MIAMI UNIVERSITY the Graduate School

MIAMI UNIVERSITY The Graduate School

Certificate for Approving the Dissertation

We hereby approve the Dissertation

Christopher Paul New

Candidate for the Degree

DOCTOR OF PHILOSOPHY

______Carole Dabney-Smith, Director

______Rick Page, Chair

______Ann Hagerman, Reader

______Gary Lorigan, Reader

______Rachael Morgan-Kiss, Graduate School Representative

ABSTRACT

ANALYSIS OF THA4 FUNCTION AND ORGANIZATION IN CHLOROPLAST TWIN ARGININE TRANSPORT

Christopher P. New

The chloroplast Twin Arginine Translocase (cpTAT) system transports fully folded proteins across the thylakoid membrane in plant cells using only energy derived from the proton motive force (PMF). Three membrane bound component proteins: cpTatC, Hcf106, and Tha4 function together in a transient manner to accomplish transport. However, clear mechanistic details of this process remain elusive such as how cpTAT utilizes energy stored in the PMF or how the individual component proteins interact during each step of transport. In addition, prior structural characterization of (cp)TAT proteins used truncated versions of the components. This dissertation describes work to develop methods to purify full-length Hcf106 for biophysical characterization. Additionally, this dissertation details the work to determine the function of a membrane embedded glutamate in the Tha4 transmembrane helix (TMH).

A series of purification trials were carried out to isolate Hcf106 fused to maltose binding protein (MBP) by the recognition sequence of tobacco etch virus protease (TEVp). Fusion protein and protease were expressed in and purified from E. coli using affinity chromatography. Multiple parameters and additives were tested during optimization of TEVp proteolysis reactions with MBP-Hcf106. TEVp and free MBP were separated from un-cleaved MBP-Hcf106 and free Hcf106 by affinity and size exclusion chromatography. Although TEVp and free MBP were removed after an optimized proteolysis reaction, free Hcf106 showed its recalcitrant nature through resistance of separation from un-cleaved MBP-Hcf106 by size exclusion chromatography in several detergent and buffer conditions.

To better understand the role of the membrane embedded Tha4 glutamate 10 (E10), Tha4 variants with glutamate to alanine (E10A) or glutamate to aspartate (E10D) substitutions were used to complement loss of cpTAT function in thylakoid membranes. Sequential glutamate substitutions in the TMH of Tha4 variant E10A were unable to restore transport while aspartate substitutions were mildly able to complement loss of function. Furthermore, organization between three structural regions in Tha4 E10/A/D variants was determined by disulfide crosslinking during various transport conditions. Tha4 E10/A/D variant oligomer formation was enhanced in the presence of functional precursor with and without PMF present. An increase in TMH hydrophobicity by alanine substitution was shown to increase Tha4 stability in isolated thylakoid membranes and to promote tighter packing interactions between adjacent Tha4 monomers. The interaction data was then used to develop a model of how Tha4 E10/A/D variant tetramers pack and reorganize in the presence of precursor.

ANALYSIS OF THA4 FUNCTION AND ORGANIZATION IN CHLOROPLAST TWIN ARGININE TRANSPORT

A DISSERTATION

Presented to the Faculty of

Miami University in partial

fulfillment of the requirements

for the degree of

Doctor of Philosophy

Department of Chemistry and Biochemistry

Christopher P. New

The Graduate School Miami University Oxford, Ohio

2020

Dissertation Director: Carole Dabney-Smith

Christopher Paul New

2020

TABLE OF CONTENTS

Chapter 1: Introduction, Routing of Thylakoid Lumen Proteins by the Chloroplast Twin Arginine Transport Pathway ...... 1 1.1 Abstract ...... 2 1.2 Introduction ...... 2 1.3 Twin Arginine Translocase: Protein components and functional requirements ...... 3 1.3.1 Protein components ...... 3 1.3.1.1 cpTatC (TatC) ...... 3 1.3.1.2 Tha4 (TatA) and Hcf106 (TatB) ...... 4 1.3.2 Functional requirements ...... 9 1.3.2.1 Precursor signal sequence ...... 9 1.3.2.2 Energetics ...... 10 1.4 Molecular mechanism of transport ...... 11 1.4.1 Receptor complex and precursor binding ...... 11 1.4.2 Translocase assembly ...... 12 1.4.3 Models of translocation ...... 13 1.5 Future perspectives ...... 16 1.6 Dissertation goals and specific aims ...... 17 1.7 References...... 19 Chapter 2: Purification of Maltose Binding Protein-Hcf106 Fusion for Structural Characterization ...... 26 2.1 Abstract ...... 27 2.2 Introduction ...... 27 2.3 Materials and Methods ...... 30 2.3.1 Generation of pMAL-c5E with TEV protease recognition sequence between MBP and full-length Hcf106 sequences ...... 30 2.3.2 MBP-Hcf106 protein expression ...... 30 2.3.3 Purification of MBP-Hcf106 fusion protein ...... 31

2.3.4 TEVpS219VHis6 expression and purification ...... 31 2.3.5 TEV protease cleavage of MBP-Hcf106 ...... 32 2.3.6 Fast protein liquid chromatography purification trials ...... 32 2.3.7 Amylose resin batchwise TEVp cleavage reactions of bound MBP-Hcf106 ...... 32

iii 2.3.8 Removal of TEV protease by Ni-NTA magnetic beads ...... 33 2.3.9 Western blot analysis of SDS-PAGE resolved proteins ...... 33 2.4 Results ...... 33 2.4.1 TEV protease cleavage recognition sequence and the mature Hcf106 sequence from garden pea were cloned into a pMAL-c5E vector ...... 33 2.4.2 Recombinant TEV protease was purified from BL21(DE3) codon plus E. coli ...... 35 2.4.3 Isolation and purification of recombinant MBP-Hcf106 from BL21(DE3) codon plus E. coli ...... 37 2.4.4 Testing and optimization of TEVp proteolysis of MBP-Hcf106 ...... 38 2.4.5 FPLC purification tests with post TEV protease-MBP-HCF106 reaction products ...... 39 2.4.6 TEV protease cleavage reactions with MBP-Hcf106 bound to amylose resin ...... 42 2.4.7 Additional proteolysis reaction tests in the presence of urea ...... 43 2.4.8 Testing additional detergents used in previously published purification trials of recombinant TatB from E. coli ...... 46 2.5 Discussion...... 46 2.6 Conclusions and Future Directions ...... 49 2.7 References...... 50 Chapter 3: Increases in Tha4 Transmembrane Helix Hydrophobicity Alter Function and Organization During Chloroplast Twin Arginine Transport ...... 53 3.1 Abstract ...... 54 3.2 Introduction ...... 54 3.3 Materials and Methods ...... 56 3.3.1 Source Plants, Chloroplast and Thylakoid Membrane Isolation ...... 56 3.3.2 Synthesis and in vitro translation of Tha4 variants, DT23, cpTatCaaa L231C, and (V-20F)tOE17 ...... 58

3.3.3 Overexpression and purification of KKtOE17His6 ...... 58 3.3.4 Functional replacement of endogenous Tha4 and complementation of cpTAT ...... 59 3.3.5 N-ethylmaleimide (NEM) blocking of endogenous cysteine residues ..... 59 3.3.6 Oxidative cross-linking between dual cysteine substituted variants ...... 59 3.3.7 Blue Native PAGE and western blot analysis ...... 60 3.3.8 Alkaline extraction of thylakoid membrane integrated Tha4 variants ...... 60 3.3.9 Import of cpTatCaaa L231C into chloroplasts and crosslinking between single cysteine Tha4 variants in thylakoid membranes ...... 61

iv 3.4 Results ...... 62 3.4.1 Tha4 from higher plants has a conserved glutamate at the 10th position in its primary sequence and is in the hydrophobic core of the thylakoid membrane ...... 62 3.4.2 Sequential glutamate substitutions in the TMH of Tha4 E10A fail to complement loss of function in αTha4 IgG treated thylakoids ...... 62 3.4.3 Aspartate substitutions in the TMH of Tha4 E10A variant weakly complement loss of function in αTha4 IgG treated thylakoids ...... 63 3.4.4 The presence of functional precursor influences Tha4 organization more so than PMF in purified thylakoid membranes ...... 65 3.4.5 Tha4 oligomer enhancement was also tested in the presence of functional precursor with urea and non-functional precursor ...... 66 3.4.6 BN-PAGE analysis revealed extensive interactions between Tha4 E10/A variants and cpTAT components ...... 69 3.4.7 The stability of thylakoid membrane integrated cys-free and double cysteine Tha4 E10/A/D variants was determined by alkaline extraction ...... 69 3.4.8 Preliminary data of crosslinking interactions between Tha4 TMH and cpTatC TM4 in the presence and absence of precursor and PMF ...... 71 3.5 Discussion...... 72 3.6 Conclusions and Future Directions ...... 77 3.7 References...... 78 Chapter 4: Conclusions ...... 83 4.1 Conclusions ...... 84 4.2 References...... Error! Bookmark not defined.

LIST OF TABLES Table 1.1 Summary of crosslink interactions between cpTAT or bacterial TAT component proteins and precursor proteins…………………………………….....6-7

Table 2.1 BCA assay determination of MBP-Hcf106 concentration following elution from amylose resin column………………………………………………...... 38

Table 3.1 BCA assay determination of KKtOE17His6 concentration after purification, pooling, and buffer exchange…………………………………………..68

LIST OF FIGURES Figure 1.1 Models of cpTatC, Hcf106, and Tha4 oriented in a thylakoid/plasma membrane generated in Chimera using sequence homology with identified TAT components (gray) as shown...... 4

Figure 1.2 Structural modeling of Hcf106-cpTatC dimer under resting state...... 5

Figure 1.3 Thylakoid precursors containing the twin arginine motif show little sequence homology in the signal peptide...... 9

Figure 1.4 Possible models for Tha4 and Hcf106 packing with cpTatC in the presence of signal peptide or precursor...... 14

Figure 1.5 Proposed model of cpTAT function...... 16

Figure 2.1 Cartoon diagrams of Hcf106 and the engineered MBP-TEVp recognition site-Hcf106 fusion protein...... 28

Figure 2.2 Insertion into and confirmation of mature Hcf106 sequence in pMAL- TEV recognition vector by various PCR techniques...... 34

Figure 2.3 Expression and purification of TEVp...... 35

Figure 2.4 Expression and purification of MBP-Hcf106...... 36

Figure 2.5 Optimization of MBP-Hcf106 cleavage by TEVp...... 39

Figure 2.6 FPLC purification trials of TEVp and MBP-Hcf106 reactions in CHAPS or C12E9 detergents...... 40

Figure 2.7 Buffer exchange and batchwise amylose resin proteolysis of MBP- Hcf106 with TEVp...... 42

Figure 2.8 Testing additional parameters for the TEVp reaction with MBP-Hcf106 including different urea concentrations and protease ratios...... 43

Figure 2.9 Removal of TEVp from the reaction mixture and subsequent purification by FPLC...... 44

Figure 2.10 Incubation of MBP-Hcf106/Hcf106 and subsequent FPLC separation with 4 M urea...... 45

Figure 2.11 Testing reaction efficiency in the presence of additional detergents...... 46

vii Figure 3.1 Tha4 has a membrane embedded glutamate in its TMH as shown by cartoon structures modeled alone in the thylakoid membrane and during interaction with receptor complex...... 57

Figure 3.2 Glutamate substitutions in the TMH of Tha4 variant E10A are unable to complement loss of cpTAT function...... 63

Figure 3.3 Aspartate substitutions in TMH of Tha4 variant E10A mildly complement loss of cpTAT function...... 64

Figure 3.4 Tha4 E10/A/D variant organization in thylakoid membranes as determined by crosslink formation between three separate structural regions.... 65

Figure 3.5 Effect of urea and functional precursor on organization of Tha4 E10/A/D variants in thylakoid membranes...... 67

Figure 3.6 Purification of non-functional precursor and its effect on Tha4 E10/A/D variant organization in thylakoid membranes...... 68

Figure 3.7 Interactions between cys-free Tha4 E10/A/D variants examined by Blue native PAGE...... 69

Figure 3.8 Cys-free and double cysteine Tha4 E10/A/D variant stability in thylakoid membranes was determined by alkaline extraction assays...... 70

Figure 3.9 Eisenberg hydrophobicity values of cys-free and double cysteine Tha4 E10/A/D variants...... 71

Figure 3.10 Preliminary crosslinking data between Tha4 TMH and cpTatC TM4 in thylakoid membranes...... 73

Figure 3.11 Models of Tha4 E10/A/D packing and reorganization in the presence of functional precursor...... 76

viii

DEDICATION

This dissertation is dedicated to my love, best friend, and inspiration Tiffany, my mother Ronda Murphy, my grandparents Ron and Barb Murphy, my aunts and uncle Karen Kessen and Paula and Chris Mate, and my brothers David and Luke.

ACKNOWLEDGEMENTS

I want to thank everyone who has helped me along the path to complete this work. First and foremost, I want to thank my advisor, Dr. Carole Dabney-Smith, for her limitless support, patience, motivation, and guidance throughout my graduate and undergraduate research career. Thank you for encouraging me to take ownership of my research projects and teaching me to never give up on myself.

Of equal importance is the love and support I have received from my family and friends. Tiffany, you are the love of my life. Thank you for your boundless love, encouragement, and support throughout this long journey. Ronda, Chris, Paula, David, Luke, Karen, Kara, and Jake, thank you for your love, encouragement, and endless support in every way imaginable. I also want to thank my Hamilton crew of friends: Aaron Pearce, Amber Anderson, Josh Folino, Laura Cook, Will Brandenburg and Andrea Brown, Alan Hawkins, Mike Shalloe, Ben and Lisa Shalloe, John Trent, Nathan Ricketts, and Steven Asher. I couldn’t have completed this journey without you all.

I also want to thank every former and present member of the Dabney-Smith lab as well as my colleagues from other Miami research groups. Martin, thank you for teaching me how to do things the right way, your way! Drs. Qianqian Ma, Amanda Storm, Debjani Pal, Lei Zhang, Gunjan Dixit, and Indra Sahu, thanks for your friendship, kindness, endless support, and encouragement while also answering the thousands of questions I know I asked each of you. Thank you Ramja Sritharan, Krystina Hird, Jorge Escobar, Katie Eudy, Thai Wright, Vidusha Weesinghe, Thilini Kankanamalage, Jessica Pax, and Andrew Abata for your friendship, kindness, advice, and humor throughout our shared time at Miami. I wish you all the best of luck!

Finally, I would like to thank my dissertation committee for their guidance, encouragement, and support throughout my graduate studies: Drs. Ann Hagerman, Rick Page, Gary Lorigan, and Rachael Morgan-Kiss.

x Chapter 1: Introduction, Routing of Thylakoid Lumen Proteins by the Chloroplast Twin Arginine Transport Pathway

Christopher Paul New,1 Qianqian Ma,1 and Carole Dabney-Smith1,2*

1Cell, Molecular, Structural Biology Graduate Program and 2Department of Chemistry and Biochemistry, Miami University, Oxford, OH 45056

*Corresponding author: Department of Chemistry and Biochemistry, Miami University, 651 East High St., Oxford, OH 45056. Tel.: 513-529-8091; E-mail: [email protected]

This work was originally published in Photosynthesis Research 2018, 138(3): 289-301

1.1 Abstract Thylakoids are complex sub-organellar membrane systems whose role in photosynthesis makes them critical to life. Thylakoids require the coordinated expression of both nuclear- and plastid-encoded proteins to allow rapid response to changing environmental conditions. Transport of cytoplasmically-synthesized proteins to thylakoids or the thylakoid lumen is complex; the process involves transport across up to three membrane systems with routing through three aqueous compartments. Protein transport in thylakoids is accomplished by conserved ancestral prokaryotic plasma membrane translocases containing novel adaptations for the sub-organellar location. This review focuses on the evolutionarily conserved chloroplast twin arginine transport (cpTAT) pathway. An overview is provided of known aspects of the cpTAT components, energy requirements, and mechanisms with a focus on recent discoveries. Some of the most exciting new studies have been in determining the structural architecture of the membrane complex involved in forming the point of passage for the precursor and binding features of the translocase components. The cpTAT system is of interest because it transports folded protein domains using only the proton motive force for energy. The implications for mechanism of translocation by recent studies focusing on interactions between membrane TAT components and with the translocating precursor will be discussed.

1.2 Introduction Chloroplasts are dependent upon nuclear gene expression for their function and as a result have specific pathways to ensure correct import and routing of proteins into the organelle. Chloroplasts often contain upwards of 3000 individual proteins (Peltier et al., 2000), of which the majority are nuclear-encoded and found in the stroma. Correct import and routing of cytoplasmically-expressed chloroplast proteins involve synthesis of proteins as higher molecular weight precursors containing cleavable, N-terminal extensions called transit peptides, which play a role in directing or targeting the precursor to the chloroplast. Import and routing into the chloroplast may require the precursor to cross up to three membrane systems (outer chloroplast membrane, inner chloroplast membrane, and thylakoid) to reach one of three aqueous compartments (the intermembrane space, the stroma, or the thylakoid lumen). Proteins destined for the thylakoid lumen use bipartite transit peptides containing a stromal targeting domain, which promotes import into the stroma, and a lumen targeting domain, which promotes transport across the thylakoid membrane to the lumen. After successful routing of the precursors, the N-terminal extension is cleaved by specific endopeptidases. The thylakoid lumen contains 80-150 different proteins (Jarvi et al., 2013; Leister and Schneider, 2003) all of which are nuclear-encoded and must be correctly routed to the compartment to reach their site of function. Two membrane-bound protein transport systems are found in the thylakoid to assist lumen proteins across the membrane: the chloroplast secretory (cpSec) pathway and the chloroplast twin arginine transport (cpTAT) pathway. The cpTAT pathway is evolutionarily conserved with TAT pathways found in the plasma membranes of many prokaryotes and the mitochondrial inner membrane of some plants. The TAT pathway is so named because of twin arginines found in the targeting signal sequence. A defining feature of the TAT pathway is the capability to transport precursors with folded domains while requiring only the proton

2 motive force (PMF) as the source of energy; i.e., independent of NTP hydrolysis (Braun et al., 2007). The cpTAT pathway (or TAT in Escherichia coli) comprises three membrane components, cpTatC (TatC), Hcf106 (TatB), and Tha4 (TatA) that work together to promote the transport of precursors across the membrane. The first cpTAT component was identified in a maize mutant containing high chlorophyll fluorescence (hcf106) (Settles et al., 1997), which quickly led to the identification of homologous genes and related operons in bacteria (Bogsch et al., 1998). We now know that the TAT pathway is widely found in prokaryotes and prokaryote-derived organelles, particularly those in photosynthetic organisms (Carrie et al., 2016; Dilks et al., 2003; Palmer and Berks, 2012). The TAT system is predicted to be responsible for transport of <10% of the total secretome in E. coli, most often transporting metal cofactor binding proteins that use cytoplasmic machinery for proper protein folding around the cofactor (Palmer and Berks, 2012). Interestingly, about 50% of thylakoid lumen proteins are predicted to use the cpTAT pathway and only few of those bind cofactors. One likely reason for maintaining the TAT pathway in chloroplasts may be that some substrates destined for the thylakoid lumen may fold rapidly in the stroma making them difficult to unfold for translocation by the cpSec system. Studies using either chloroplasts or prokaryotes, such as E. coli, have provided insight into the mechanistic capabilities of the system. As we learn more about the mechanistic capabilities, we are only now beginning to appreciate nuanced differences for each system as it is adapted to its unique environment

1.3 Twin Arginine Translocase: Protein components and functional requirements 1.3.1 Protein components 1.3.1.1 cpTatC (TatC) The mature form of cpTatC comprises six transmembrane helical (TM) domains with an apparent molecular weight of ~30 kDa. The N- and C-termini are found on the stromal side of the thylakoid (Figure 1.1). We now know that cpTatC exists as a heterotrimer with Hcf106 and Tha4, which serves as the cpTAT receptor complex and is responsible for signal peptide binding. Signal peptide binding occurs on the stromal proximal portions of TM1 and TM2 of cpTatC (Ma and Cline, 2010). Mutational analysis of cpTatC has also shown that variations in TM5 directly impact overall complex assembly (Ma and Cline, 2010) by altering the site of Hcf106 (TatB) binding (Figure 1.2) (Habersetzer et al., 2017). Despite mutational analyses and protein interaction studies of cpTatC (TatC), concrete biophysical structural data had been lacking until recently. Currently, two solved structures of TatC purified from the thermophilic bacterium Aquifex aeolicus (Ramasamy et al., 2013; Rollauer et al., 2012) have guided recent mechanistic studies. Both structures of TatC were similar and presented an overall glove or cupped hand shape in the arrangement of the transmembrane helices where the concave face was suggested to serve as the nucleation site for the Tha4 (TatA) oligomer (Ramasamy et al., 2013; Rollauer et al., 2012). The transmembrane helices are arranged such that TM2, TM4, and TM5 make up the palm of the glove/hand while TM1, TM3, and TM6 bolster the convex portion of the proteins structure. Functional analysis of cpTatC and bacterial TatC has also provided insight into the requirement for specific residues in the receptor complex. For example, there is a conserved polar

Figure 1.1 Models of cpTatC, Hcf106, and Tha4 oriented in a thylakoid/plasma membrane generated in Chimera using sequence homology with identified TAT components (gray) as shown. Five iterations of cpTatC (blue) were modeled on the TatC crystal structure from A. aeolicus (Ramasamy et al., 2013) PDB: 4HTT by alignment of the 4HTT sequence with last 230 residues from P. sativum (Uniprot: Q94G17). Two iterations of Hcf106 (green) were modeled on the NMR structure of TatB from E. coli (Zhang et al., 2014b) (PDB: 2MI2) from the sequence alignment between 2MI2 and Hcf106 residues 87-261 (Uniprot: Q94G16); three additional Hcf106 structures were omitted due to extreme mismatch with TatB structural features. Five iterations of Tha4 (pink) were modeled on the solution NMR structure of TatAd from B. subtilis (Hu et al., 2010) PDB: 2L16 from the sequence alignment of 2L16 and the Tha4 residues 55-137 from P. sativum (Uniprot: Q9XH46). residue (glutamine in cpTatC/glutamate in TatC) that extends into the hydrophobic core of the membrane bilayer that when substituted in E. coli inhibits TAT transport but has a minimal effect on the thylakoid system (Holzapfel et al., 2007; Ma and Cline, 2013; Zoufaly et al., 2012). These observations taken in context with molecular dynamics simulations using the structures of the A. aeolicus TatC show that this residue is hydrated and is able to form a transient water path through the membrane which has led to the suggestion that the polar residue is able to connect the polar phases of the stroma (cytoplasm) and the lumen (periplasm) (Ramasamy et al., 2013; Rollauer et al., 2012).

