouri 65211, United States
| |
Collapse
|
10
|
Sikkema HR, Gaastra BF, Pols T, Poolman B. Cell Fuelling and Metabolic Energy Conservation in Synthetic Cells. Chembiochem 2019; 20:2581-2592. [PMID: 31381223 DOI: 10.1002/cbic.201900398] [Citation(s) in RCA: 24] [Impact Index Per Article: 4.8] [Reference Citation Analysis] [Abstract] [Key Words] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/18/2019] [Indexed: 12/14/2022]
Abstract
We are aiming for a blue print for synthesizing (moderately complex) subcellular systems from molecular components and ultimately for constructing life. However, without comprehensive instructions and design principles, we rely on simple reaction routes to operate the essential functions of life. The first forms of synthetic life will not make every building block for polymers de novo according to complex pathways, rather they will be fed with amino acids, fatty acids and nucleotides. Controlled energy supply is crucial for any synthetic cell, no matter how complex. Herein, we describe the simplest pathways for the efficient generation of ATP and electrochemical ion gradients. We have estimated the demand for ATP by polymer synthesis and maintenance processes in small cell-like systems, and we describe circuits to control the need for ATP. We also present fluorescence-based sensors for pH, ionic strength, excluded volume, ATP/ADP, and viscosity, which allow the major physicochemical conditions inside cells to be monitored and tuned.
Collapse
Affiliation(s)
- Hendrik R Sikkema
- Department of Biochemistry, University of Groningen, Nijenborgh 4, 9747 AG, Groningen, The Netherlands
| | - Bauke F Gaastra
- Department of Biochemistry, University of Groningen, Nijenborgh 4, 9747 AG, Groningen, The Netherlands
| | - Tjeerd Pols
- Department of Biochemistry, University of Groningen, Nijenborgh 4, 9747 AG, Groningen, The Netherlands
| | - Bert Poolman
- Department of Biochemistry, University of Groningen, Nijenborgh 4, 9747 AG, Groningen, The Netherlands
| |
Collapse
|