1.3.1.2 Tha4 (TatA) and Hcf106 (TatB) Tha4 and Hcf106 are the additional protein components of the chloroplast twin arginine translocase. The bacterial homologs for these proteins are TatA and TatB, respectively. Tha4 (TatA) and Hcf106 (TatB) are quite similar in their overall predicted structure and sequence homology in both plants and bacteria (Alcock et al., 2016) and are essentially different branches of a larger TatA family proteins. Hcf106 (TatB) and Tha4 (TatA) are made up of four regions: a thylakoid lumen proximal N-terminal short TM domain, followed by a short hinge region, an amphipathic helix (APH), and an unstructured C- terminal tail (Figure 1.1). Sequence and phylogenetic analyses have suggested that the sequence divergence of these proteins arose from a gene duplication event (Yen et al., 2002). Although the structures of Tha4 (TatA) and Hcf106 (TatB) are nearly similar, the stochastic ratios of these proteins are quite different between the bacterial and plant systems. In the model organism Pisum sativum (garden pea), the ratio of Tha4:Hcf106:cpTatC is 21:4:1 (Celedon and Cline, 2012) while in E. coli the ratio of TatA:TatB:TatC is 75:2.5:1 (Berks et al., 2003; Jack et al., 2001; Sargent et al., 2001). The significance of such a difference is unclear.

The Hcf106 component of the cpTAT complex is found in two pools in the membrane: a heterotrimeric complex with cpTatC and Tha4 called the receptor complex and a separate free pool. Hcf106 (High chlorophyll fluorescence106) was the first protein component of the chloroplast TAT system to be identified (Settles et al., 1997). The mature form of Hcf106 has an apparent molecular weight of ~29 kDa. Recent biophysical characterization of bacterial TatB by nuclear magnetic resonance (NMR) spectroscopy largely confirmed predicted structures of the N-terminal half showing the presence of the hinge region such that the TM and APH form an L-shape (Zhang et al., 2014b). Several crosslinking studies and suppressor mutant analyses under non- transporting conditions have shown that the TM of E. coli TatB binds to a location on TM5 of TatC and in plants (Figure 1.2, Table 1.1) as was proposed previously (Blummel et al., 2015; Kneuper et al., 2012; Rollauer et al., 2012).

Figure 1.2 Structural modeling of Hcf106-cpTatC dimer under resting state. Model is viewed from the side (left) and top (Schleiff et al., 2002) of cpTatC. The Hcf106 TM is positioned in the vicinity of the cpTatC TM5 (left). The central APH of Hcf106 is positioned in the vicinity of the S1 and S2 of cpTatC. Manual docking simulation to generate an Hcf106-cpTatC docking model based on biochemically determined interactions. All modeling calculations were performed using the Rosetta 3 molecular modeling suite. An N-terminally truncated amino acid sequence (72-303) of cpTatC from P. sativum (GenBank accession no. AAK93948) was homology modeled onto the backbone structure of A. aeolicus TatC using PDB code 4B4A (Rollauer et al., 2012). A C-terminally truncated amino acid sequence (1-107) of mature Hcf106 from P. sativum (GenBank accession no. AAK93949) was homology modeled onto the structure of E. coli TatB using PDB code 2MI2 (Zhang et al., 2014b). The resulting models were used for docking exercises with Rosetta 3 (Kahraman et al., 2013) using known points of chemical crosslinking as constraints for docking. In brief, 1000 decoys were generated in low-resolution centroid mode and clustered in to four clusters. The lowest energy model of each cluster was then used for high resolution docking simulations with a request of 2500 decoys. More than 32,600 decoys were generated and were clustered using a custom script to pull the top 5% of structures by interface energy to generate 9 clusters. Three of the 5 lowest energy structures by interface score were found in a single cluster.

Table 1.1 Summary of crosslink interactions between cpTAT or bacterial TAT component proteins and precursor proteins. cpTAT Interaction cpTatC Hcf106 Tha4 Residue Precursor Crosslink Reference Residue Residue Residue Type cpTatC TM5 – A268C A12C Disulfide QM and CDS, personal Hcf106 TM communication A269C A12C, L13C, Disulfide QM and CDS, personal V14C, I15C communication V270C A12C, L13C, Disulfide QM and CDS, personal V14C communication cpTatC L2 – N203C F3C Disulfide (Aldridge et al., 2014) Tha4 TM cpTatC L3 – T275C F3C, P9C Disulfide (Aldridge et al., 2014) Tha4 TM cpTatC TM4 – L231C F3C, P9C Disulfide (Aldridge et al., 2014) Tha4 TM cpTatC TM5 – V270C F3C, P9C Disulfide (Aldridge et al., 2014) Tha4 TM Tha4 TM – F3C, F4C, V8C, V13C, Disulfide (Dabney-Smith et al., Tha4 TM A18C, L20C, V21C 2006) Tha4 APH – K26C, P28C, E29C, Disulfide (Dabney-Smith et al., Tha4 APH K39C, S40C, Q43C, 2006) K46C, E47C Tha4 C-tail – T59C, A65C, Q68C, Disulfide (Dabney-Smith et al., Tha4 C-tail T78C 2006) Tha4 Oligomers V8C P9C, A18C L20C, Disulfide (Dabney-Smith and Cline, A65C T78C 2009) cpTatC Stroma E73C, D78C (-G25C) Disulfide (Ma and Cline, 2013) loop 1 – tOE17 (-25C-20F) cpTatC – tOE17 (-V20) Tmd-Phe + (Gerard and Cline, 2006) UV Hcf106 – (-G10), Tmd-Phe + (Gerard and Cline, 2006) Signal peptide h (-G9) UV domain of tOE17 cpTatC – cpTatC L68C Disulfide (Ma and Cline, 2013) Stroma loop 1 cpTatC – cpTatC L126C Disulfide (Ma and Cline, 2013) Lumen loop 1

Table 1.1 (continued) Summary of crosslink interactions between cpTAT or bacterial TAT component proteins and precursor proteins. cpTAT Interaction cpTatC Hcf106 Tha4 Residue Precursor Crosslink Reference Residue Residue Residue Type cpTatC – cpTatC T164C Disulfide (Ma and Cline, 2013) Stroma loop 2 cpTatC – cpTatC V270C Disulfide (Ma and Cline, 2013) (TM5) cpTatC – cpTatC T275C Disulfide (Ma and Cline, 2013) Lumen loop 3 cpTatC – cpTatC G293C Disulfide (Ma and Cline, 2013) (TM6)

Bacterial TAT TatC TatB Residue TatA Residue Precursor Crosslink Reference Interaction Residue Residue Type TatC TM5 – M205C, L9C Disulfide (Habersetzer et al., 2017; TatB TM L206C, Kneuper et al., 2012) F213C TatC TM6 – S214C, L9C Disulfide (Habersetzer et al., 2017) TatB TM Q215C TatC TM5 – V212C, L9C Disulfide (Habersetzer et al., 2017) TatA TM F213C TatC – V202, L2016, Bpa + UV (Blummel et al., 2015) Tor(mCherry) T208 TatC – TatB F2, V3, I4, F6, Bpa + UV (Blummel et al., 2015) I10, I14 TatC – TatC G144C, Disulfide (Cleon et al., 2015) M205C TatC TM1 – L21C V18C Disulfide (Alcock et al., 2016) TatB TM TatB TM – L9C Disulfide (Kneuper et al., 2012) TatB TM TatB TM – F8, L16, V21 Tmd-Phe + (Alami et al., 2003) h domain of UV preSuf1 TatB TM – I4 Bpa + UV (Ulfig et al., 2017) h domain of TorA

For example, a docking simulation between models of Hcf106 and cpTatC constrained by experimentally determined crosslinks demonstrates the interactions between transmembrane regions (TM5 of cpTatC) and the APH of Hcf106 with stromal facing loops (Figure 1.2, Table 1.1). Additional evidence for this interaction was recently shown through co-sequence evolution and molecular modeling analysis (Alcock et al., 2016) where the polar residue E8 in the TM of E. coli TatB was modelled to form hydrogen bond interactions with three residues of TatC (M205, T208, and Q215) in what the authors termed the polar cluster (Alcock et al., 2016). These contacts were confirmed by cysteine substitution and disulfide crosslinking with the strongest interactions occurring between TatB L9 and TatC M205 and F213 (Habersetzer et al., 2017). Furthermore, topological studies of both Hcf106 and TatB have shown that the APH and C-tail regions are localized to the stromal/cytoplasmic membrane interface (Koch et al., 2012; Lee et al., 2006; Mori et al., 1999) suggesting interactions via the Hcf106 (TatB) APH and cytoplasmic loops of cpTatC (TatC). Finally, TatB-TatC crosslinks have been shown between the cytoplasmic loop 1 and the periplasmic loop 2 of TatC and the TM of TatB which are proposed to be crucial for the proper conformation of the signal peptide binding pocket (Blummel et al., 2015; Zoufaly et al., 2012) and (Figure 1.2, Table 1.1).

Tha4 is the smallest protein in the cpTAT complex with an apparent molecular weight of ~9 kDa. Tha4 has a structural organization similar to Hcf106 as seen in the published structures of bacterial TatA showing an L-shaped structure with a hinge region between the TM and APH (Hu et al., 2010; Rodriguez et al., 2013; Walther et al., 2010). In the same studies, TatA was shown to have shorter APH and C-tail regions than TatB (Hu et al., 2010; Rodriguez et al., 2013; Walther et al., 2010). Despite having very similar secondary structure motifs, Tha4 and TatA have noticeable differences in their primary sequences that impact TAT function in their respective membrane environments (Hauer et al., 2017). Tha4 (TatA) chimeras in which the TM of either the plant or bacterial protein was substituted for the opposing TM were each tested for active transport. Results showed that substituting the three amino acids in the TatA TM, S5, W7, and Q8, with the corresponding Tha4 residues, G, P, and E, was enough to restore transport with the thylakoidal TAT system (Hauer et al., 2017). Furthermore, comparing hydrophobicity of the TMs of Tha4 and TatA, showed that the TM of E. coli TatA was relatively more hydrophobic than Tha4 (Hauer et al., 2017). Therefore, as residues of the Tha4 TM were substituted by those from the TatA TM, the hydrophobicity of the Tha4 TM increased with a concomitant decrease in cpTAT transport efficiency in thylakoid membranes (Hauer et al., 2017). The overall implication of this work is that the hydrophobicity of the TM region has evolved separately to function in the given membrane lipid composition. Thylakoid and bacterial cytoplasmic membranes have stark differences in lipid composition and as a consequence, membrane fluidity (Cronan, 2003; Sprague and Staehelin, 1984).

In both plants and bacteria, Tha4 (TatA) exists mostly as a large pool of small homo- oligomers (Dabney-Smith and Cline, 2009; Leake et al., 2008). Specifically, in the thylakoid membrane during non-transporting conditions Tha4 exists as homo-tetrameric complexes that are maintained through interactions between the TMs of the individual

8 monomers (Dabney-Smith and Cline, 2009). Upon binding of a signal peptide of a precursor protein to the receptor complex and accumulation of the PMF, Tha4 polymerizes further into large complexes (Dabney-Smith and Cline, 2009; Mori and Cline, 2002). Studies of direct contacts between Tha4 (TatA) and cpTatC (TatC) showed that the N-terminus of Tha4 (TatA) contacts lumen loops (periplasmic loops) 2 and 3 of cpTatC (TatC), the TM helix contacts TM5 of cpTatC (TatC), and the APH interacts with the stromal/cytoplasmic regions of cpTatC (TatC) (Aldridge et al., 2014; Blummel et al., 2015; Zoufaly et al., 2012). In addition to these constitutive interactions between Tha4 and cpTatC, Tha4 was shown to form additional contacts under transport conditions with TM4 of cpTatC in a manner that suggests E10 of Tha4 is aligned with cpTatC Q234 (Aldridge et al., 2014) indicating a binding location for Tha4 in the active translocase (e.g., presence of RR signal peptide, a PMF, and the E10 of Tha4) (Aldridge et al., 2014). Further exploration of the TM contacts of TatA with TatC in E. coli was carried out by sequence co-evolution analysis and molecular modeling. The results of this study identified several proposed interactions that occur between TatA and TatC that are very similar to the interacting locations shown between TatB and TatC (Alcock et al., 2016).

1.3.2 Functional requirements 1.3.2.1 Precursor signal sequence Proteins that are transported across biological membranes must be targeted to the proper transport system and are often synthesized as higher molecular weight precursor proteins containing N-terminal, cleavable targeting sequences. The functional requirement of TAT-mediated transport is a specific signal peptide sequence for the initial binding with the translocase receptor complex. All known precursor substrates for the cpTAT pathway contain bipartite targeting sequences: the N-terminal portion serves

Figure 1.3 Thylakoid precursors containing the twin arginine motif show little sequence homology in the signal peptide. WebLogo sequence plot (Crooks et al., 2004) generated from 20 chloroplast TAT precursor signal peptides (Peltier et al., 2002). Underneath the plot is the general architecture of chloroplast signal peptides with an extend N-terminal region, the RR motif N domain, the primarily hydrophobic H domain, and the C-terminal domain leading to the protease cleavage site. Finally, the consensus of the cpTAT signal peptides are shown where x represents any residue, h represents hydrophobic residues, and u represents an uncharged residue.

9 as a stromal targeting factor and is cleaved after import of the precursor into chloroplasts, while the C-terminal portion serves as a lumen targeting signal peptide sequence to direct precursors to the thylakoid membrane. TAT signal peptides contain three distinct regions: an amino proximal n-domain containing the obligate twin arginine (RR) motif for which the pathway is named, a hydrophobic h-domain, and a polar c- domain (Figure 1.3) (Berks et al., 2014; Berks et al., 2000). While there is a broad consensus in the physicochemical properties between signal peptides of bacterial and plant TAT precursors, the amino acid sequences vary. The general chloroplast twin arginine motif consensus sequence is R-R-x-h-h/u where x represents any amino acid, h represents a hydrophobic amino acid, and u represents an uncharged amino acid (Figure 1.3) (Peltier et al., 2002). In the bacterial system, the twin arginine motif consensus sequence is S/T-R-R-x-F-L-K (Berks et al., 2000). Following the c-domain is the cleavage site for a signal processing peptidase that recognizes a general A-x-A motif (Cline, 2015). The n-domain has been found to be of variable length with examples of signal peptides that extended beyond the RR motif such as TorA in E. coli and the chloroplast precursor OE17 (PsbQ) (Berks et al., 2000; Peltier et al., 2002). Studies that have investigated the amino-terminal N domain in plants have shown that truncation of this extended domain improved in vitro transport efficiency for several cpTAT precursor proteins (Henry et al., 1997; Ma and Cline, 2000). The RR motif is critical for transport under normal conditions where it has been shown that a mutation to twin lysines prevents signal peptide interactions with the receptor complex in several different model organisms (Chaddock et al., 1995; Gerard and Cline, 2006; Henry et al., 1997; Niviere et al., 1992). Recent work has also highlighted the importance of the signal peptide h-domain as discussed below.

1.3.2.2 Energetics The transport of substrate proteins across biological membranes through the TAT system requires only the presence of the proton motive force (PMF) across the thylakoid or cytoplasmic membrane and does not depend upon the hydrolysis of nucleotide triphosphates (Cline et al., 1992; Mould and Robinson, 1991). Further investigations into the energetics of TAT transport were able to show that contributions from both ΔpH and Δψ (transmembrane electric potential) can facilitate active translocation in both plants and bacteria (Bageshwar and Musser, 2007; Braun et al., 2007). Experiments quantifying the PMF contribution to TAT transport showed that it facilitates transport using a fraction of the available gradient (i.e., the PMF is more than capable of supporting multiple rounds of transport without depleting the gradient, especially given that it would be most active during the daylight) and that the PMF requirement is different for each substrate (Alder and Theg, 2003; Aldridge et al., 2014; Bageshwar and Musser, 2007; Braun and Theg, 2008; Brock et al., 1995). It is unclear how this occurs, however. Two mechanisms have been proposed: one suggests that cpTAT proton utilization is due to a proton leak while another posits that a protein- proton motor facilitates transport (Alder and Theg, 2003; Berks, 2015). Experiments aimed at understanding the role of the PMF have led to the observation that initial substrate binding to the receptor complex is not PMF dependent (Ma and Cline, 2000). However, the substrate-triggered oligomerization of Tha4 (TatA) does require a PMF and it was shown that TatA oligomers dissipate following the removal of the PMF

(Alcock et al., 2013; Dabney-Smith et al., 2006; Mori and Cline, 2002; Rose et al., 2013). Other experiments, by contrast, provided evidence that these oligomers assemble in some transport-inactive bacterial strains in the absence of a PMF (Alcock et al., 2013; Leake et al., 2008). This has led to speculation over the role of the PMF in energizing TAT transport with a focus on protonation events of key residues in the TAT protein components, as the PMF has not been shown to be required during the transport step. One hypothesis is that the PMF is able to protonate the membrane embedded glutamate 10 of Tha4 such that hydrogen bonding with glutamine 234 in cpTatC is energetically more probable (Aldridge et al., 2014).

1.4 Molecular mechanism of transport 1.4.1 Receptor complex and precursor binding The receptor complex in the chloroplast system was originally shown to comprise a cpTatC (TatC) and Hcf106 (TatB) heterodimer in chloroplasts as well as E. coli via signal peptide binding studies and blue native polyacrylamide gel electrophoresis (Alami et al., 2003; Mori and Cline, 2001). Many of the initial binding studies of precursor in thylakoid used a modified substrate tOE17-20F containing a substitution of a phenylalanine at the -20 position, which increased the binding affinity (Celedon and Cline, 2012; Gerard and Cline, 2007). These studies revealed that there are two binding modes for cpTAT substrate signal sequences: a weak binding mode and a tightly bound, more deeply inserted mode (Celedon and Cline, 2012; Gerard and Cline, 2007). It was also noted that the deep binding mode was triggered by the accumulation of the PMF, which was also shown in E. coli (Bageshwar and Musser, 2007; Blummel et al., 2015; Gerard and Cline, 2007). Several studies including crosslinking, calorimetric binding, and genetic studies have been used to elucidate the precise binding location of the signal peptide in the receptor complex (Kreutzenbeck et al., 2007; Ma and Cline, 2013; Rollauer et al., 2012; Strauch and Georgiou, 2007; Zoufaly et al., 2012). In plants, it has been shown that the signal peptide primarily interacts with the N-terminal region and the first stromal loop of cpTatC (Gerard and Cline, 2007; Ma and Cline, 2013). Furthermore, the hydrophobic region (h-domain) of the signal peptide contacts Hcf106 (TatB) in an inverted hairpin shape after binding (Alami et al., 2003; Fincher et al., 1998; Frobel et al., 2012; Gerard and Cline, 2006; Hou et al., 2006). This configuration of the signal peptide places the h-domain in the membrane in close proximity to the TM of Hcf106 (TatB) which has been confirmed by photo-crosslinking experiments (Alami et al., 2003; Blummel et al., 2015; Gerard and Cline, 2006). More recent experiments in E. coli demonstrate that unfolding of the hairpin conformation of the bound signal peptide is critical for transport of the mature domain (Hamsanathan et al., 2017). Recent work also in E. coli has made the case for the role of the h-domain in precursor binding to the receptor complex (Huang and Palmer, 2017; Ulfig et al., 2017). These results showed that increases in the hydrophobicity of the h-region could improve transport efficiency of precursors containing lysine (or other) substitutions for the arginines (Huang and Palmer, 2017; Ulfig et al., 2017). These observations suggest that TAT signal peptides evolved to contain the twin arginine motif and an overall lower hydrophobicity in the h- region as a means to prevent targeting to the Sec translocon (Huang and Palmer, 2017; Ulfig et al., 2017). Additional precursor binding studies in plants demonstrated that each cpTatC in the complex was capable of binding substrate proteins in a non-cooperative

11 manner such that each can transport substrate independently (Celedon and Cline, 2012). These results support a model in which cpTAT operates more efficiently during the biogenesis of thylakoids and the chloroplast maturation process, which would have high demand for protein transport (Celedon and Cline, 2012). However, a cooperative manner of cpTatC (TatC) function cannot be ruled out, as evidenced by the ability of (cp)TAT to transport crosslinked precursors or heterodimeric precursors with two signal peptides, which suggest that at least two receptor complexes may function together to transport homo-oligomeric substrate proteins (James et al., 2013; Ma and Cline, 2010).

In addition to studies focused on the interactions between substrate precursors and the receptor complex, recent work has taken aim at further defining the contacts between cpTatC (TatC) and Hcf106 (TatB). In thylakoid, the TM of Hcf106 can be shown to interact with TM5 of cpTatC and to a lesser extent TM1/2 (QM and CDS personal communication and Table 1.1). Comparable results were demonstrated in E. coli both computationally by sequence co-evolution analysis and molecular modeling as well as experimentally (Alcock et al., 2016). Interactions between adjacent TatC monomers were also modeled where the periplasmic cap of one TatC monomer interacted with another TatC monomer matching crosslinking evidence found in several studies (Alcock et al., 2016; Blummel et al., 2015; Cleon et al., 2015; Ma and Cline, 2013; Zoufaly et al., 2012). These TatC-TatC contacts lead to a model of the translocase that functions as multimers of TatBC. The receptor complex has been shown to exist as a large oligomer of 3-4 heterodimers (Bolhuis et al., 2001; Cline and Mori, 2001; Tarry et al., 2009). The requirement of multiple receptor complexes functioning together as one was tested by introducing mutations that disrupted interactions between Hcf106 (TatB) and cpTatC (TatC), which prevented substrate binding and transport, while mutations in the RR binding location of cpTatC (TatC) still allowed translocase assembly (Buchanan et al., 2002; Ma and Cline, 2013). The presence of a large oligomeric receptor complex is also supported by data showing that disulfide crosslinking of individually bound substrate after binding to the receptor did not prevent TAT function (Aldridge et al., 2014; Ma and Cline, 2010; Ma and Cline, 2013).

1.4.2 Translocase assembly Studying the assembly of the TAT system prior to transport is an active area of research and recent studies in both thylakoid and bacteria have shed some light on the process. Translocase assembly begins with the binding of the precursor signal peptide to the receptor complex followed by PMF-dependent Tha4 (TatA) oligomerization and assembly with the precursor-bound receptor (Dabney-Smith and Cline, 2009; Dabney- Smith et al., 2006; Mori and Cline, 2002). Recent work has focused on gaining a better understanding of the molecular mechanisms of translocase assembly. Biochemical crosslinking assays, in both thylakoid and bacteria, showed that the binding of the signal peptide to the receptor complex altered interactions between cpTatC (TatC) and Hcf106 (TatB), which then promoted an interaction of Tha4 (TatA) with the core of the translocase (Aldridge et al., 2014; Blummel et al., 2015). In addition, studies in E. coli examined suppressor mutants in TatB that restored the transport of precursor in cells containing variants of TatC unable to bind the RR motif of the signal peptide or precursors lacking a signal peptide (Huang et al., 2017). Furthermore, it was shown that

12 the suppressor mutation, F13Y, in TatB was able to restore TatA oligomerization with the receptor complex without being able to biochemically detect the binding of substrate to the translocase (Huang et al., 2017), suggesting that this suppressor mutant causes a conformational change in the receptor complex that promotes TatA oligomerization at the receptor despite the lack of bound substrate (Huang et al., 2017). Multiple TatAC and TatBC interactions were also predicted based on sequence co-evolution analysis and molecular dynamics simulations (Alcock et al., 2016). TatBC interactions were predicted to occur between the TM of TatB and specific residues of the TatC TM5-loop- TM6 regions (Alcock et al., 2016) and experimentally confirmed in E. coli (Habersetzer et al., 2017). These data suggest a model of translocase assembly in which TatB is displaced from the polar cluster binding site upon precursor binding, which then allows TatA to enter the translocase to bind at the translocase active position in the cleft between TatC TM2 and TM4 (Alcock et al., 2016; Habersetzer et al., 2017). Binding of the TM of Tha4 to TM4 of cpTatC was shown in thylakoid (Aldridge et al., 2014); however, Tha4 switching or exchanging with Hcf106 has not been demonstrated in thylakoid. Stable interactions between the TM of Tha4 and TM5 of cpTatC are enhanced in the presence of signal peptide binding while still being present in its absence (Aldridge et al., 2014), which suggests the models of translocase assembly in plants and E. coli may have nuanced differences. In plants, the current evidence suggests a model of translocase assembly in which Hcf106 acts as gate that prevents the flow of Tha4 into the translocase. Upon signal peptide binding to the receptor complex, the receptor complex undergoes a conformational change that allows additional Tha4 to enter into and oligomerize in the translocase core; e.g., near TM4 of cpTatC (Figure 1.4). In E. coli, the evidence gathered supports a model where binding of the precursor displaces TatB from its binding site at the polar cluster which is followed by the binding of TatA in the same region; the swapping mechanism (Figure 1.4) (Alcock et al., 2016).

1.4.3 Models of translocation Two key criteria must be satisfied for the transport of precursors. First, the system must be able to accommodate folded domains of precursors with various cross-sectional diameters, and second, the membrane must not lose its ion impermeability because of transport to maintain the PMF. Several models have been suggested to describe how the translocase might adapt to the different sizes and shapes of transporting substrates while maintaining the ion impermeability of the membrane. One model suggests that the transport event occurs by inversion of the APH of Tha4 (TatA) into the pore such that the hydrophilic residues line the channel in a charge-zipper mechanism (Walther et al., 2013). However, this model seems highly unlikely and has been countered by studies in plant and bacterial systems that have shown no inversion of the Tha4 (TatA) APH occurs during transport (Alcock et al., 2017; Aldridge et al., 2012; Koch et al., 2012; Pal et al., 2013). Furthermore, recent in vivo analysis in E. coli demonstrated that several amino acid substitutions that would disrupt the charge zipper interactions failed to prevent TatA oligomerization demonstrating that when key residues implicated in salt bridge formation had their charges reversed, transport efficiency was not restored as would be expected in the charge-zipper mechanism (Alcock et al., 2017).

Figure 1.4 Possible models for Tha4 and Hcf106 packing with cpTatC in the presence of signal peptide or precursor. Visible cpTatC (blue) TMs are labeled 1–5, while TM6 is hidden behind TM5. A The constitutive binding positions of Tha4 (pink) and Hcf106 (green) TMs on TM5 of cpTatC prior to precursor binding. B No swapping of Hcf106 and Tha4 TMs, but rather an additional Tha4 positioned on cpTatC in an active thylakoid translocase. C Position swapping of Hcf106 and Tha4 TMs mapped onto cpTatC based on a bacterial model (Alcock et al., 2016; Habersetzer et al., 2017). A second model suggests that cpTatC (TatC) and Hcf106 (TatB) form the channel and that Tha4 adjusts the size of the opening to accommodate the precursor (Robinson and Bolhuis, 2004). A third model suggests that cpTatC (TatC) pulls precursor through bilayer regions that are destabilized by the amphipathic helices and atypically short transmembrane regions of a Tha4 (TatA) oligomer (Bruser and Sanders, 2003). In this third model, the presence of excess Tha4 (TatA) would cause localized disruption of the bilayer through a pinching or compressing of the bilayer. Support for this model was shown through experiments aimed at determining the stability of TatA oligomers in detergents and lipid bilayers. It remains to be determined if cpTatC (TatC) pulls precursor through the membrane, but aside from that aspect of this model, there is much that accounts for the experimental evidence. For example, molecular dynamics

14 simulations suggested that tetramers and nonamers of TatA were capable of disrupting lipid bilayers (Rodriguez et al., 2013). Further evidence for the strength of this model has been shown by topology studies in thylakoid membranes in which the APH of Tha4 was shown to be angled in a manner that relieved the hydrophobic mismatch under non-transporting conditions but then shifted to a more even partitioning at the stromal interfacial region during transport (Aldridge et al., 2012). Crosslinking analysis has also shown that the Tha4 TM binds the TM4 of cpTatC in manner that would localize the APH to the interfacial region during transport (Aldridge et al., 2014). Further evidence has been collected for the quantity of Tha4 (TatA) in an active translocase by fluorescence imaging and kinetic analyses where the total is ~25 monomers (Celedon and Cline, 2012; Leake et al., 2008). Additional quantification of Tha4 in an active translocase was determined by disulfide crosslinking studies in the presence or absence of precursor or signal peptide where the total monomeric count of Tha4 was 8- 16 (Dabney-Smith and Cline, 2009). Thus, the quantities of Tha4 (TatA) present in the active translocase can feasibly disorganize the lipid environment enough to facilitate precursor protein transport with the previously mentioned proton flow but without complete disruption of the membrane.

A fourth model suggests that Tha4 (TatA) oligomerizes to form the primary point of passage in the membrane that then facilitates transport (Gohlke et al., 2005; Mori and Cline, 2001; Sargent et al., 2006). This latter model describes a mechanism where cpTatC (TatC) and Hcf106 (TatB) receptors in complexes serve as a nucleation site for Tha4 (TatA) assembly and as a proteinaceous annulus to contain the assembled Tha4, placing the point of passage on the inside of the ring of receptor protein as drawn from data collected showing multiple, differently-sized Tha4 (TatA) oligomers that were isolated from transporting or non-transporting membranes using detergent extraction (Alcock et al., 2016; Dabney-Smith and Cline, 2009; Dabney-Smith et al., 2006; Gohlke et al., 2005; Sargent et al., 2006).

Another likely model is a combination of the second and fourth models where cpTatC- Hcf106 (TatC-TatB) heterodimers associate as higher ordered complexes that form a proteinaceous annulus around the oligomers of Tha4 (TatA) to restrict the area of disruption by the accumulated Tha4 (Figure 1.5) (Cline, 2015). This is an attractive model because it takes into account the necessity to maintain the ion impermeability of the membrane, while allowing a weakening or thinning of the membrane for translocation to occur. At rest the large (e.g., tetramer or higher of a cpTatC-Hcf106- Tha4 trimer) receptor complex is held together through the presence of Hcf106 and/or Tha4, which interact with TM1 on one cpTatC and TM5 of another to create a ring. Presence of the precursor and binding of its signal peptide disrupts the interaction between Hcf106 and TM1 of a cpTatC, disrupting the large ring and promoting the influx and interaction of Tha4 with TM4 of cpTatC. At this stage the PMF provides the neutralization of charge in the TM regions of Tha4 and Hcf106 to promote packing and oligomerization of Tha4 protomers. Tightly packed Tha4 in its transport active conformation (Aldridge et al., 2012) in the interior of the receptor ring would provide the membrane instability through thinning or weakening to allow precursor mature domain to pass. How the precursor would pass through the membrane at that point is still

15 unknown and is an area of active investigation. Regardless of the actual process of passage, it seems likely that the signal peptide remains attached and unfolds or unhinges as the precursor traverses the membrane (Gerard and Cline, 2006; Hamsanathan et al., 2017).

Figure 1.5 Proposed model of cpTAT function. A cpTAT in the receptor complex state where each cpTatC is bound to Hcf106 and Tha4. B Precursor proteins bearing the RR motif bind to the receptor complex. In this depiction of active cpTAT, two precursors are bound to opposing RR binding sites. C Upon stimulation of cpTAT by the PMF, Tha4 nucleates in the interior cavity of the translocase to facilitate transport through the thylakoid membrane. D The signal peptide is cleaved from the transported protein by a luminal peptidase and the new mature protein diffuses away to its location of function. The additional Tha4 also dissociates from the central cavity and the translocases returns to the receptor complex state, ready for a new round of precursor binding and transport.

1.5 Future perspectives Despite the new evidence that has clarified the interactions between the receptor complex proteins, Tha4 (TatA), the signal peptide, and the precursor protein, questions remain to be answered regarding the twin arginine translocase. One such question is what is the molecular mechanism by which Tha4 (TatA) oligomers facilitate protein

16 transport through the membrane bilayer? Answering this question may involve manipulation of the native lipid environment to determine the degree of membrane fluidity required for transport. Another avenue of study will be to examine the apparent difference between Hcf106 (TatB) function during the assembly of the translocase in chloroplasts and E. coli. Another area yet to be examined is the in vivo state of Tha4 (TatA) oligomers in an active translocase at physiological levels. A more specific avenue of study is to determine if a polar cluster mediates the interactions between cpTatC and Hcf106/Tha4 in a comparable manner to E. coli. Another avenue of research into the overall structure of the twin arginine translocase is to purify a stalled translocase described by Aldridge et al. (2014) and subject it to a battery of biophysical techniques. This is of course easier said than done given the transient nature and inherent heterogeneity of the TAT pathway.

1.6 Dissertation goals and specific aims The Twin Arginine Transport system is a deeply important translocase in plants as well as most bacteria and archaea as this introduction chapter has stressed. In planta, cpTAT plays an integral role in transporting fully folded protein cargo that are required for maintenance and repair of photosystem II by transporting such proteins as the 23 kDa and 17 kDa oxygen-evolving enhancer proteins, OE23 (PsbP) and OE17 (PsbQ) (Cline, 2015; Lu, 2016). In bacteria and archaea, the TAT system has been implicated in the virulence of certain pathogens (Berks, 2015; De Buck et al., 2008). Thus, it is of importance to better understand the precise details of (cp)TAT function to aid the advancement of improving crop resilience to photosystem stress as well as antibiotic targets in microbial pathogens. Despite the wealth of published literature, the exact molecular mechanistic details of how (cp)TAT functions remain elusive including but not limited to defining the transient interactions between individual component proteins and how (cp)TAT is able to transport fully folded proteins using only the PMF. In order to answer these lingering questions, we determined that purification and structural characterization of cpTAT component proteins is required. Previous work in the Dabney-Smith lab made use of solid-phase peptide synthesis to gain meaningful structural and lipid interaction data of both the transmembrane and amphipathic helices of Hcf106 (Zhang et al., 2013; Zhang et al., 2014a). However, these studies only characterized individual Hcf106 helices and not the full-length protein. So, we still need to complete these studies with full-length Hcf106. In addition to using biophysical techniques to study the cpTAT proteins, we can take advantage of the robust biochemical techniques used previously in our lab such as in vitro crosslinking and transport complementation assays in isolated chloroplast and thylakoid membranes (Aldridge et al., 2014; Aldridge et al., 2012; Dabney-Smith and Cline, 2009; Dabney- Smith et al., 2003; Dabney-Smith et al., 2006; Pal et al., 2013). Specifically, we sought to gain a better understanding of why Tha4 requires an obligate transmembrane glutamate for transport function in active cpTAT (Dabney-Smith et al., 2003). Substitution of this glutamate to alanine abolished transport (Dabney-Smith et al., 2003). Additionally, substitution of this glutamate with aspartate (maintaining the side chain chemistry) moderately recovered transport (Dabney-Smith et al., 2003). Currently, there is a gap in our collective understanding of cpTAT function because we don’t know the role of this glutamate in Tha4 function, specifically if it this residue is linked to Tha4

17 organization or Tha4 function relative to the other cpTAT components cpTatC and Hcf106. Thus, the work detailed in this dissertation is presented as two separate projects to satisfy the need for additional Hcf106 structural characterization as well as a detailed examination of Tha4 function and interaction relative to the transmembrane glutamate residue and substitutions therein.

Chapter 2 describes the development and optimization of methods to express and purify sufficient quantities of full-length, recombinant Hcf106 from E. coli. Detailed solid state NMR structural and lipid interaction data was previously collected for the transmembrane and amphipathic helices of Hcf106 (Zhang et al., 2013; Zhang et al., 2014a). In addition to these studies of solid-phase synthesized Hcf106 helices, other researchers solved the structure of a truncated variant of E. coli TatB in DPC micelles (Zhang et al., 2014b). These studies provide meaningful structural models of this protein, but we still lack a high-resolution structure of full-length Hcf106. In order to improve our chances of purifying this a full-length version of this protein, we generated a plasmid that when induced, led to expression of Hcf106 with an N-terminally linked maltose binding protein (MBP) (Lebendiker and Danieli, 2011). MBP was also linked to Hcf106 with the cleavage recognition sequence of tobacco etch virus (TEV) protease (Tropea et al., 2009). We purified MBP-Hcf106 and TEVp from E. coli using affinity chromatography (Lebendiker and Danieli, 2011; Tropea et al., 2009). We then optimized proteolytic cleavage reaction by testing several conditions and additives. In addition to reaction parameters, further purification of reaction products was carried out using multiple affinity and size exclusion chromatography techniques. Despite our best efforts, we were unable to completely isolate full-length Hcf106 from un-cleaved fusion protein but were able to remove free MBP and TEVp. Future directions for this project are discussed in detail.

Chapter 3 describes our work to better understand the role of the Tha4 transmembrane helix (TMH) glutamate in cpTAT function and Tha4 organization. Prior studies have shown that the Tha4 TMH glutamate is required for translocation of proteins by cpTAT (Dabney-Smith et al., 2003). When this residue is substituted with an aspartate, transport function is moderately restored in thylakoid membranes that have had endogenous Tha4 sequestered by αTha4 immunoglobulins (Dabney-Smith et al., 2003). Tha4 with alanine and glutamine substitutions for this glutamate do not complement loss of cpTAT function when endogenous Tha4 has been sequestered in thylakoid (Dabney-Smith et al., 2003). Furthermore, alanine substitutions of this glutamate were shown to alter Tha4 oligomer formation under transport conditions including functional precursor or signal peptide alone (Dabney-Smith and Cline, 2009). To expand upon these studies, we carried out complementation and oligomerization assays using glutamate, alanine, and aspartate substituted Tha4 variants. Complementation assays were used to determine how glutamate and aspartate spatial positioning in the TMH were able restore loss of transport function in Tha4 E10A variants. Glutamate substitutions closer to the N-terminus of Tha4 (closer to the lumen) were not tolerated in any position while aspartate substitutions in Tha4 E10A were weakly tolerated in positions on the same face of the TMH as the glutamate 10 position. Oligomer formation by Tha4 monomer interaction was also probed in glutamate, alanine, and aspartate

18 variants in three structural regions of the protein: the lumen proximate and stromal proximate helix regions and the C-terminal tail in the presence or absence of functional and non-functional precursor. Interactions between the TMH were enhanced in the presence of functional precursor in the glutamate, aspartate, and alanine variants. In the C-tail, Tha4 interactions were largely present regardless of the presence of precursor and PMF for each variant tested. Non-functional precursor and the addition of urea to functional precursor were both unable to promote increases in Tha4 oligomer formation. We were also able to correlate changes in the hydrophobic character of the transmembrane helix to oligomer formation and complementation efficiency using hydropathy calculations and alkaline extraction assays. Finally, the preliminary studies to examine interactions between the Tha4 E10/A/D variant TMH and cpTatC are discussed and future directions of this project are proposed.

1.7 References Alami, M., I. Luke, S. Deitermann, G. Eisner, H.G. Koch, J. Brunner, and M. Muller. 2003. Differential interactions between a twin-arginine signal peptide and its translocase in Escherichia coli. Mol Cell. 12:937-946. Alcock, F., M.A. Baker, N.P. Greene, T. Palmer, M.I. Wallace, and B.C. Berks. 2013. Live cell imaging shows reversible assembly of the TatA component of the twin- arginine protein transport system. Proc Natl Acad Sci U S A. 110:E3650-3659. Alcock, F., M.P. Damen, J. Levring, and B.C. Berks. 2017. In vivo experiments do not support the charge zipper model for Tat translocase assembly. Elife. 6. Alcock, F., P.J. Stansfeld, H. Basit, J. Habersetzer, M.A. Baker, T. Palmer, M.I. Wallace, and B.C. Berks. 2016. Assembling the Tat protein translocase. Elife. 5. Alder, N.N., and S.M. Theg. 2003. Energetics of protein transport across biological membranes. a study of the thylakoid DeltapH-dependent/cpTat pathway. Cell. 112:231-242. Aldridge, C., X. Ma, F. Gerard, and K. Cline. 2014. Substrate-gated docking of pore subunit Tha4 in the TatC cavity initiates Tat translocase assembly. J Cell Biol. 205:51-65. Aldridge, C., A. Storm, K. Cline, and C. Dabney-Smith. 2012. The chloroplast twin arginine transport (Tat) component, Tha4, undergoes conformational changes leading to Tat protein transport. J Biol Chem. 287:34752-34763. Bageshwar, U.K., and S.M. Musser. 2007. Two electrical potential-dependent steps are required for transport by the Escherichia coli Tat machinery. J Cell Biol. 179:87- 99. Berks, B.C. 2015. The twin-arginine protein translocation pathway. Annu Rev Biochem. 84:843-864. Berks, B.C., S.M. Lea, and P.J. Stansfeld. 2014. Structural biology of Tat protein transport. Curr Opin Struct Biol. 27:32-37. Berks, B.C., T. Palmer, and F. Sargent. 2003. The Tat protein translocation pathway and its role in microbial physiology. Adv Microb Physiol. 47:187-254. Berks, B.C., F. Sargent, and T. Palmer. 2000. The Tat protein export pathway. Mol Microbiol. 35:260-274. Blummel, A.S., L.A. Haag, E. Eimer, M. Muller, and J. Frobel. 2015. Initial assembly steps of a translocase for folded proteins. Nat Commun. 6:7234.

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Chapter 2: Purification of Maltose Binding Protein-Hcf106 Fusion for Structural Characterization

Christopher Paul New1, Aman Habtemichael2, Gwendolyn Thomas2, Katherine Siemen2, Carole Dabney-Smith1,2,*

1Graduate program in Cell, Molecular, and Structural Biology, Miami University, Oxford, Ohio 45056

2Department of Chemistry and Biochemistry Miami University, Oxford, Ohio 45056

*Corresponding author: Department of Chemistry and Biochemistry, Miami University, 651 East High St., Oxford, OH. Tel.: 513-529-8091; E-mail: [email protected]

Author contributions: CPN and CDS contributed to data analysis and writing the manuscript; CPN and AH contributed to molecular biology; CPN, AH, GT, and KS contributed to cell growth, overexpression of proteins, and purification trials.

2.1 Abstract The chloroplast Twin Arginine Transport (cpTAT) system is a vital protein transporting complex that functions in thylakoid membranes. It plays a major role in maintaining the photosynthetic systems of the chloroplast by transporting fully folded proteins of these photosystem complexes using only energy derived from the proton motive force. This pathway is also homologous to the TAT pathway in several species of bacteria and archaea which is responsible for secretion of fully folded proteins primarily related to virulence. While (cp)TAT systems share homologous component proteins and seemingly similar molecular mechanisms, differences exist between the two such as the molar ratio of their component proteins and the type and function of the transported substrate proteins. To better define the specific interactions between the three (cp)TAT component proteins, purification and biophysical characterization studies of the individual (cp)TAT proteins such as Hcf106 (TatB) have been pursued and published. Although prior investigations of this protein have been successful in purification and structural characterization, the studied protein was truncated for ease, ultimately lacking the unstructured C-tail domain. To purify full-length Hcf106, we generated an Hcf106 fusion protein linked to maltose binding protein by molecular biology and PCR techniques. Additionally, maltose binding protein was linked to Hcf106 (MBP-Hcf106) by the recognition sequence of tobacco etch virus protease (TEVp) for ease of affinity tag removal. MBP-Hcf106 and TEVp were expressed in E. coli and purified using various affinity chromatography techniques. We then optimized proteolysis reaction conditions by varying duration, protease concentration, temperature, and reaction additives. Purification trials of the proteolysis reaction products were tested by fast protein liquid chromatography using different exclusion sizes and detergents. A proteolysis reaction was also carried out using MBP-Hcf106 bound to amylose resin. The results of each purification trial are discussed despite mixed success in using amylose resin affinity chromatography. TEVp removal was attained using Ni-NTA magnetic beads. New directions and avenues for purification of full-length Hcf106 are also discussed.

2.2 Introduction Biological membranes contain transport systems that facilitate the movement of proteinaceous cargo across lipid bilayers. One of these protein transport systems is the Twin Arginine Transport (TAT) system, so named because of the conserved twin arginine (RR) motif in the signal peptides of TAT specific substrates that serve as the recognition sequence for the translocase, recently reviewed (Hamsanathan and Musser, 2018; New et al., 2018). TAT systems exist in bacterial and archaeal membranes as well as in the chloroplasts of plants (Hamsanathan and Musser, 2018; New et al., 2018). In plants, chloroplast (cp)TAT functions to transport proteins from stroma to the lumen of the thylakoid membrane. The cpTAT system is comprised of three protein components, cpTatC, Hcf106, and Tha4, that transport fully folded substrate proteins with only energy derived from the proton motive force (PMF) (Cline et al., 1992; Dabney-Smith et al., 2003; Ma and Cline, 2010; Settles et al., 1997). The bacterial and archaeal TAT systems have homologous component proteins: TatC (cpTatC), TatB (Hcf106), and TatA (Tha4) (Berks, 2015; Hamsanathan and Musser, 2018). In thylakoid membranes, cpTatC, Hcf106, and Tha4 work together as a heterotrimeric receptor complex responsible for recognizing and binding with the twin arginine motif signal

27 peptides of cpTAT substrate proteins prior to transport (Aldridge et al., 2014; Gerard and Cline, 2006; Gerard and Cline, 2007). The proposed structure of Hcf106 resembles that of Tha4; both proteins have an N-terminal transmembrane helix (TMH), a hinge region that links the TMH to an amphipathic helix (APH) followed by an unstructured, soluble C-terminal tail (Aldridge et al., 2012; Hu et al., 2010; Zhang et al., 2014b) (Figure 2.1A).

Figure 2.1 Cartoon diagrams of Hcf106 and the engineered MBP-TEVp recognition site-Hcf106 fusion protein. A Illustration of full-length Hcf106 in a thylakoid membrane showing the individual motif regions. B TEV protease recognition sequence highlighted in the MBP-Hcf106 fusion construct. TEV protease cleaves MBP from Hcf106 between the Gln-Gly peptide bond as indicated by the arrow.

Recent studies have contributed to structural models of Hcf106 and its prokaryotic homolog, TatB (Zhang et al., 2013; Zhang et al., 2014a; Zhang et al., 2014b). In these studies, recombinant protein purification or solid phase peptide synthesis were used to produce proteins for biophysical characterization by nuclear magnetic resonance (NMR) spectroscopy in detergent micelles and liposomes (Zhang et al., 2013; Zhang et al., 2014a; Zhang et al., 2014b). Although these experiments were able to determine the structure of Hcf106/TatB as well as their interactions with membrane lipids, full-length Hcf106/TatB was not used. For example, the TMH and APH of Hcf106 were separately synthesized and studied as individual helices (Zhang et al., 2013; Zhang et al., 2014a). As a result, the influence of the linked TMH-APH on the global structure of Hcf106 as well as its orientation and partitioning depth parameters in a membrane bilayer were not determined. In the prokaryotic system, the structure of truncated TatB was solved in detergent micelles which can alter the structure and topology of membrane proteins relative to a native membrane environment (Cross et al., 2011; Zhang et al., 2014b). In response to the lack of full-length structural data, the primary aim of this work was to develop an optimized expression and purification scheme to generate a sufficient quantity of homogenously pure, full-length Hcf106 for study by electron paramagnetic resonance (EPR) spectroscopy and other biophysical techniques such as precision force microscopy or solid-state nuclear magnetic resonance (ssNMR) (Matin et al.,

2017; Sahu and Lorigan, 2018; Sahu et al., 2013; Zhang et al., 2013; Zhang et al., 2014a).

Hcf106 is a thylakoid membrane protein with a single transmembrane helix (TMH) linked to an amphipathic helix (APH) ending in an unstructured, soluble carboxy- terminal tail (Figure 2.1A). Given the hydrophobic character of the amino acids that comprise the TMH and segments of the APH, Hcf106 should be relatively insoluble in aqueous buffers requiring solubilization with detergents or lipids (Figure 2.1A). In order to overcome this limitation and improve the probability of purification, we decided to generate a fusion protein linking Hcf106 to maltose binding protein (MBP). MBP has previously been used as a solubility enhancing protein affinity tag ultimately enabling purification by affinity chromatography methods using commercially available amylose resin (Lebendiker and Danieli, 2011). Therefore, MBP fusion to Hcf106 (MBP-Hcf106) would improve the likelihood of purifying a full-length version of this protein as it would increase the solubility of Hcf106 without the addition of detergent or lipid (Lebendiker and Danieli, 2011). An added benefit of using this affinity tag is that fusion of MBP to target proteins has been shown to improve expression of the target protein in bacterial hosts (Lebendiker and Danieli, 2011). By employing an MBP fusion tag to purify Hcf106, we then had to consider how to remove the affinity tag from the protein of interest following purification. We achieved these aims by using a commercial protein expression vector that generates MBP fusion proteins, pMAL-c5E (NE Biolabs), and used site-directed mutagenesis by PCR to overwrite the enterokinase recognition sequence with a tobacco etch virus protease recognition sequence (Lebendiker and Danieli, 2011; Tropea et al., 2009) (Figure 2.1B). The catalytic protease fragment of the tobacco etch virus, TEVpS219VHis6 (TEVp) was chosen for use instead of enterokinase because TEVp can be easily expressed in and purified from E. coli (Tropea et al., 2009). An added benefit of using TEVp is that it has been shown to be compatible for proteolysis in a wide variety of detergents and lipids containing buffers (Lundback et al., 2008; Vergis and Wiener, 2011).

In this study, we generated a pMAL-TEV plasmid and inserted the nucleotide sequence of mature, full-length Hcf106 into the vector by two-step megaprimer PCR (Bryksin and Matsumura, 2010). After successfully sequencing our new expression plasmid, separate induction screen trials were conducted to isolate individual clones for expression of TEVp and MBP-Hcf106. Expression and purification of TEVp was completed according to a previously reported protocol through the use of affinity and size exclusion chromatography (Tropea et al., 2009). Expression and purification of MBP-Hcf106 was completed using a modified version of the protocol published by Lebendiker and Danieli (2011) (Lebendiker and Danieli, 2011). Proteolysis optimization trials were conducted at various protein:protease ratios for different durations and at different temperatures. Multiple protease reaction additives were also tested. Further purification of Hcf106 from non-cleaved fusion protein, free MBP, and TEVp was attempted using fast protein liquid chromatography (FPLC). We also conducted an on-column proteolysis trial. Finally, we were able to remove TEVp from the reaction mixture following proteolysis through the use of magnetized Ni-NTA resin but purification of free Hcf106 was unsuccessful in subsequent FPLC trials. Although ultimately unsuccessful, the work presented here will

29 serve as a guide and starting point for future expression and purification of full-length Hcf106. We examine and discuss these future perspectives at length in this work.

2.3 Materials and Methods 2.3.1 Generation of pMAL-c5E with TEV protease recognition sequence between MBP and full-length Hcf106 sequences Site directed mutagenesis was used to replace the enterokinase cleavage sequence present in the commercial pMAL-c5E plasmid (NEBiolabs) with a tobacco etch protease cleavage sequence (residues: ENLYFQG) (Kunkel, 1985). Phusion polymerase (NEBiolabs) was used in all PCR site directed mutagenesis reactions. The overwrite and insert sequences were confirmed by BigDye Terminator Cycle sequencing (ThermoFisher) at the Center for Bioinformatics and Functional Genomics (CBFG) at Miami University (Oxford, OH). After confirming that the enterokinase recognition sequence was changed to the TEV protease recognition sequence, overlap extension PCR cloning was used to insert the mature sequence of Hcf106 (MTPSLAIA…) from a pGEM donor plasmid to the newly created pMAL-c5TEV recognition sequence plasmid (Bryksin and Matsumura, 2010; Geiser et al., 2001). Briefly, chimeric primers were designed that would overlap ~25 nucleotides downstream or upstream of the desired insertion site on the pMAL-c5TEV plasmid while also overlapping the first or last ~25 nucleotides of Hcf106 in a pGEM donor vector. The first round of PCR was used to generate a dual strand megaprimer that consisted of the Hcf106 sequence flanked on both ends by primer sequences that will bind to and insert Hcf106 into the pMAL-c5TEV plasmid. The megaprimer was purified from a 2% (w/v) agarose gel using a Wizard PCR kit (Promega) according to the manufacturer’s protocol. A second round of PCR using the purified Hcf106 megaprimer inserted the coding sequence of Hcf106 into the pMAL-c5TEV plasmid. After the reaction was completed, the restriction enzyme Dpn1 (NEBiolabs) was used according to the manufacturer’s protocol to degrade the methylated plasmid template. The completed MBP-Hcf106 plasmid was then isolated by colony PCR with Phusion polymerase (NEBiolabs) and NEB5α E. coli competent cells (NEBiolabs) according to the manufacturer’s instruction. After isolation, sequence of MBP-Hcf106 was confirmed as previously described at the Miami University CBFG.

2.3.2 MBP-Hcf106 protein expression Briefly, chemically competent BL21(DE3) codon plus E. coli (NEBiolabs) were transformed with the MBP-Hcf106 expression plasmid and grown overnight lysogeny broth (LB) agar plates with ampicillin (amp, 150 μg/mL) selectivity at 37°C. Individual colonies were isolated and screened for protein overexpression in LB media with amp selectivity (150 μg/mL) in 5 mL cultures. MBP-Hcf106 overexpression was induced by the addition of isopropyl β-D-1-thiogalactopyranoside (IPTG) to a final concentration of 1 mM. Initial expression was continued for 3 hr at 37°C in a shaking incubator. After confirming overexpression through SDS-PAGE and Coomassie blue staining, large scale overexpression was carried out in 2 L baffled flasks containing 1 L LB media with amp selectivity. Cultures were grown at 37°C until an OD600 nm of ~0.4-0.6 was reached. Large scale cultures were then induced for protein overexpression with IPTG at a final concentration of 1 mM. MBP-Hcf106 expression was completed following incubation for 3 hours at 37°C in a shaking incubator. Following the duration of protein expression, the

30 bacteria were collected in cell pellets by centrifugation at 3700 x g for 30 minutes and stored at -80°C until lysis.

2.3.3 Purification of MBP-Hcf106 fusion protein The purification protocol of MBP-Hcf106 was adapted from a previously published report (Lebendiker and Danieli, 2011). The cell paste collected from ~250 mL of induced culture was resuspended in ~30 mL of the appropriate buffer; MBP-Hcf106 in amylose resin binding buffer [50 mM Tris-HCl pH 7.4 with 200 mM NaCl, 1 mM EDTA, 1 mM dithiothreitol (DTT), and 1 mM phenylmethylsulfonyl fluoride (PMSF)] and TEVp in TEVp FPLC/reaction buffer [25 mM sodium phosphate (pH 7.5), 100 mM NaCl, 10% glycerol, 1 mM DTT, 1 mM PMSF)]. Lysis was executed using a French cell press at 20000 psi. After two passes, the cell lysate was clarified by centrifugation at ~40000 x g for 30 minutes at 4°C. The clarified supernatant was then passed through a gravity flow column packed with ~10 mL amylose resin bed (NEBiolabs). The eluate was saved, and the column was washed with 10 column volumes (CVs) of amylose resin binding buffer. Bound MBP-Hcf106 was then eluted in 2 mL aliquots in a total of 2 CVs of amylose resin elution buffer (amylose binding buffer or TEVp FPLC/reaction buffer + 10 mM maltose). Each aliquot was tested for protein content by SDS-PAGE with Coomassie blue staining. Fractions containing the fusion protein were pooled. The buffer containing MBP-Hcf106 was exchanged into amylose resin binding buffer by desalting through G- 25 sephadex resin (GE Healthcare) gravity flow column. The concentration of MBP- Hcf106 was determined using a BCA assay kit (Pierce) on a BioMate UV/Vis spectrophotometer according to manufacturer’s instruction.

2.3.4 TEVpS219VHis6 expression and purification The catalytic fragment of tobacco etch virus protease-S219V-His6 (TEVp) was prepared according to the previously published protocol (Tropea et al., 2009). pRK793 was a gift from David Waugh via Addgene (plasmid # 8827). Briefly, competent BL21(DE3) codon plus E. coli were transformed with the pRK793 vector and grown in 1 L LB cultures in 2 L baffled flasks in a 37°C shaking incubator until OD600 nm = ~0.6 was reached. Induction of protein overexpression was initiated by addition of IPTG to a final concentration of 1 mM. Overexpression was carried out for 4 hours at 30°C. Post induction, the cells were pelleted by centrifugation at 3700 x g for 30 min into 250 mL aliquots followed by freezing at -80°C. The cell paste of one 250 mL aliquot was then resuspended in cold lysis/IMAC equilibration buffer (50 mM sodium phosphate, pH 8.0, 200 mM NaCl, 10% glycerol, 25 mM imidazole) and lysed by 2 passes through a French cell press at 20000 psi. Cell lysates were clarified by centrifugation at ~40000 x g at 4°C for 30 minutes. Following clarification, the supernatant (soluble) fraction was decanted for further purification by Ni-affinity chromatography. The supernatant was added to 10 mL of packed Ni-NTA resin (Qiagen) in a column and capped. The resin was incubated with lysate in the capped column on a tilt table in a 4°C cold room for 1 hr. First, the flow- through lysate was eluted and collected. The column was then washed with 10 CVs of lysis/equilibration buffer and the eluted wash buffer was collected for SDS-PAGE analysis. After the washing step, TEV protease was eluted from the Ni-NTA resin with 2 CVs of elution buffer in 2 mL aliquots (50 mM sodium phosphate, pH 8.0, 200 mM NaCl, 10% glycerol, 250 mM imidazole). Following the elution process, the eluate aliquot

31 fractions were analyzed by SDS-PAGE; aliquots containing purified TEVp were pooled. TEVp was then concentrated and the buffer was exchanged into gel filtration buffer (25 mM sodium phosphate (pH 7.5), 100 mM NaCl, 10% glycerol) by 3 kDa MWCO spin concentrators (GE Healthcare) by centrifugation according to the manufacturer’s instruction. The volume of TEVp was reduced to ~1 mL and was further purified by filtration through a HiLoad 16/600 Superdex 75 pg prepacked column with an ATKA FPLC system (GE Healthcare). Aliquots corresponding to increases in absorbance at 280 nm were collected and analyzed by SDS-PAGE and Coomassie blue staining. The fractions containing purified TEVp were then concentrated by spin concentrators as before and stored in 100 μL fractions at -80°C until use.

2.3.5 TEV protease cleavage of MBP-Hcf106 Purified MBP-Hcf106 cleavage trials were carried out using purified TEVp. Cleavage reactions were screened for compatibility with urea (Fisher Scientific), 3-((3- cholamidopropyl) dimethylammonio)-1-propanesulfonate (CHAPS, ACROS Organics), nonaethylene glycol monododecyl ether (C12E9, Sigma-Aldrich), Anzergent 3-14 (AZ314, Anatrace), or FOS choline 12 (DPC, Anatrace) in cleavage reaction buffer (50 mM Tris-Cl pH 8, 200 mM NaCl). Reactions were carried out at either 4°C or 25°C at various time durations. Aliquots were drawn at various time points and were examined for completion of proteolysis by SDS-PAGE and Coomassie blue staining.

2.3.6 Fast protein liquid chromatography purification trials TEVp cleavage reactions of MBP-Hcf106 were carried out as in 2.3.5 followed by buffer exchanged by dialysis using 3000 MWCO SnakeSkin tubing (Thermo Scientific) into gel filtration buffer with added detergent (50 mM Tris-HCl pH 7.5, 1 mM DTT, 0.1% CHAPS or 0.1% C12E9). After buffer exchange, the reaction mixture was separated by either a Superdex 200 Increase 10/300 GL or HiLoad 16/600 Superdex 75 pg column with an ATKA Pure system (GE Healthcare). The collected fractions were then analyzed for protein content by SDS-PAGE and Coomassie blue staining. Urea additive buffer based FPLC experiments were carried out using a HiLoad 16/600 Superdex 200 pg prepacked column with an ATKA Pure system (GE Healthcare) with either 300 mM or 4 M urea added to the phosphate based reaction buffer (25 mM sodium phosphate (pH 7.5), 100 mM NaCl, 10% glycerol).

2.3.7 Amylose resin batchwise TEVp cleavage reactions of bound MBP-Hcf106 MBP-Hcf106 was purified from BL21 E. coli using the lysis protocol presented previously. Initially, 5 mL of clarified lysate containing MBP-Hcf106 was passed through 2 mL of packed amylose resin (NEBiolabs) in a gravity flow column (Bio-Rad). The column was then washed with 10 CV of a modified amylose resin binding buffer (50 mM Tris pH 7.4, 1 mM EDTA, 1 mM DTT, 0.075 mM CHAPS). The amount of TEVp added to the reaction was based on the average binding capacity of amylose resin determined by the NEBiolabs for a MBP5*-paramyosin ΔSal fusion, ~4 mg fusion protein/mL of resin. The cleavage reaction of MBP-Hcf106 was initiated upon the addition of TEVp to the buffer head of the amylose resin to yield a 1:10 ratio of TEVp to hypothetically calculated concentration of bound MBP-Hcf106 (~8 mg for 2 mL of packed resin). The gravity column was then capped and mixed on a tilt table for 16 hours in a 4°C room.

After the cleavage reaction incubation period, the column was set upright and allowed to settle. Once settled, the eluate from the cleavage reaction was collected and concentrated by spin concentrators. Samples for SDS-PAGE and Coomassie blue staining were collected from the lysate, the post binding flow-through eluate, the buffer exchange eluate, and the post cleavage eluate.

2.3.8 Removal of TEV protease by Ni-NTA magnetic beads His Mag Sepharose Ni (GE Healthcare) magnetic beads were used to remove TEVp from a 50:1 (mg/mL:mg/mL) MBP-Hcf106:TEVp reaction according to the manufacturer’s instructions. Briefly, the beads from ~600 μL of suspended magnetic bead slurry were pulled to the side of a snap cap tube using a magnet lined rack. The supernatant was removed, and the beads were equilibrated with TEV protease reaction buffer including 300 mM urea and 10 mM imidazole. After equilibration, ~750 μL of a 2 day long, 25°C, 50:1 MBP-Hcf106:TEVp (mg/mL:mg/mL) reaction was added to the beads and incubated at 25°C for 45 mins rotating end over end. Following the incubation, the magnetic beads were sequestered to the side of the tube by the magnet rack and the supernatant sample was collected. The beads were then washed with equilibration buffer and then bound proteins were eluted from the beads after incubation with TEVp reaction buffer containing 300 mM urea and 500 mM imidazole. This binding, wash, and elution process was then repeated once. Collected samples were analyzed by SDS-PAGE.

2.3.9 Western blot analysis of SDS-PAGE resolved proteins Western blot analysis of SDS-PAGE resolved proteins was initially carried out by wet transfer of proteins to nitrocellulose blotting membranes (Amersham). After transferring resolved proteins results, nitrocellulose membranes were blocked with 5% non-fat dry milk dissolved in 1x Tris-buffered saline with 0.1% Tween 20 (TBS-T, 50 mM Tris-HCl, pH 7.6, 150 mM NaCl, 0.1% Tween 20). Primary antibody treatment with a 1:5000 dilution of serum from rabbit containing anti-Hcf106 antibodies in 1x TBS-T, 5% (m/v) powdered milk. Primary antibody decorated membranes were washed with 1x TBS-T and incubated with the secondary antibody, goat-antirabbit conjugated to horse radish peroxidase (Bio-Rad) at 1:20000 titer in 5% milk in 1x TBS (lacking Tween 20). Chemiluminescence was generated by incubation of primary and secondary antibody decorated membranes with Clarity Western ECL Substrate (Bio-Rad). Image was captured at 15 min exposure in UVP multi-user imaging cabinet and camera.

2.4 Results 2.4.1 TEV protease cleavage recognition sequence and the mature Hcf106 sequence from garden pea were cloned into a pMAL-c5E vector We used two step PCR to clone the nucleotide sequence of mature Hcf106 from a pGEM plasmid into the destination pMAL-c5E with a TEVp recognition sequence vector. In order to do so, chimeric primer pairs were generated such that half of the sequence of the primer binds the 5’ and 3’ regions of the top and bottom strands of the Hcf106 sequence while the other half of the primer binds to the desired insertion site in the pMAL-c5E plasmid (Bryksin and Matsumura, 2010; Geiser et al., 2001). First, the optimal annealing temperature of the megaprimer was determined by gradient PCR.

Figure 2.2 Insertion into and confirmation of mature Hcf106 sequence in pMAL-TEV recognition vector by various PCR techniques. A Annealing temperature gradient PCR results for Hcf106 insert megaprimer synthesis. An annealing temperature gradient from 45-55°C was used, lanes 1-8. Lane 9 is a 100 bp DNA step ladder (NEBiolabs). Lane 10 is pGEM-Hcf106 template only. B Scaled-up PCR reaction to generate the Hcf106 megaprimer. Lane 1 is 100 bp DNA step ladder (NEBiolabs). Lanes 2-3 show Dpn1 (NEBiolabs) and non-Dpn1 treated megaprimer samples. Lane 4 shows higher concentration pGEM-Hcf106 template. C Megaprimer PCR with annealing temperature gradient. Lanes 1-2 show the megaprimer and the pGEM-Hcf106 template. Lane 3 is a 1 kbp step ladder (NEBiolabs). Lanes 4-11 show the annealing temperature gradient from 50-65°C. D Colony PCR amplifying the Hcf106 insert in the newly purified MBP-Hcf106. Lanes 1-7 are individual colonies, showing a positive reaction in lane 6. The megaprimer was successfully synthesized as shown by agarose gel electrophoresis (Figure 2.2A). The complete megaprimer is 585 bp which corresponds to its migration between the bands corresponding to 500 and 600 bp (Figure 2.2A). Following optimization, the reaction was scaled up to a 50 μL reaction volume for use in the second round of PCR using 55°C as the annealing temperature. The Dpn1 digestion was successful in removing the remaining template DNA following the megaprimer synthesis PCR reaction (Figure 2.2B). After purification of the megaprimer, another PCR experiment was carried out to clone the sequence of Hcf106 into the pMAL-c5TEV destination vector. Annealing temperature gradient PCR was used to optimize the annealing temperature of megaprimer binding to pMAL-c5TEV. The results of the PCR gradient showed that the productive megaprimer PCR annealing temperature was in the range from 51.4-62.1°C with the optimal being at 59.4°C (Figure 2.2C). Following the optimization of the megaprimer annealing temperature, the reaction was scaled up followed by a colony PCR assay to ensure that the colony of choice contained the proper pMAL-Hcf106 vector (Figure 2.2D). The result of the colony PCR was that #6

34 contained the sequence for mature Hcf106 (Figure 2.2D). After the confirmation by of insertion of Hcf106 by colony PCR, the MBP-Hcf106 plasmid was sequenced for accuracy.

2.4.2 Recombinant TEV protease was purified from BL21(DE3) codon plus E. coli After successfully designing and synthesizing a vector to produce the MBP-Hcf106 fusion protein linked by a TEV protease recognition sequence, we began protein purification trials. We first set out to purify recombinant TEVp that would be used to proteolytically cleave the MBP affinity tag from Hcf106. The variant of TEVp used in this study is a truncated version of the full-length protease containing only the catalytic domain with an S219V substitution (David Waugh, Addgene plasmid #8827). This TEVp variant is expressed as a maltose binding protein fusion to the S219V substituted catalytic domain with a C-terminal hexahistidine tag. After the induction of

Figure 2.3 Expression and purification of TEVp. A Screening for TEVp expression in BL21 E. coli. Lane 1, EZ-Run prestained protein ladder (Thermo Fisher). Lanes 2-9 are samples from four transformed colonies pre-induction by IPTG and post IPTG and 3-hour expression. B Purification of TEVp by Ni-NTA resin. Lanes 1-6 are samples from elution fractions from TEVp bound Ni-NTA. C Chromatogram showing absorbance at 280 nm of TEVp polishing FPLC experiment. D SDS-PAGE analysis of pooled TEVp polishing FPLC elution fractions, lanes 1-3 correspond to fractions 21-22 (2.3C, peak 1), 25-28 (2.3C, peak 2), and 32-37 (2.3C, peak 3), respectively.

35 overexpression, the MBP-TEVp-His6 fusion accumulates as a soluble protein which then undergoes autolysis (Tropea et al., 2009). This process removes the MBP tag from the catalytic domain which can then be recovered by immobilized metal ion affinity chromatography (IMAC) using Ni2+ bound NTA resin (Qiagen) (Tropea et al., 2009).

An initial induction screen of four BL21 E. coli colonies each transformed with the pRK793 plasmid (David Waugh, Addgene #8827) was carried out to find a suitable starting colony for larger scale TEVp purification (Figure 2.3A). Each colony was capable of overexpressing S219V TEVp so colony #4 was chosen arbitrarily for a scaled-up purification process (Figure 2.3A). After overexpression of TEVp according the published protocol, the cells were harvested and lysed followed by lysate clarification (Tropea et al., 2009). The clarified lysates from two separate TEVp cultures were incubated with Ni-IMAC resin in separate gravity flow columns. Purified TEVp was eluted from the columns in several fractions that were analyzed by SDS-PAGE (Figure 2.3B). Following purification and analysis of the TEVp fractions collected during IMAC purification, the fractions were pooled according to starting culture (1 or 2) and were concentrated by centrifugal spin concertation units. The concentrated TEVp samples were then purified by FPLC and analyzed by SDS-PAGE (Figure 2.3C-D). FPLC fractions 32-37 were then pooled and concentrated following the completion of SDS- PAGE (Figure 2.3D). The final TEVp concentration of 0.25 mg/mL was determined by NanoDrop spectrophotometer (Thermo Fisher).

Figure 2.4 Expression and purification of MBP-Hcf106. A Screening for MBP-Hcf106 expression in BL21(DE3) codon plus E. coli. Lane 1, EZ-Run pre-stained protein ladder (ThermoFisher). Lanes 2-9 are samples from four transformed colonies before induction by IPTG and 3 hours of expression. B Comparison of soluble and insoluble fractions of MBP-Hcf106 after cell lysis. Lanes 1-2 show the pre- and post IPTG induction lysate fractions. Lanes 3-5 are 1x, 0.1x, and 0.01x dilutions of the supernatant (soluble) fraction following lysate clarification. Lanes 6-8 are 1x, 0.1x, and 0.01x dilutions of the dissolved insoluble pellet fraction. C Purification of MBP-Hcf106 by amylose resin chromatography. Lanes 1-2 are the flow-through and wash fractions from the column. Lanes 3-8 are the six elution fractions of MBP-Hcf106 after the addition of maltose.

2.4.3 Isolation and purification of recombinant MBP-Hcf106 from BL21(DE3) codon plus E. coli The first step in the expression and purification of MBP-Hcf106 from E. coli was an induction screen. The induction screen allowed us to analyze protein production of four MBP-Hcf106 plasmid transformed BL21 E. coli colonies upon induction by IPTG (Figure 2.4A). Colony one produced a product that was not the correct size for the ~75 kDa MBP-Hcf106 fusion while colonies 2-4 expressed a fusion protein of the correct size (Figure 2.4A). Colony number 2 was chosen for large scale MBP-Hcf106 production. After the induction of protein expression, cell harvest, and mechanical lysis by French pressure cell press (Thermo Fisher), the recovered lysate was clarified by centrifugation and the resulting supernatant and pellet were tested for protein content by SDS-PAGE. Once again, the induction of MBP-Hcf106 expression was confirmed (Figure 2.4B, lanes 1-2). The results of SDS-PAGE also showed that the MBP-Hcf106 is primarily located in the soluble fraction post lysis and not the pellet of insoluble material (Figure 2.4B, lanes 3-5). A large amount of free MBP was also present in the soluble and insoluble fractions indicating lysis by endogenous protease (Figure 2.4B, lanes 3-8). To prevent this lysis in the future, additional isolation and purification attempts included a protease inhibitor in the buffer. Phenylmethanesulfonyl fluoride (PMSF) was chosen as it will not inhibit TEVp but will inhibit the endogenous E. coli serine proteases present in the cell lysate (Mohanty et al., 2003; Vergis and Wiener, 2011). TEVp is a trypsin-like cysteine protease and is only strongly inhibited by iodoacetamide and specific detergents (Phan et al., 2002). After determining which protease inhibitor to use, another large-scale expression culture of MBP-Hcf106 was completed. Following the lysis and clarification of the lysate, the supernatant was passed over a packed gravity amylose resin column (NEBiolabs) to bind the MBP tagged fusion protein. The lysate contained a multitude of bacterial proteins in addition to MBP-Hcf106 (Figure 2.4C, lane 1) which were washed from the resin by 10 column volumes of binding buffer (Figure 2.4C, lane 2). After a thorough series of wash steps, MBP-Hcf106 was eluted from the column by the addition of maltose to the binding buffer (elution buffer = amylose binding buffer with 10 mM maltose added) into six fractions that were sampled on SDS-PAGE (Figure 2.4C, lanes 3-8). After we confirmed the presence of MBP-Hcf106 in these fractions, the concentration of protein in each was determined by a bicinchoninic acid (BCA) protein concentration assay (Pierce). After generating a standard curve for bovine serum albumin concentrations ranging from 0.1 mg/mL to 0.5 mg/mL, the linear regression of the data points yielded the line y = 3.076x + 0.03242. The 1x and 0.1x MBP-Hcf106 fractions fell outside of the high end of the calibration curve but the 0.01x concentrations were within the curve and were used to calculate the final protein concentration of each eluate fraction. The final fraction concentrations are presented in Table 2.1.

Table 2.1 BCA assay determination of MBP-Hcf106 concentration following elution from amylose resin column. Calibration curve: y = 3.076x + 0.03242 Amylose Resin Absorbance 0.01x concentration 1x concentration Eluate Fraction # 562 nm (mg/mL) (mg/mL) 1 0.17576 0.0466 4.66 2 0.17022 0.0448 4.48 3 0.31695 0.0925 9.25 4 0.12839 0.0312 3.12 5 0.10286 0.0229 2.29 6 0.07671 0.0144 1.44

2.4.4 Testing and optimization of TEVp proteolysis of MBP-Hcf106 Prior to any proteolysis reactions, TEVp and MBP-Hcf106 were both exchanged into the initial cleavage buffer listed in Tropea et al. (50 mM Tris-HCl pH 8.0, 0.5 mM EDTA, 1 mM DTT) by 3 kDa MWCO centrifugal concentrators (GE Healthcare) (Tropea et al., 2009). Once exchanged into reaction buffer, several parameters of the proteolysis reaction were tested. The first cleavage reaction condition tested was recommended in the protocol of Tropea et al. (2009) that suggested TEVp reactions could be carried out in (mg/mL):(mg/mL) ratios between 100:1 to 5:1 substrate:protease (Tropea et al., 2009). In addition to the ratio of substrate:protease, the suggested cleavage duration and temperature were also compared (Figure 2.5). For the reactions carried out at 4°C, five substrate:protease ratios were tested: 100:1, 50:1, 25:1, 10:1, and 5:1 (Figure 2.5A, lanes 1-5). The 4°C reactions were carried out for 16 hours (Figure 2.5A, lanes 1- 5) as this duration was recommended for initial TEVp cleavage reactions at this temperature (Tropea et al., 2009). In addition to those carried out at 4°C, cleavage reactions were conducted at 25°C for a duration of 2 and 4 hours at substrate:protease ratios of 10:1 and 25:1 (Figure 2.5A, lanes 6-9). We were able to monitor the progress and completeness of TEVp proteolysis by appearance of two bands on polyacrylamide gels near the 43 kDa and the 26 kDa protein markers corresponding to free maltose binding protein (42.5 kDa) and free Hcf106 (migrates on 12.5% SDS-PAGE at ~27 kDa) (Figure 2.5A). The 4°C reactions show an increase in the completeness of the proteolysis reaction as the ratio of protease:substrate was increased as expected (Figure 2.5A). The reactions that were carried out 25°C had slightly different degrees of completion. The 10:1 cleavage reaction showed minimal differences between the completeness of the reaction between the 2- and 4-hour time points (Figure 2.5A). However, in the 25:1 reaction, there was a marked increase in cleavage of the fusion protein between the 2- and 4-hour time points (Figure 2.5A). After determining that cleavage of MBP-Hcf106 was incomplete after ~24 hr at 4°C, the reaction duration was increased to 48 hr while also testing various substrate:protease ratios (Figure 2.5B). The results of the 48 hr cleavage reactions were similar to 24 hr reactions as the 10:1 reactions were more complete relative to the other ratios tested (Figure 2.5B).

Despite these increases in reaction duration, the cleavage of MBP-Hcf106 by TEVp was hardly reaching completion as shown by large amount of remaining fusion protein under all tested conditions (Figure 2.5A-B). Thus, we needed to improve the efficiency of the reaction. We decided to test improving the solubility of free Hcf106 after being separated from MBP by adding detergent to the reaction. We also hypothesized that a small addition of denaturant to the reaction may improve the ability of TEVp to access the cleavage recognition sequence. We tested the addition of urea or CHAPS to the reaction mixture while maintaining similar duration and temperature parameters. We tested 300 mM urea and 15 mM CHAPS in reactions with a 10:1 ratio of substrate:protease (Figure 2.5C). The addition of urea to the reaction shifted the cleavage reaction towards completion, especially as the concentration of protease was increased (Figure 2.5C, compare lanes 7 and 9 with Figure 2.5C, lanes 5-8). The addition of 15 mM CHAPS also appeared to shift the reaction towards completion as the concentration of protease was increased (Figure 2.5C, compare lanes 7 and 9 with Figure 2.5C, lanes 10-13). However, each additive failed to push the reaction to completion (Figure 2.5C).

Figure 2.5 Optimization of MBP-Hcf106 cleavage by TEVp. A SDS-PAGE of initial trials of MBP removal from Hcf106 by TEVp. Lanes 1-5, 16-hour proteolysis reaction at 4°C with substrate:protease ratios 100:1, 50:1, 25:1, 10:1, & 5:1. Lane 6, EZ run protein ladder (Thermo Fisher). Lanes 6-7, 10:1 ratio at 25°C for 2 hr and 4 hr. Lanes 8-9, 25:1 ratio at 25°C for 2 hr and 4 hr. B Extended duration TEVp reactions with MBP-Hcf106. Lane 1 is MBP-Hcf106. Lane 2 is the TEVp and MBP-Hcf106 reaction mixture at time = 0. Lanes 3-6 are samples from 10:1, 25:1, 50:1, 100:1 reactions after 48 hr at 4°C. C SDS-PAGE of urea or CHAPS addition to proteolysis reaction. Lane 1, MBP-Hcf106. Lanes 2-3, TEVp at 25:1 and 10:1 ratios. Lanes 5-8, addition of 300 mM urea to 2 hr, 25°C reactions at 100:1, 50:1, 25:1, 10:1. Lanes 10-13, addition of 15 mM CHAPS to 2 hr, 25°C reactions at 100:1, 50:1, 25:1, 10:1. 2.4.5 FPLC purification tests with post TEV protease-MBP-HCF106 reaction products Having pushed the cleavage reaction further towards completion with the addition of 15 mM CHAPS, we carried out separation of the reaction products by fast protein liquid chromatography (FPLC). Separation of a 10:1 substrate:protease reaction was

Figure 2.6 FPLC purification trials of TEVp and MBP-Hcf106 reactions in CHAPS or C12E9 detergents. A Chromatogram and SDS- PAGE analysis of MBP-Hcf106 and TEVp (10:1) proteolysis reaction in 15 mM CHAPS buffer on an S200 Superdex Increase 10/300 GL column (GE Healthcare). Lane 1, 10:1 reaction after 2-hour at 25°C. Lanes 2-9 are acetone-precipitated samples from FPLC elution fractions 8-10 (peak 1), 14-16 (peak 2), 19 (peak 3), and 21 (peak 4). Lane 10 is the eluate from an SDS buffer wash of an in-line 1 mL HisTrap column (GE Healthcare) that preceded the size exclusion column. B Chromatogram and SDS-PAGE analysis of MBP-Hcf106 and TEVp (10:1) proteolysis reaction in 0.1% C12E9 buffer on a HiLoad 16/600 Superdex 75 pg column (GE Healthcare). Lane 1, diluted TEVp. Lane 2, MBP- Hcf106. Lanes 3 is 10:1 reaction, 2-hour at 25°C. Lanes 4-10 are acetone-precipitated protein samples from FPLC fractions 18-19 (peak 1), 23-25 (peak 2), and 30-31 (peak 3). C Chromatogram and SDS-PAGE analysis of MBP-Hcf106 and TEVp of a 10:1 reaction in 15 mM CHAPS buffer using a HiLoad 16/600 Superdex 75 pg column (GE Healthcare). Lane 1, 10:1 reaction after 2 hr at 25°C. Lanes 2-12 are acetone precipitations corresponding to elution fractions 22-25 (peak 1), 29-31 (peak 2), 55 (peak 3), and 66-68 (peak 4).

40 attempted using either a Superdex 200 Increase or HiLoad 16/600 Superdex 75 pg (GE Healthcare) prepacked size exclusion column (Figure 2.6). The purification assay that used the Superdex 200 Increase column also had an in-line HisTrap (GE Healthcare) column to capture the His-tagged TEVp. The results of FPLC experiment with the Superdex 200 column were that the initial elution fractions from the column contained un-cleaved fusion protein and Hcf106 (Figure 2.6A, peak 1, lanes 2-3). The second protein elution fractions from the column contained only free maltose binding protein (Figure 2.6A, peak 2, lanes 5-7). The elution fractions corresponding to the 3rd and 4th chromatogram absorbance peaks did not contain any detectable protein as shown by SDS-PAGE (Figure 2.6A, peaks 3-4, lanes 8-9). The inline HisTrap column (GE Healthcare) was also checked for protein retention by eluting any remaining proteins by incubation with 2% SDS and did not contain any detectable protein (Figure 2.6A, lane 10). These conditions allowed for the removal of nearly all free MBP but failed to separate the Hcf106 from noncleaved fusion protein. As such, two additional FPLC purification trials were carried out with different parameters.

We carried out our second and third FPLC purification trials using a HiLoad 16/600 Superdex 75 pg column as it has a smaller protein mass exclusion size (3-70 kDa) than the S200 Superdex Increase column (10-600 kDa). The difference in mass exclusion size between these two columns increases resolution in the apparent mass range of free Hcf106 (~19.7 kDa). In the second purification trial, the cleavage reaction and following FPLC separation was also performed with the non-polar detergent, nonaethylene glycol monododecyl ether (C12E9, Sigma-Aldrich), instead of the zwitterionic CHAPS (Figure 2.6B). The elution fractions corresponding to the first peak contained un-cleaved fusion protein, MBP, and Hcf106 (Figure 2.6B, peak 1, lanes 4- 5). The absorbance of the second eluate peak was much lower in comparison to the first and third peaks and, correspondingly, showed trace amounts of protein (Figure 2.6B, peak 2, lanes 6-8). The final elution fractions contained MBP and TEVp (Figure 2.6B, peak 3, lanes 9-10). Ultimately, C12E9 was unable to improve separation of Hcf106 from the other proteins in the post-cleavage reaction mixture.

In our third FPLC separation experiment, we tested CHAPS detergent with the smaller mass exclusion range HiLoad 16/600 Superdex 75 pg column (Figure 2.6C). The first elution fractions contained un-cleaved fusion protein, free MBP, and Hcf106 (Figure 2.6C, peak 1, lanes 2-5). The elution fractions that correspond to the second peak on the chromatogram contained only MBP (Figure 2.6C, peak 2, lanes 6-8). Finally, the fractions corresponding to the third and fourth peaks on the chromatogram showed no visible protein by SDS-PAGE analysis (Figure 2.6C, peaks 3-4, lanes 9-12). The results of a separation of the 10:1 reaction mixture in CHAPS by S75 Superdex were nearly identical to the results of the S200 attempt (Compare Figure 2.6A with Figure 2.6C). After confirmation that these FPLC techniques were unable to separate Hcf106 from the other cleavage reaction products, we conducted a cleavage trial in which MBP-Hcf106 was bound to amylose resin.

2.4.6 TEV protease cleavage reactions with MBP-Hcf106 bound to amylose resin We also tested a resin-bound or batchwise cleavage reaction in which MBP-Hcf106 was first bound to the amylose matrix followed by the addition TEVp directly to the resin bed. Before our reaction trial experiment, we determined how buffer exchange from the initial lysis/binding buffer to a modified TEVp cleavage buffer would affect the resin bound MBP-Hcf106 (Figure 2.7A). MBP-Hcf106 was bound to amylose resin and then washed with more binding buffer (Figure 2.7A, lanes 1-2). Following this the MBP-Hcf106 bound amylose resin was exchanged into cleavage buffer and washed several times with the same buffer (Figure 2.7A, lanes 3-4). Immediately following the lysis of MBP- Hcf106 fusion expressing E. coli, the clarified lysate was incubated with the amylose resin in a gravity column (Figure 2.7B, lane 1). The eluate of the lysate from the amylose column contained fusion protein and free MBP (Figure 2.7B, lane 2). After the fusion protein was bound to the amylose resin, the column was washed with a low [CHAPS] cleavage buffer which caused elution of fusion protein and MBP (Figure 2.7B, lane 3). After equilibration with cleavage buffer, the cleavage reaction was carried out by the addition TEV protease. Despite the successful cleavage reaction in the column, the non-cleaved fusion protein and free MBP was not retained on the column (Figure 2.7B, lanes 4-5).

Figure 2.7 Buffer exchange and batchwise amylose resin proteolysis of MBP-Hcf106 with TEVp. A Lane 1, amylose resin bound MBP-Hcf106 in binding buffer (20 mM Tris pH 7.4, 200 mM NaCl, 1 mM EDTA, 1 mM DTT, 1 mM PMSF, 0.075 mM CHAPS). Lane 2, amylose resin washed with binding buffer. Lane 3, flow-through after exchange of MBP-Hcf106 bound amylose resin into cleavage buffer (50 mM Tris pH 7.4, 1 mM EDTA, 1 mM DTT, 0.075 mM CHAPS). Lane 4, fraction from washing amylose resin with cleavage buffer. B Lane 1, clarified lysate from BL21 E. coli expressing MBP-Hcf106. Lane 2, flow- through from passing MBP-Hcf106 lysate over amylose resin column. Lane 3, exchange of MBP-Hcf106 bound amylose resin into cleavage buffer. Lane 4, buffer flow through following TEVp 24-hour cleavage reaction at 4°C. Lane 5, 5x concentrate of buffer flow-through post proteolysis.

2.4.7 Additional proteolysis reaction tests in the presence of urea The addition of 300 mM urea to the reaction improved the overall efficiency of TEVp (Figure 2.5C), so we tested additional urea concentrations at two reaction temperatures and timepoints (Figure 2.8). New batches of MBP-Hcf106 and TEVp were prepared according the materials and methods covered previously. The results of 10:1 MBP- Hcf106:TEVp reactions at 25°C and 4°C were quite similar (Figure 2.8A). Regardless of the presence or absence of 300 mM urea, the 25°C reactions were more complete after 18 hr than after 4 hr as shown by a decrease in substrate and an increase in free MBP and Hcf106 (Figure 2.8A). The reactions carried out at 4°C also show the time dependent increase in products and decrease in substrate regardless of the individual urea concentrations (Figure 2.8A). In response the results of the 10:1 reactions, we also tested 25:1 and 50:1 reactions in TEVp reaction buffer with 300 mM urea (Figure 2.8B). The results of the cleavage reaction at 25°C following 4 hr and 36 hr of reaction time are very similar to those for the 10:1 reactions (Figure 2.8).

Figure 2.8 Testing additional parameters for the TEVp reaction with MBP-Hcf106 including different urea concentrations and protease ratios. A 15% SDS-PAGE analysis of 10:1 MBP- Hcf106:1 reactions at 25°C (lanes 2-9) and 4°C (lanes 11-18). Lanes 1 and 10 are TEVp alone at the 10:1 dilution. Four concentrations of urea were tested in these reactions: 0 mM (lanes 2-3, 11-12), 100 mM (lanes 4-5, 13-14), 300 mM (lanes 6-7), and 500 mM (lanes 8-9, 17-18). Individual 25°C reaction samples were drawn after 4 hr (lanes 2, 4, 6, 8) and 18 hr (lanes 3, 5, 7, 9). Individual 4°C reaction samples were drawn after 18 hr (lanes 11, 13, 15, 17) and 32 hr (lanes 12, 14, 16, 18). B 15% SDS- PAGE results of 25:1 (lanes 2-3) and 50:1 (lanes 5-6) MBP-Hcf106:TEVp reactions at 25°C after 4 hr (lanes 2, 5) and 36 hr (lanes 3, 6) in reaction buffer containing 300 mM urea. Lane 1 is MBP-Hcf106 diluted to be equivalent to the quantity of starting reaction substrate. Lane 4 is TEVp diluted to be equivalent to the quantity of protease in a 50:1 reaction.

As the 50:1 reaction progressed to almost the same degree as the 10:1 reaction, we then tested a method for the removal of TEVp from the post-cleavage reaction mixture using the hexahistidine tag once again. To make this process as quick and efficient as possible, we used HisPur Ni-NTA magnetic beads (Thermo Scientific) to capture the

Figure 2.9 Removal of TEVp from the reaction mixture and subsequent purification by FPLC. A 15% SDS-PAGE showing removal of TEVp from a 32 hr, 50:1 reaction with 300 mM urea using His Mag Sepharose Ni (GE Healthcare). Lane 1 is EZ-Run Pre-stained Rec protein ladder (Fisher). Lane 2 is the 50:1 reaction. Lanes 3-4 are the supernatant collected after incubation with the Ni-NTA magnetic beads and the eluate with 500 mM imidazole. Lanes 6-7 are the supernatant and eluate fractions after a second incubation with Ni-NTA magnetic beads. B mAU280nm chromatogram following the elution of protein fractions after protease removal in (A). C FPLC fractions from 2.9B analyzed by 12.5% SDS- PAGE and Coomassie Blue staining. Lane 1 is TEVp at 50:1 dilution. Lane 2 is the reaction mixture after TEVp removal from 2.9A. Lanes 3-5 are pooled fractions from FPLC in 2.9B. Lane 3 is fractions 11-15 (2.9B, peak 1), lane 4 is fractions 33-38 (2.9B, peak 2), and lane 5 is fractions 51-58 (2.9B, peak 3). D Western blot analysis of the same SDS-PAGE samples from 2.9C with αHcf106 IgG (see Materials and Methods 2.3.9).

TEVp from a 50:1 reaction in the presence of 300 mM urea containing binding and elution buffers (Figure 2.9A). All of the TEVp in the reaction was bound to the magnetic Ni-NTA beads after the first incubation as subsequent binding and elution steps showed no additional protease remaining (Figure 2.9A, lanes 4-7). After successful removal of TEVp from the reaction mixture, FPLC was used to further separate free Hcf106 from the remaining MBP-Hcf106 and MBP (Figure 2.9B). A HiLoad 16/600 Superdex 200 pg column (GE Healthcare) was used with the TEVp reaction buffer containing 300 mM urea (Figure 2.9B). Three peaks corresponding to protein elution fractions were reported on the chromatogram (Figure 2.9B). The fractions that correspond to each peak on the chromatogram were pooled and prepared for further analysis. SDS-PAGE with Coomassie blue staining (Figure 2.9C) and immunoblot analysis (Figure 2.9D) were used to identify the contents of each set of pooled fractions. The fractions corresponding to first elution peak contained Hcf106 and MBP-Hcf106 as shown by the appropriate molecular masses for each Coomassie stained band and the high degree of chemiluminescence (Figure 2.9B, peak 1, 2.9C-D, lane 3). The second peak and corresponding FPLC elution fractions contained only free MBP (Figure 2.9B, peak 2, 2.9C-D, lane 4). The final eluate fractions that correspond to the third peak on the chromatogram contained no Coomassie blue stained proteins nor any detectable chemiluminescence (Figure 2.9B, peak 3, 2.9C-D, lane 5).

Although we were able to remove TEVp and MBP from the reaction mixture, free Hcf106 was not separated from intact MBP-Hcf106 using FPLC with the 300 mM urea reaction buffer (Figure 2.9). In order to disrupt the interactions between free Hcf106 and MBP-Hcf106, we increased the urea concentration to 4 M in the pooled fractions and

Figure 2.10 Incubation of MBP-Hcf106/Hcf106 and subsequent FPLC separation with 4 M urea. A Chromatogram of absorbance at 280 nm monitoring the elution of proteins from the pooled fractions 11-15 from the previous FPLC separation (Figure 2.9C, lane 3) after incubation in 4 M urea reaction buffer. B 4 M buffer FPLC elution fractions were pooled and analyzed by 15% SDS-PAGE. Lane 1 is pooled fractions 11-15 after incubation with 4 M urea prior to FPLC separation. Lane 2 is the contents of pooled fractions 12-16 (2.9A, peak 1). Lane 3 is the contents of pooled fractions 52-56 (2.9A, peak 2). Lane 4 is from pooled fractions 58-63 (2.9A, peak 3).

45 repeated the FPLC separation experiment with the same urea concentration in the column buffer (Figure 2.10A). The elution fractions that correspond to the first peak on the chromatogram contained free Hcf106 and MBP-Hcf106 (Figure 2.10A, peak 1, 2.10B, lanes 1-2). The elution fractions that correspond to the second and third chromatogram peaks contained no protein as shown by a lack of staining with Coomassie blue (Figure 2.10A, peaks 2-3, 2.10B, lanes 3-4).

2.4.8 Testing additional detergents used in previously published purification trials of recombinant TatB from E. coli Our attempts at separating free Hcf106 from noncleaved MBP-Hcf106 by FPLC in the presence of 300 mM and 4 M urea were unsuccessful (Figure 2.10). As such, we decided to test two detergents that were used to purify recombinant, truncated TatB from E. coli; these detergents are Anzergent 3-14 (AZ314) and Fos-choline-12 (DPC) (Anatrace) (Zhang et al., 2014b). To start this process, we exchanged the already purified MBP-Hcf106 and TEVp into reaction buffer with either 0.5% AZ314 or 0.5% DPC using Vivaspin concentrators with a 3000 MWCO (Millipore). After several filtration steps, 50:1 MBP-Hcf106:TEVp reactions were conducted at both 25°C and 4°C with samples drawn to monitor reaction progress after 4 hr and 24 hr (Figure 2.11). TEVp activity was markedly decreased in the presence of either detergent tested as compared to no urea or 100/300/500 mM urea (Figure 2.8 and 2.11).

Figure 2.11 Testing reaction efficiency in the presence of additional detergents. 15% SDS-PAGE results of 50:1 MBP-Hcf106:TEVp reactions in the presence of either 0.5% AZ314 (lanes 2-5) or 0.5% DPC (Lanes 6-9). Reactions containing each detergent were tested at 4°C (lanes 2-3, 6-7) or 25°C (lanes 4-5, 8-9). Samples from each reaction were drawn after 4 hr (lanes 2, 4, 6, 8) and 24 hr (lanes 3, 5, 7, 9). Lane 1 is TEVp diluted to be equivalent to the amount of protease in each 50:1 reaction.

2.5 Discussion Despite recent advances in determining the structure of Hcf106/TatB through overexpression or synthesis followed by biophysical characterization, these studies have been limited to truncated versions on the protein in mimetic membrane environments (Zhang et al., 2013; Zhang et al., 2014a; Zhang et al., 2014b). The work presented here set out to add to and expand upon these previous studies by purifying and characterizing a full-length version of Hcf106. To accomplish this aim, we engineered a protein expression vector to generate a full-length version of Hcf106 fused

46 to a solubility enhancing protein (Figure 2.2). We chose MBP as it has been previously shown to enhance the solubility of membrane proteins when fused to it (Lebendiker and Danieli, 2011). In addition to improving solubility, MBP-linked fusion proteins have been easily purified by affinity chromatography using commercially available amylose resins (Lebendiker and Danieli, 2011). We successfully generated a fusion construct in a pMAL-c5E vector using molecular biology techniques ultimately linking MBP to full- length Hcf106 with a TEV protease recognition sequence (Figure 2.2). We utilized molecular biology techniques to overwrite the enterokinase recognition sequence with a TEVp recognition sequence in the commercially available pMAL-c5E plasmid. A protease recognition sequence is required so that we could remove MBP from Hcf106 following purification. We then used a previously published method to express and purify TEVp (Figure 2.3) from BL21 E. coli (Tropea et al., 2009). Finally, we were able to overexpress this MBP-Hcf106 fusion protein in E. coli and isolate it from lysate using amylose resin affinity chromatography (Figure 2.4).

Although we were able to express and purify both MBP-Hcf106 and TEVp in sufficient quantities from E. coli, we struggled to achieve efficient proteolysis of the fusion protein. Even at the highest ratio of protease:substrate, all of the fusion protein present in the reaction was not cleaved by TEVp (Figure 2.5, 2.8). To improve the efficiency of proteolysis we added urea to a final concentration of 300 mM in the reaction buffer (Figure 2.5C, 2.8). Urea has been previously utilized to improve the efficiency of TEVp cleavage of dengue virus NS4A peptide fusions to a glutathione S-transferase (GST) tag (Hung et al., 2014). This improvement in efficiency is likely due to the mild protein denaturing properties of urea which ultimately allows easier or increased access to the protease recognition sequence. In our case, the transmembrane domain of Hcf106 is hydrophobic (similar to other membrane anchoring protein regions and membrane transmembrane helices) and thus is likely to interact with other hydrophobic protein domains such as other Hcf106 TM region. These interactions could then lead to steric occlusion of the TEVp recognition sequence ultimately slowing or preventing proteolysis. By adding urea to the reaction buffer, any protein-protein reactions that would prevent access to the recognition sequence would be diminished and TEVp would have better access to the TEVp recognition sequence leading to increased cleavage efficiency. Although urea improved the efficiency of the cleavage reaction, complete cleavage of MBP-Hcf106 was not attained with this denaturant additive, so we also tested the addition of detergents to improve the proteolytic efficiency of TEVp. (Figure 2.5C, 2.8).

We hypothesized that the addition of detergent to the reaction would solubilize and prevent any aggregation any form of Hcf106 present whether free or still fused to MBP. We decided to test CHAPS, C12E9, and AZ314 as these detergents had been previously used to solubilize bacterial TAT proteins including the Hcf106 homolog TatB (de Leeuw et al., 2002; De Leeuw et al., 2001; Sargent et al., 2001). CHAPS and C12E9 detergents have also been shown to be compatible with TEVp activity (Lundback et al., 2008; Vergis and Wiener, 2011). The addition of the zwitterionic detergent CHAPS to the proteolysis reaction also improved cleavage efficiency but to a lesser degree than the addition of urea (Figure 2.5C). The hydrophobic detergent

C12E9 was also examined for compatibility with Hcf106 following TEVp cleavage reactions but was less effective at improving the efficiency of the reaction than urea or CHAPS (Figure 2.6).

Despite being unable to achieve complete proteolysis of the MBP-Hcf106 fusion protein, we also tested the feasibility of separation of the reaction products using FPLC with varied column exclusion limits and with buffers containing C12E9 and CHAPS detergents (Figure 2.6). The first separation by FPLC was carried out on a Superdex 200 Increase column with a separation range from 10000 to ~600000 Da, relative molecular mass (Mr) (GE Healthcare) in CHAPS detergent (Figure 2.6A). The first protein containing eluant fractions contained a mixture of MBP-Hcf106 and Hcf106, possibly due to interactions with the fusion protein and free Hcf106 (Figure 2.6A). The only other meaningful protein eluant fraction contained a strongly stained band corresponding to MBP and two faint bands, one of which corresponds to Hcf106 (~24- 25 kDa) that appears beneath the band corresponding to TEVp (~27 kDa) (Figure 2.5, 2.6). We also used a HiLoad 16/600 Superdex 75 pg column (GE Healthcare) with a separation range from ~3000 to 70000 Mr. The benefit of using this column was that we could load up to 5 mL of reaction mixture. However, similar results were attained using this larger column in both C12E9 and CHAPS detergents (Figure 2.6B-C). In both cases, the first protein eluate fractions contained MBP-Hcf106 and Hcf106 (Figure 2.6B-C). The results of the C12E9 detergent FPLC trial also showed free MBP that was eluted in the first protein fractions (Figure 2.6B). The latter fractions contained most of the free MBP for both detergents tested (Figure 2.6B-C). However, a band that corresponds to Hcf106 based on the previous proteolysis SDS-PAGE results eluted with free MBP in C12E9 detergent buffer (Figure 2.6B). Each attempt at FPLC was unable to separate free Hcf106 from the rest of the reaction components. Ultimately, the reaction buffer conditions still need to be optimized for maximal TEVp efficiency while ensuring that the hydrophobic domains of Hcf106 are solubilized after the removal of the MBP tag.

In order to simplify the purification process, we tried to cleave the MBP-Hcf106 fusion protein while the MBP tag was bound to amylose resin as has been previously carried out (Zhu et al., 2017). By cleaving Hcf106 from MBP bound to the resin, free Hcf106 and TEVp would be the only two proteins washed from the resin with buffer lacking maltose. First, we used amylose resin to bind MBP-Hcf106 in binding buffer and then exchanged the fusion protein into a TEVp cleavage buffer containing CHAPS (Figure 2.7A). The exchange into CHAPS containing buffer did remove a slight amount of nonspecifically bound MBP-Hcf106 (Figure 2.7A). We then were able to test the batchwise cleavage reaction (Figure 2.7B). However, following the cleavage reaction, the buffer collected from the column resin contained all four protein components as well as many nonspecific bands that are likely proteolytic fragments (Figure 2.7B). The major issue with this approach is that the batchwise buffer exchange process disrupted the interactions between MBP and the resin leading to elution of the free MBP and MBP-Hcf106 prior to as well as after the proteolysis reaction. This process could still be utilized but new buffer/detergent considerations must be tested and optimized.

We also further explored the addition of urea to the proteolysis reaction. Newly purified MBP-Hcf106 was efficiently cleaved by TEVp in reactions with ratios from 10:1 to 50:1 at 25°C and 4°C (Figure 2.8). The reaction even proceeded at 10:1 without the addition of urea (Figure 2.8A). After further optimization of cleavage reaction parameters, we were successful in completely removing TEVp with magnetic Ni-NTA beads from a 50:1 reaction (Figure 2.9A). Although we were able to remove TEVp, the subsequent FPLC trials using buffers containing either 300 mM or 4 M urea were unsuccessful in separating MBP-Hcf106 from free Hcf106 (Figure 2.9B-D, 2.10). This is likely due to poor solubilization of free Hcf106 after the proteolysis reaction as these reactions contain no detergent or lipid additives to interact with the hydrophobic portions of the protein (Figure 2.9B-D, 2.10).

To tackle the problems of Hcf106 solubility, we carried out cleavage reactions in the presence of the detergents AZ314 or DPC previously used to solubilize recombinant TatB (Zhang et al., 2014b). The efficiency of TEVp was markedly decreased in reactions containing either detergent under all tested conditions (Figure 2.11). Our results agree with a published comprehensive study of TEVp and 5 other proteases that showed changes proteolytic efficiency in the presence of 94 different detergents (Vergis and Wiener, 2011). TEVp was shown to have relatively low activity for soluble fusion protein substrates in reactions containing either AZ314 or DPC (Vergis and Wiener, 2011). Thus, we need to screen additional detergents in cleavage reactions that have been shown to not impact TEVp efficiency (Vergis and Wiener, 2011).

2.6 Conclusions and Future Directions Although we were unable to purify full-length Hcf106, the work presented in this chapter is a foundation for future purification trials of MBP-Hcf106. First, we established a reproducible, high yield expression and purification protocol for our fusion protein. We also optimized several cleavage reaction parameters. Future studies can remove multiple conditions that fail to produce pure Hcf106 from consideration, for example, confirming TEVp detergent incompatibility. Finally, we were able to remove TEVp and free MBP from Hcf106 and intact fusion protein using commercially available and easily adapted techniques such as magnetic Ni-NTA resin and FPLC. So, one of the first future avenues of investigation would be attempt unfolding Hcf106 and MBP-Hcf106 soluble aggregates in concentrated guanidinium HCl buffer. However, we would then need to determine if Hcf106 can be refolded and further purified from intact fusion protein.

Another future direction for this project is to use a protease that is compatible with AZ314 and DPC. Although TEVp is only modestly active in AZ314 or DPC, the proteases thrombin and enterokinase were previously shown to be highly active in these detergents (Vergis and Wiener, 2011). In addition to testing alternative detergent- protease combinations, there are other TEVp variants that can be purified from E. coli or purchased commercially (Sanchez and Ting, 2020; Zhu et al., 2017). These TEVp variants have higher overall proteolytic efficiency than the version we used in this study as well as having drastically improved affinity for resin bound fusion proteins making ideal for use in our batchwise cleavage assays (Zhu et al., 2017). We would still need to

49 test the detergent compatibility of new TEVp variants and then optimize the cleavage reaction once again.

Finally, an alternative methodology for purifying Hcf106 from E. coli was recently published (Zinecker et al., 2020). In this work, the authors purified full-length Hcf106 with a chimeric N-terminal His/S-tag, showed that it integrates into thylakoid membranes, and complements loss of transport function (Zinecker et al., 2020). In their scheme, Hcf106 was cleaved from the chimeric His/S-tag by BrCN treatment and further purified by an extensive multistep process including preparative SDS-PAGE, Zn- imidazole staining, electroelution, reversed phase HPLC, and refolding by dialysis (Zinecker et al., 2020). Although the expressed Hcf106 integrated into and functioned in thylakoid, there was a consistent loss of protein throughout the purification process, likely due to aggregation or misfolding (Zinecker et al., 2020). So, this protein purification process may be of use for structural characterization of Hcf106 as the authors report that they routinely achieved ~38 mg/L (2 mM) concentrations of purified protein.

2.7 References Aldridge, C., X. Ma, F. Gerard, and K. Cline. 2014. Substrate-gated docking of pore subunit Tha4 in the TatC cavity initiates Tat translocase assembly. J Cell Biol. 205:51-65. Aldridge, C., A. Storm, K. Cline, and C. Dabney-Smith. 2012. The chloroplast twin arginine transport (Tat) component, Tha4, undergoes conformational changes leading to Tat protein transport. J Biol Chem. 287:34752-34763. Berks, B.C. 2015. The twin-arginine protein translocation pathway. Annu Rev Biochem. 84:843-864. Bryksin, A.V., and I. Matsumura. 2010. Overlap extension PCR cloning: a simple and reliable way to create recombinant plasmids. Biotechniques. 48:463-465. Cline, K., W.F. Ettinger, and S.M. Theg. 1992. Protein-specific energy requirements for protein transport across or into thylakoid membranes. Two lumenal proteins are transported in the absence of ATP. J Biol Chem. 267:2688-2696. Cross, T.A., M. Sharma, M. Yi, and H.X. Zhou. 2011. Influence of solubilizing environments on membrane protein structures. Trends Biochem Sci. 36:117-125. Dabney-Smith, C., H. Mori, and K. Cline. 2003. Requirement of a Tha4-conserved transmembrane glutamate in thylakoid Tat translocase assembly revealed by biochemical complementation. J Biol Chem. 278:43027-43033. de Leeuw, E., T. Granjon, I. Porcelli, M. Alami, S.B. Carr, M. Muller, F. Sargent, T. Palmer, and B.C. Berks. 2002. Oligomeric properties and signal peptide binding by Escherichia coli Tat protein transport complexes. J Mol Biol. 322:1135-1146. De Leeuw, E., I. Porcelli, F. Sargent, T. Palmer, and B.C. Berks. 2001. Membrane interactions and self-association of the TatA and TatB components of the twin- arginine translocation pathway. FEBS Lett. 506:143-148. Geiser, M., R. Cebe, D. Drewello, and R. Schmitz. 2001. Integration of PCR fragments at any specific site within cloning vectors without the use of restriction enzymes and DNA ligase. Biotechniques. 31:88-90, 92.

Gerard, F., and K. Cline. 2006. Efficient twin arginine translocation (Tat) pathway transport of a precursor protein covalently anchored to its initial cpTatC binding site. J Biol Chem. 281:6130-6135. Gerard, F., and K. Cline. 2007. The thylakoid proton gradient promotes an advanced stage of signal peptide binding deep within the Tat pathway receptor complex. J Biol Chem. 282:5263-5272. Hamsanathan, S., and S.M. Musser. 2018. The Tat protein transport system: intriguing questions and conundrums. FEMS Microbiol Lett. 365. Hu, Y., E. Zhao, H. Li, B. Xia, and C. Jin. 2010. Solution NMR structure of the TatA component of the twin-arginine protein transport system from gram-positive bacterium Bacillus subtilis. J Am Chem Soc. 132:15942-15944. Hung, Y.F., O. Valdau, S. Schunke, O. Stern, B.W. Koenig, D. Willbold, and S. Hoffmann. 2014. Recombinant production of the amino terminal cytoplasmic region of dengue virus non-structural protein 4A for structural studies. PLoS One. 9:e86482. Kunkel, T.A. 1985. Rapid and efficient site-specific mutagenesis without phenotypic selection. Proc Natl Acad Sci U S A. 82:488-492. Lebendiker, M., and T. Danieli. 2011. Purification of proteins fused to maltose-binding protein. Methods in molecular biology (Clifton, N.J.). 681:281-293. Lundback, A.K., S. van den Berg, H. Hebert, H. Berglund, and S. Eshaghi. 2008. Exploring the activity of tobacco etch virus protease in detergent solutions. Anal Biochem. 382:69-71. Ma, X., and K. Cline. 2010. Multiple precursor proteins bind individual Tat receptor complexes and are collectively transported. EMBO J. 29:1477-1488. Matin, T.R., K.P. Sigdel, M. Utjesanovic, B.P. Marsh, F. Gallazzi, V.F. Smith, I. Kosztin, and G.M. King. 2017. Single-Molecule Peptide-Lipid Affinity Assay Reveals Interplay between Solution Structure and Partitioning. Langmuir. 33:4057-4065. Mohanty, A.K., C.R. Simmons, and M.C. Wiener. 2003. Inhibition of tobacco etch virus protease activity by detergents. Protein Expr Purif. 27:109-114. New, C.P., Q. Ma, and C. Dabney-Smith. 2018. Routing of thylakoid lumen proteins by the chloroplast twin arginine transport pathway. Photosynth Res. Phan, J., A. Zdanov, A.G. Evdokimov, J.E. Tropea, H.K. Peters, 3rd, R.B. Kapust, M. Li, A. Wlodawer, and D.S. Waugh. 2002. Structural basis for the substrate specificity of tobacco etch virus protease. J Biol Chem. 277:50564-50572. Sahu, I.D., and G.A. Lorigan. 2018. Site-Directed Spin Labeling EPR for Studying Membrane Proteins. Biomed Res Int. 2018:3248289. Sahu, I.D., R.M. McCarrick, K.R. Troxel, R. Zhang, H.J. Smith, M.M. Dunagan, M.S. Swartz, P.V. Rajan, B.M. Kroncke, C.R. Sanders, and G.A. Lorigan. 2013. DEER EPR measurements for membrane protein structures via bifunctional spin labels and lipodisq nanoparticles. Biochemistry. 52:6627-6632. Sanchez, M.I., and A.Y. Ting. 2020. Directed evolution improves the catalytic efficiency of TEV protease. Nat Methods. 17:167-174. Sargent, F., U. Gohlke, E. De Leeuw, N.R. Stanley, T. Palmer, H.R. Saibil, and B.C. Berks. 2001. Purified components of the Escherichia coli Tat protein transport system form a double-layered ring structure. Eur J Biochem. 268:3361-3367.

Settles, A.M., A. Yonetani, A. Baron, D.R. Bush, K. Cline, and R. Martienssen. 1997. Sec-independent protein translocation by the maize Hcf106 protein. Science. 278:1467-1470. Tropea, J.E., S. Cherry, and D.S. Waugh. 2009. Expression and purification of soluble His(6)-tagged TEV protease. Methods in molecular biology (Clifton, N.J.). 498:297-307. Vergis, J.M., and M.C. Wiener. 2011. The variable detergent sensitivity of proteases that are utilized for recombinant protein affinity tag removal. Protein Expr Purif. 78:139-142. Zhang, L., L. Liu, S. Maltsev, G.A. Lorigan, and C. Dabney-Smith. 2013. Solid-state NMR investigations of peptide-lipid interactions of the transmembrane domain of a plant-derived protein, Hcf106. Chem Phys Lipids. 175-176:123-130. Zhang, L., L. Liu, S. Maltsev, G.A. Lorigan, and C. Dabney-Smith. 2014a. Investigating the interaction between peptides of the amphipathic helix of Hcf106 and the phospholipid bilayer by solid-state NMR spectroscopy. Biochim Biophys Acta. 1838:413-418. Zhang, Y., L. Wang, Y. Hu, and C. Jin. 2014b. Solution structure of the TatB component of the twin-arginine translocation system. Biochim Biophys Acta. 1838:1881- 1888. Zhu, K., X. Zhou, Y. Yan, H. Mo, Y. Xie, B. Cheng, and J. Fan. 2017. Cleavage of fusion proteins on the affinity resins using the TEV protease variant. Protein Expr Purif. 131:27-33. Zinecker, S., M. Jakob, and R.B. Klosgen. 2020. Functional reconstitution of TatB into the thylakoidal Tat translocase. Biochim Biophys Acta Mol Cell Res. 1867:118606.

Chapter 3: Increases in Tha4 Transmembrane Helix Hydrophobicity Alter Function and Organization During Chloroplast Twin Arginine Transport

Christopher Paul New1, Krystina Hird2, Vidusha Weesinghe2, Jessica Pax2, Andrew Abata2, Carole Dabney-Smith1,2,*

1Graduate program in Cell, Molecular, and Structural Biology, Miami University, Oxford, Ohio 45056

2Department of Chemistry and Biochemistry Miami University, Oxford, Ohio 45056

*Corresponding author: Department of Chemistry and Biochemistry, Miami University, 651 East High St., Oxford, OH. Tel.: 513-529-8091; E-mail: [email protected]

Author contributions: CPN and CDS performed the data analysis and wrote the manuscript; CPN generated the protein variants, performed the bioinformatics analysis, generated the protein models, and completed the BN-PAGE experiments; CPN and KH performed the complementation assays; CPN, KH, VW, JP, and AA performed oligomerization experiments; CPN and VW performed the alkaline extraction assays.

To be submitted to Plant Physiology

3.1 Abstract Tha4, a cpTAT component protein, has an obligate glutamate in its transmembrane helix. Substitution of this glutamate to alanine prevents transport while substitution to aspartate moderately recovers transport. It has also been shown that this Tha4 alanine variant has a different oligomerization profile than wild type Tha4 in the presence of the proton motive force and functional/non-functional substrate proteins. We designed and carried out a series of experiments to determine if substitution of this glutamate into lumen proximate positions in the transmembrane helix of the non-functional Tha4 alanine complemented loss of transport function. Other than in the wild type positions, glutamate substitutions were not tolerated in the alanine background while aspartate variants were slightly tolerated. We also developed assays to test Tha4 glutamate/alanine/aspartate variant oligomer formation in the presence or absence of substrate and PMF. Tha4 oligomer formation is enhanced by substitution of transmembrane glutamate with alanine and diminished by aspartate substitution due to changes in the interactions between individual Tha4 variant monomers. The changes in Tha4 packing and organization were linked to altered monomer and oligomer stability in the thylakoid membrane by modulation of transmembrane helix hydrophobicity.

3.2 Introduction Chloroplasts require nuclear and organellar gene expression for proper function. These genes encode for nearly 3000 different proteins in the organelle, most of which are translated in the cytosol of the plant cell. These newly synthesized proteins are transported into the organelle by the translocons on the outer and inner chloroplast membranes (TOC and TIC, respectively) (Paila et al., 2015; Thomson et al., 2020). After import by TOC/TIC, a subset of these proteins is further routed to the thylakoid membrane or lumen. One of two systems are used to transport proteins from the stromal compartment of the chloroplast to the lumen of the thylakoid (Cline and Dabney- Smith, 2008; New et al., 2018). The first system is the chloroplast secretory pathway (cpSec) which uses energy derived from ATP hydrolysis and a proton motive force (PMF) to thread unfolded precursor proteins into the thylakoid lumen through a SecY/E channel with the aid of the SecA protein (Albiniak et al., 2012; Yuan et al., 1994). The second pathway, called the chloroplast twin arginine transport (cpTAT) system, translocates fully folded proteins across the thylakoid membrane using only the energy of the trans-thylakoidal PMF (Mould and Robinson, 1991; New et al., 2018). The cpTAT system is also homologous to bacterial and archaeal TAT systems (Berks, 2015; Hamsanathan and Musser, 2018; Yen et al., 2002). The name of the TAT system comes from the presence of the conserved twin arginine motif (RR) found in the N- terminal signal peptide sequences of TAT specific substrate proteins (Berks et al., 2000; Peltier et al., 2002). The cpTAT pathway is composed of three proteinaceous components. The first (cp)TAT component protein is (cp)TatC, a multi-pass membrane protein with six linked transmembrane helices that has an overall glove-like shape (Mori et al., 2001; Ramasamy et al., 2013; Rollauer et al., 2012). The other two cpTAT component proteins are Hcf106 (Settles et al., 1997) and Tha4 (Walker et al., 1999). In bacteria and archaea, these proteins are TatB and TatA, respectively (Berks, 2015; Berks et al., 2003; Muller and Klosgen, 2005). Additionally, not all TAT systems have a TatB component while others have an additional TatE component (Berks, 2015). In

cpTAT, Tha4 and Hcf106 have similar overall structures and domains: each has an N- terminal transmembrane α-helix (TMH) linked to an amphipathic α-helix by a short hinge region followed by a soluble, unstructured carboxy-terminal “tail” (Fincher et al., 2003; Mori and Cline, 2001). cpTAT-mediated transport of substrate proteins operates in a cyclical fashion. cpTatC- Hcf106-Tha4 function as a heterotrimeric precursor receptor complex that interacts with and binds the twin arginine motif located in the signal peptide of (cp)TAT specific substrate/cargo proteins (Alcock et al., 2016; Cline and Mori, 2001; Habersetzer et al., 2017). This initial “shallow” binding is followed by a “deep” signal peptide insertion into the (cp)TAT binding pocket in the presence of PMF (Gerard and Cline, 2006; Gerard and Cline, 2007). Following this deep binding, Tha4 (TatA) from a separate pool in the thylakoid membrane organizes into oligomers at the receptor complex, possibly via nucleation at Tha4 (TatA) that is constitutively bound with cpTatC-Hcf106 (TatC-TatB) (Alcock et al., 2016; Aldridge et al., 2014; Gerard and Cline, 2007; Habersetzer et al., 2017). After the oligomerization and assembly of additional Tha4, translocation of the mature domain of the substrate protein occurs. The signal peptide is then cleaved by a signal processing peptidase which frees the mature substrate protein to diffuse from the (cp)TAT complex to its location of function be it in the thylakoid lumen or outside of the plasma membrane in bacteria/archaea (Cline, 2015; Frobel et al., 2012; Luke et al., 2009). Tha4 (TatA) oligomers then dissociate from the receptor complex to reset the (cp)TAT system for the subsequent rounds of substrate binding and transport (Alcock et al., 2013; Dabney-Smith and Cline, 2009).

For proper cpTAT-mediated substrate transport, Tha4 requires the presence of a transmembrane glutamate (Tha4 E10) (Dabney-Smith et al., 2003). Substitutions of that glutamate to an alanine (Tha4 E10A) or glutamine (Tha4 E10Q) failed to restore transport while substitution to aspartate (Tha4 E10D) restored transport, although to less than half that of wild type (Dabney-Smith et al., 2003). Substitution of this glutamate to alanine also increased the resistance of Tha4 to alkaline extraction from the thylakoid membrane (Dabney-Smith et al., 2003). In addition, Tha4 oligomerization between adjacent TMHs or carboxy-terminal tails was enhanced in the presence of both the PMF and cpTAT specific signal peptides or full-length precursor proteins (Dabney- Smith and Cline, 2009). However, oligomerization between Tha4 monomers in both of these structural regions decreased when the PMF was depleted even in the presence of a truncated cpTAT signal peptide (Dabney-Smith and Cline, 2009). Likewise, the mature domain of cpTAT substrates appears to stabilize the formation of Tha4 oligomers through interactions between the amphipathic helix of Tha4 and the mature domain of precursor (Pal et al., 2013). These findings led to us to further question the role of the Tha4 TMH glutamate in transport function and Tha4 organization in cpTAT.

Currently, the contribution of the transmembrane glutamate (glutamate 10, E10) to Tha4 function in cpTAT is unknown. Specifically, we do not know if proper positioning of this glutamate in the Tha4 TMH is a requirement for function. This led us to question if the loss of transport recovery with Tha4 E10A would be restored if a glutamate (or aspartate) was introduced into the TMH in positions nearer to lumen. Our rationale

being that there are protonated and deprotonated populations of Tha4 in the thylakoid membrane. In the presence of the PMF, the population of protonated Tha4 would increase while Tha4 E10A wouldn’t be affected. So, reintroduction of an acidic amino acid to the TMH of Tha4 E10A could lead to the reformation of these two subpopulations. We could then test these variants for complementation of function in cpTAT. This would allow us to determine if Tha4 function is related to the position of the glutamate or merely the presence of a protonatable residue in the TMH.

In addition to determining how the position of a protonatable residue in the Tha4 TMH relates to transport recovery, we also wanted to determine if these substitutions impacted Tha4 organization prior to and during transport. Specifically, we wanted to test how the oligomer formation of each Tha4 E10/A/D variant is affected by presence or absence of PMF and full-length precursor. Based on these questions, we formed the hypothesis that the Tha4 transmembrane glutamate may function as a sensor for the formation of PMF prior to cpTAT-mediated transport (Figure 3.1A). We reason that as the environment around this residue becomes acidified during the formation of the PMF, there is an increase in protonated Tha4 glutamate residues (Figure 3.1A). Once the R- group carboxylate is neutralized by protonation, the movement of Tha4 further into the hydrophobic membrane core to interact with cpTatC TM4 is more energetically favorable as previously proposed (Aldridge et al., 2014; Aldridge et al., 2012) (Figure 3.1A-B).

In this study, we set out to determine if glutamate 10 senses the PMF/pH gradient across the thylakoid membrane leading to changes in Tha4 oligomerization and function. We found that Tha4 E10A with glutamate substitutions in the TMH were unable to restore cpTAT-mediated transport of substrate. When we made aspartate mutations in the TMH of Tha4 E10A cpTAT function was weakly restored. As for oligomerization of Tha4, interaction between adjacent Tha4 C-tails occurs independently of the PMF in Tha4 E10/A/D variants. We also determined that interactions between adjacent Tha4 TMH domains are enhanced in the presence of precursor protein regardless of PMF dissipation. The data led us to propose a model of Tha4 TMH conformational change.

3.3 Materials and Methods 3.3.1 Source Plants, Chloroplast and Thylakoid Membrane Isolation Chloroplasts from 10- to 11-day old pea plants (Little Marvel) were isolated according to published procedures (Cline et al., 1993). Intact chloroplasts were suspended in import buffer (IB: 50 mM HEPES-KOH, pH 8.0, 300 mM sorbitol,) to 1 mg/mL chlorophyll as determined by UV-Vis absorbance at 663 and 645 nm (Arnon, 1949). Thylakoids were isolated from intact chloroplasts by hypotonic lysis as described previously (Cline et al., 1993). Briefly, pelleted, intact chloroplasts were suspended in hypotonic lysis buffer (20 mM HEPES-KOH, pH 8, 10 mM MgCl2) and incubated on ice for 10 min before centrifugation at 3000 x g for 8 min. The recovered supernatant was clarified by further centrifugation at 42000 x g for 30 min and saved as “stromal extract.” Clarified stromal extract was used to dilute in vitro translated precursor proteins. Recovered thylakoids

were resuspended to 1 mg/mL chlorophyll in IBM (IB with10 mM MgCl2) and stored on ice until use.

Figure 3.1 Tha4 has a membrane embedded glutamate in its TMH as shown by cartoon structures modeled alone in the thylakoid membrane and during interaction with receptor complex. A Tha4 (pink) primary sequence residues 55-137 from P. sativum (Uniprot: Q9XH46) were modeled onto the solution NMR structure of TatAd from B. subtilis (Hu et al., 2010) PBD: 2L16. The Tha4 conformational shift and how the thylakoid membrane is proposed to compress by hydrophobic mismatch and during generation of the PMF is shown with generic cartoon lipids. Alignment structures were created using Modeller (Sali and Blundell, 1993) in UCSF Chimera (Pettersen et al., 2004). B Proposed interaction between Tha4 TMH (pink) and cpTatC (blue) in the active translocase (Aldridge et al., 2014). cpTatC structural model by homology alignment between the final 230 residues from P. sativum (Uniprot: Q94G17) and the TatC crystal structure from A. aeolicus (Ramasamy et al., 2013) PDB: 4HTT. Hcf106 TMH (green) was modeled on the TatB NMR structure from E. coli (Zhang et al., 2014a) PDB: 2MI2. Alignment structures were created using Modeller (Sali and Blundell, 1993) in UCSF Chimera (Pettersen et al., 2004). C Multiple sequence alignment and sequence logo plot of Tha4 from garden pea (Uniprot: Q9XH46), Arabidopsis (Uniprot: Q9LKU2), maize (Uniprot: Q9XFJ8), and rice (Uniprot: Q75GK3) generated using the MUSCLE alignment algorithm in MegAlign Pro™ (Version 15.3.0. DNASTAR. Madison, WI). Amino acids are color coded according to side chain chemistry (yellow = aromatic, red = acidic, blue = basic, nonpolar = orange, and green = polar).

3.3.2 Synthesis and in vitro translation of Tha4 variants, DT23, cpTatCaaa L231C, and (V-20F)tOE17 Single and double cysteine substituted Tha4 E10/A/D were generated by primer-based site directed mutagenesis using the Phusion polymerase (NE Biolabs) according to the manufacturer’s instruction. Each Tha4 variant was confirmed by sequencing. In vitro translation of Tha4 variants, cpTatCaaa L231C, and (V-20F)tOE17 (a modified signal peptide variant of oxygen evolving complex protein 17 kDa with a phenylalanine substituted for the valine 20 residues ahead of the initial amino acid of mature OE17) was carried out using the Promega Wheat Germ Extract translation kit from in vitro transcribed, capped mRNA with or without [3H]-leucine (Perkin-Elmer or Moravek) as described previously (Dabney-Smith and Cline, 2009). After 1 hr incubation, in vitro translation reactions were diluted with an equal volume of 60 mM leucine in 2x IB and kept on ice until use. Precursor translation reactions were further diluted with an equal volume of clarified stromal extract to a final dilution of 1:4.

3.3.3 Overexpression and purification of KKtOE17His6 Briefly, chemically competent BL21(DE3) codon plus E. coli (NEBiolabs) were transformed with a pET-23a+ expression plasmid containing the coding sequence for the intermediate form of OE17 from maize where the RR motif was substituted with twin lysines (KK) with a C-terminal hexahistidine (His6) tag. The pET-23a+ vector with KKtOE17His6 was a generous gift from Dr. Steven Theg (UC Davis). After overnight growth on lysogeny broth (LB) agar plates with research grade ampicillin (amp, 150 μg/mL) selectivity at 37°C, individual colonies were isolated and screened for protein overexpression LB with amp selectivity (150 μg/mL) in 5 mL cultures. KKtOE17His6 expression was induced by the addition of isopropyl β-D-1-thiogalactopyranoside (IPTG) to a final concentration of 1 mM. Initial expression was continued for 3 hr at 37°C in a shaking incubator. After confirming protein expression by SDS-PAGE and Coomassie blue staining, large scale overexpression was carried out in 2 L baffled flasks containing 1 L terrific broth (TB) with amp selectivity. Cultures were grown at 37°C until an OD600 nm = ~0.6-0.8 was reached. Protein expression in 1 L cultures was induced with IPTG at a final concentration of 1 mM. KKtOE17His6 expression was carried out for 3 hr at 37°C in a shaking incubator. Following expression, the bacteria were collected in cell pellets by centrifugation at 3700 x g for 30 minutes and stored at -80°C until lysis. The cell paste collected from ~500 mL of induced culture was resuspended in 30 mL Ni-NTA resin binding buffer (20 mM Na3PO4, pH 7.9, 500 mM NaCl, 5 mM imidazole, 1 mM PMSF) and cells were lysed by two passes through a French cell press at 20000 psi. The cell lysate was clarified by centrifugation at ~40000 x g for 30 minutes at 4°C. The clarified supernatant was then passed through a gravity flow column with ~5 mL Ni-NTA resin (GoldBio) bed equilibrated with Ni-NTA binding buffer. The flow-through fraction was collected, and the column was washed with 10 column volumes (CVs) of Ni-NTA wash buffer (20 mM Na3PO4, pH 7.9, 500 mM NaCl, 60 mM imidazole). Bound KKtOE17His6 was eluted from the column matrix in 5 separate 1 mL aliquots with Ni- NTA elution buffer (20 mM Na3PO4, pH 7.9, 500 mM NaCl, 500 mM imidazole). Each collected fraction was tested for protein content by SDS-PAGE. Elution fractions were pooled and clarified as before to remove any residual insoluble material. The clarified elution sample was then exchanged into phosphate buffered saline (PBS, 137 mM

NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4), and concentrated by spin filtration in a 3000 MWCO Amicon® Ultra-15 centrifugal filter (Millipore Sigma). The final concentration of KKtOE17His6 was determined using a using a BCA assay kit (Pierce) on a BioMate UV/Vis spectrophotometer according to manufacturer’s instruction.

3.3.4 Functional replacement of endogenous Tha4 and complementation of cpTAT The functionality of Tha4 E10A glutamate and aspartate TMH variants was tested using an in vitro complementation assay as described previously (Dabney-Smith et al., 2003). Briefly, endogenous Tha4 was inhibited in isolated thylakoids (1 mg/mL) by treatment with purified anti-Tha4 immunoglobulin G (IgG) followed treatment with protein A from Staphylococcus aureus (Sigma-Aldrich). In vitro translated Tha4 variants were integrated into αTha4 treated thylakoids by incubation for 20 min at 15°C. Thylakoids were then washed and recovered to test for transport efficiency. In vitro translated DT23 (oxygen evolution protein 23 kDa with a modified signal peptide) was used as precursor protein to test transport by the cpTat system under illumination (~100 μmol/m2/s) at 15°C for 15 min. Transport reactions were arrested by incubation on ice. Thylakoids were recovered and incubated with thermolysin to degrade non-transported DT23. Protein samples were then analyzed by SDS-PAGE and gel fluorography (Cline, 1986). Briefly, SDS-PAGE gels were prepared for fluorography by internal deposition of 2,5- diphenyloxazole (ACROS) in DMSO (ThermoFisher) and dried on filter paper followed by exposure to X-ray film (Carestream Health)

3.3.5 N-ethylmaleimide (NEM) blocking of endogenous cysteine residues Isolated thylakoids were suspended in 1x IB with 0.5 mM DTT and incubated on ice for 15 min before treatment with 2.5 mM NEM. NEM was freshly prepared in 95% EtOH prior to each experiment. NEM treatment was quenched with 3x volumes of 10 mM DTT in 1x IB. Thylakoids were recovered by centrifugation and resuspended to the initial volume with IBM.

3.3.6 Oxidative cross-linking between dual cysteine substituted variants In vitro translated dual cysteine Tha4 variants were integrated into NEM-treated thylakoids by incubation at 15°C for 20 min. Double cys-substituted Tha4 integrated thylakoids were then recovered by centrifugation and washed with excess 1x IBM. Individual crosslinking reactions contained 50 μM ATP, 0.1 mM DTT, and 10 μM methyl viologen in IB, 3.3 mM MgCl2. Reactions in which the PMF was dissipated also contained 0.5 μM nigericin and 1 μM valinomycin. Urea containing crosslinking reactions had the addition of 300 mM urea in 1x IB. The transport reaction was initiated by the addition of either 15 μL of in vitro translated (V-20F)tOE17, KKtOE17His6 (1.5 μM final concentration), or 1x IB as a control and the reactions were incubated at 15°C in a circulating water bath with ~100 μmol/m2/s white light illumination. After 5 min, oxidative crosslinking was induced by the addition of 0.25 mM copper phenanthroline (Dabney- Smith et al., 2003). The cross-linking reaction was continued for 5 min followed in the water bath. The reaction was halted and quenched by the addition 1x IB, 14 mM EDTA and 50 mM NEM. Thylakoids were recovered by centrifugation and normalized to equal chlorophyll concentrations according to Arnon (1949) by addition of non-reducing

sample solubilization buffer (2x: 100 mM Tris-HCl, pH 6.8, 5 mM EDTA, 5% SDS, 30% glycerol, and 8 M urea). Samples were analyzed by 5-13.5% gradient Tris-Tricine SDS- PAGE and gel fluorography, as described above.

3.3.7 Blue Native PAGE and western blot analysis BN-PAGE was carried out as described previously (Cline and Mori, 2001; Schagger and von Jagow, 1991). Briefly, Tha4 variants were integrated into isolated thylakoids and treated to the same conditions as the oligomerization assays as described without the addition of oxidant. After completion of the transport reaction, Tha4 variant integrated thylakoids were solubilized by the addition of digitonin to a final concentration of 0.5% in solubilization buffer (25 mM BisTris-HCl, pH 7, 20% glycerol, 5% Serva blue G). Solubilization was carried out by incubation at 4°C for 1 hr rotating end over end. The soluble fractions following digitonin solubilization were recovered after ultracentrifugation at 100000 x g for 30 min. Soluble fraction samples were mixed with 0.1 volumes of BN-PAGE sample buffer (100 mM BisTris-HCl, pH 7.0, 0.5 M 6-amino-n- caproic acid, 30% glycerol) and resolved on a 5-13.5% acrylamide gradient native gel. Ferritin, bovine serum albumin, and β-amylase were used as the molecular weight standard proteins for mass comparison. BN-PAGE gels were then prepared for gel fluorography as described previously. Western blot analysis of BN-PAGE was carried out by transferring resolved proteins to nitrocellulose blotting membranes (Amersham). After transferring BN-PAGE results, nitrocellulose membranes were blocked with 5% non-fat dry milk dissolved in 1x Tris-buffered saline with 0.1% Tween 20 (TBS-T, 50 mM Tris-HCl, pH 7.6, 150 mM NaCl, 0.1% Tween 20). Primary antibody treatment was with 1:10000 dilution serum containing anti-cpTatC antibodies in 5% milk in 1x TBS-T from rabbit. Primary antibody decorated membranes were washed with 1x TBS-T and incubated with the secondary antibody, goat-antirabbit conjugated to horse radish peroxidase (Bio-Rad) at 1:20000 titer in 5% milk in 1x TBS (lacking Tween 20). Chemiluminescence was generated by incubation of primary and secondary antibody decorated membranes with Clarity Western ECL Substrate (Bio-Rad). Image was captured at 15 min exposure in UVP multi-user imaging cabinet and camera.

3.3.8 Alkaline extraction of thylakoid membrane integrated Tha4 variants Alkaline extraction assays were carried out as described previously (Dabney-Smith et al., 2003). Briefly, Tha4 E10/A/D and Tha4 E10/A/D transmembrane domain double cysteine variants were integrated into thylakoids as previously described above. Each in vitro translated Tha4 variant was diluted 1:6 in 1 x IB (final volume 120 μL) and incubated with 1 mg/mL thylakoids in 1 x IBM with 1 mM DTT. Tha4 variants were integrated into thylakoid membranes for 15 min at 15°C in a water bath and the thylakoid membranes were recovered by centrifugation (3200 x g, 8 min, 4°C). Thylakoids were then suspended in 275 μL of IBM and divided into 5 x 50 μL aliquots for each reaction condition. One aliquot was treated as the integration control and pelleted as before by centrifugation and kept on ice. Two aliquots were also pelleted for the alkaline extraction process. Each of these aliquots were resuspended in 200 μL of either freshly prepared 0.1 M NaOH or 0.2 M NaCO3 buffer. These samples were then incubated on ice for 1 hr. Following the alkaline buffer extraction process, extracted thylakoid membranes were recovered by ultracentrifugation (100000 x g, 15 min, 4°C)

with a TLA-120 rotor in an Optima Max-XP Ultra benchtop centrifuge (Beckman- Coulter). The final two aliquots of each variant were treated or mock-treated with thermolysin in 10 mM CaCl2 buffer. Thermolysin (5 mg/mL in 10 mM CaCl2) was added to give a final concentration of 0.1 mg/mL. Mock-treated samples included the addition of equivalent CaCl2 without the protease. Lysis and mock-lysis reactions were carried out for 40 min with end-over-end rotation at 4°C. After the incubation period, lysis and mock-lysis reactions were quenched by addition of 750 μL of 14 mM Na2EDTA in 1 x IB. Thylakoids were recovered by centrifugation as before and washed with 500 μL of the same buffer. Following thermolysin or alkaline buffer treatment, thylakoid membranes were recovered and suspended in 20 μL 5 mM Na2EDTA in 1 x IB and 20 μL 2 x SSB [(0.1 M Tris-HCl pH 6.8, 10% (v/v) β-mercaptoethanol, 5% (w/v) SDS, 30% (v/v) glycerol, 0.1% (w/v) bromophenol blue]. Samples were resolved by 12.5% acrylamide SDS-PAGE. SDS-PAGE gels were then treated as mentioned above for x-ray film fluorography.

3.3.9 Import of cpTatCaaa L231C into chloroplasts and crosslinking between single cysteine Tha4 variants in thylakoid membranes In vitro translated cpTatCaaa L231C (the aaa cpTatC variant has had its endogenous cysteine residues substituted with alanine) was imported in chloroplasts suspended in 1x IB, 5 mM Mg-ATP, and 5 mM DTT in a circulating water bath at 25°C with ~100 μmol/m2/s white light illumination for 40 mins. Intact chloroplasts were recovered by centrifugation through a 35% Percoll cushion and washed with excess 1x IB, 10 mM MgCl2. Washed chloroplasts were lysed as described above. Single cysteine Tha4 variants were integrated into cpTatCaaa L231C imported thylakoids as described above. cpTatCaaa L231C and Tha4 variant integrated thylakoids were used in crosslinking assays with the same buffers and additives as described for the oligomerization assays above. Preincubation reactions were initiated by the addition of precursor or IB and stored on ice in the dark for 10 min, warmed to 15°C in the dark for 3 min, and placed in an illuminated water bath at 15°C for 3 mins. Crosslinking was initiated by the addition of copper phenanthroline as described above and continued for 5 mins. Reactions were terminated by the addition of NEM and 1x IB, 14 mM EDTA as described previously. Non-preincubated samples were added directly to the light bath and crosslinking was carried out exactly as for the oligomerization assays as described above. Individual reactions were normalized for chlorophyll content with non-reducing 2x SSB as before. Aliquots of each reaction were reduced by addition of β- mercaptoehtanol to 5% total. Reduced and non-reduced samples were then analyzed by SDS-PAGE and gel fluorography.

3.4 Results 3.4.1 Tha4 from higher plants has a conserved glutamate at the 10th position in its primary sequence and is in the hydrophobic core of the thylakoid membrane Nearly all TatA family proteins (Tha4 and Hcf106 included) have a polar amino acid in their TMH (Berks, 2015; Cline, 2015; New et al., 2018). In garden pea and other higher plants such as rice, Arabidopsis, and maize, the polar Tha4 TMH residue is a glutamate (Figure 3.1C). Prior biochemical studies showed that optimal Tha4 function requires a glutamate residue at the 10th position (E10) (Dabney-Smith et al., 2003; Fincher et al., 2003). Precursor transport was abolished in functional complementation assays when Tha4 E10 was changed to either alanine (Tha4 E10A) or glutamine (Tha4 E10Q) (Dabney-Smith et al., 2003). However, when E10 was substituted with an aspartate (Tha4 E10D), substrate proteins were transported by cpTAT (Dabney-Smith et al., 2003), suggesting that this transmembrane glutamate (and possibly aspartate) in Tha4 might function as a sensor for the presence of the pH gradient (proton motive force, PMF) formed across the thylakoid membrane previously demonstrated to be required for active cpTAT transport; recently reviewed (Berks, 2015; Cline, 2015; Hamsanathan and Musser, 2018; New et al., 2018). Our reasoning was that if the Tha4 TMH glutamate functions as a PMF sensor, any changes to its location relative to the of the face of the thylakoid membrane bilayer in the acidified lumen would impact transport function.

3.4.2 Sequential glutamate substitutions in the TMH of Tha4 E10A fail to complement loss of function in αTha4 IgG treated thylakoids In order to better understand the local residue environment of the Tha4 TMH glutamate, we constructed a helical wheel projection (Figure 3.2A) and a model of the transmembrane helix using Modeller in UCSF Chimera (Figure 3.2B) (Pettersen et al., 2004; Sali and Blundell, 1993). To test our hypothesis that the proximity of this glutamate to the thylakoid lumen impacts Tha4 function, we generated several Tha4 variants containing an alanine for glutamate substitution at the 10th position (Tha4 E10A). We then added sequential glutamate or aspartate substitutions in the TMH towards the N-terminus of the protein. First, we made glutamate substitutions in the E10A background at positions P9, V8, G7, or L6 (P9E, V8E, G7E, and L6E). We screened these Tha4 variants for their ability to restore transport of a precursor in a thylakoid membrane biochemical complementation assay (Figure 3.2C) (Dabney-Smith et al., 2003). The precursor, DT23, contains a modified signal peptide linked to the 23 kDa mature domain of the oxygen-evolving complex of photosystem II protein (OE23, aka PsbP) and enhances transport efficiency of the precursor relative to wild type (Henry et al., 1997). Transport of precursor is demonstrated by the appearance of a lower molecular weight band corresponding to mature OE23 (mOE23) as analyzed by SDS-PAGE and fluorography (Figure 3.2C). Additionally, thylakoids were treated with an external protease, thermolysin, to digest non-transported precursor leaving behind only mOE23 that was protected in the lumen (Figure 3.2C). Transport occurred in untreated, isolated thylakoid membranes, while transport was prevented in thylakoids treated with αTha4 IgGs (Figure 3.2C, lanes 2-3 vs. 4-5). Transport was restored, however, if we integrated in vitro translated wild type Tha4 into the thylakoids prior to addition of precursor (Figure 3.2C, lanes 7-8). As expected and previously shown, Tha4

E10A was unable to restore transport (Figure 3.2C, lanes 10-11) (Dabney-Smith et al., 2003). The substitution of glutamate into the TMH of Tha4 E10A at the 9th through 6th positions did not restore transport of DT23 (Figure 3.2C, lanes 14, 17, 20, 23).

Figure 3.2 Glutamate substitutions in the TMH of Tha4 variant E10A are unable to complement loss of cpTAT function. A Helical wheel projection of mature Tha4 residues 2-18 with Met at position 1 (ProteanTM. Version 15.3. DNASTAR. Madison, WI). Aromatic residues are shown in blue, non-polar are black, non-charged in green, and acidic in red. B Tha4 N-terminal transmembrane helix residues (pink) including the glutamate (blue) and glycine (gray) generated in USCF Chimera (Pettersen et al., 2004) as in Figure 3.1A. C Fluorography results of glutamate substituted Tha4 E10A variants. Radiolabeled DT23 (lane 1) was the precursor protein used for transport function assay; maturation of DT23 to mOE23 following successful transport is shown by a decrease in apparent molecular mass. Isolated thylakoid (lanes 2-3) were used as a positive transport control while the rest of the reactions were conducted with αTha4 IgG treated membranes (lanes 4-5, 7-8, 10-11, 13-14, 16-17, 19-20, 22- 23). IVT corresponds to in vitro translated (3H)Tha4 variants before integration in thylakoid. Bands on the gel appearing below the size of recombinant Tha4 are likely proteolytic fragments following thermolysin treatment. Complementation assays were carried out as described in Materials and Methods and analyzed by 12.5% acrylamide Tris-glycine SDS-PAGE under reducing conditions.

3.4.3 Aspartate substitutions in the TMH of Tha4 E10A variant weakly complement loss of function in αTha4 IgG treated thylakoids Previous work had shown that substitution of glutamate 10 with an aspartate (Tha4 E10D) was able to weakly restore transport (Dabney-Smith et al., 2003). Thus, we asked whether Tha4 function was linked to the acidic character of glutamate or the specific position of the residue in the TMH. To tease apart these two contributions, we performed the complementation assays with Tha4 E10A variants containing aspartate substitutions sequentially along the TMH from positions 8-4. Positive (Figure 3.3, lanes 2-3) and negative (Figure 3.3, lanes 4-5) controls for transport indicated functional thylakoids. Integration of wild type Tha4 was able to restore transport of precursor and Tha4 E10A was unable to complement cpTAT (Figure 3.3, lanes 6-11). A band

corresponding to mOE23 was present in the Tha4 E10A integrated sample but was degraded by thermolysin treatment indicating that it wasn’t completely transported into and protected by intact thylakoid membranes (Figure 3.3, compare lane 10 with 11). In our work and as shown previously, Tha4 E10D was able to complement cpTAT transport of DT23 in αTha4 treated thylakoids (Dabney-Smith et al., 2003) albeit at a decreased efficiency when compared to integrated wild type Tha4 and isolated thylakoids (Figure 3.3, compare lanes 13-14 with 1-2 & 7-8). Intriguingly, when an aspartate was substituted into the 8th, 7th, 6th, and 4th positions in the Tha4 E10A TMH, transport was weakly restored (Figure 3.3, lanes 16-17, 19-20, 22-23, 25-26, 28-29). Aspartate substitution in the 5th position in the Tha4 E10A TMH did not restore transport of precursor (Figure 3.3, lanes 16-17).

Figure 3.3 Aspartate substitutions in TMH of Tha4 variant E10A mildly complement loss of cpTAT function. (3H)DT23 (lane 1) was the precursor used in transport complementation assays; maturation of DT23 to mOE23 is shown by a decrease in apparent molecular mass. Isolated thylakoid (lanes 2-3) were used as a positive transport control while the rest of the reactions were conducted with αTha4 IgG treated membranes and the corresponding Tha4 E10A aspartate substituted variant (lanes 4-5, 7-8, 10-11, 13-14, 16-17, 19-20, 22-23, 25-26, 28-29). IVT corresponds to in vitro translated (3H)Tha4 variants before integration into thylakoid. Bands on the gel appearing below the size of recombinant Tha4 are likely proteolytic fragments following thermolysin treatment Complementation assays were carried out as described (see Materials and Methods) and analyzed by 12.5% acrylamide Tris-glycine SDS-PAGE under reducing conditions.

3.4.4 The presence of functional precursor influences Tha4 organization more so than PMF in purified thylakoid membranes Part of the role Tha4 has in transport relates to organization at the precursor-bound receptor complex (Dabney-Smith and Cline, 2009). Because some aspartate substitutions in the TMH of Tha4 weakly restored transport, we wondered if these substitutions altered the ability of Tha4 to oligomerize at the receptor complex. To test this, we used a crosslinking strategy with double cysteine substituted Tha4 as described previously (Dabney-Smith and Cline, 2009). Three structural regions of Tha4 (the unstructured carboxy-terminal tail, stroma proximal TMH, and lumen proximal TMH) were chosen to report on oligomerization (Figure 3.4A). The three pairs of cysteine substitutions (C-tail: A65C T78C; stromal TMH: A18C L20C; lumen TMH: V8C P9C) were chosen based on prior work that showed these substitutions do not interfere with transport (Figure 3.4A) (Dabney-Smith and Cline, 2009). We conducted these assays in the presence or absence of precursor and/or dissipation of the PMF by protonophore/ionophore treatment (Figure 3.4B-D). For these assays, we used the

Figure 3.4 Tha4 E10/A/D variant organization in thylakoid membranes as determined by crosslink formation between three separate structural regions. A Primary sequence diagram of Tha4 in a cartoon membrane. Glutamate 10 is highlighted in orange. Cysteine pairs in the TMH and the C-tail regions of Tha4 are color coded: V8C P9C are highlighted in green, A18C L20C in purple, and A65C T78C in blue. B, C, D Crosslinking interactions formed by interactions between Tha4 E10/A/D variants with B C-tail (A65C T78C), C stroma proximal TMH (A18C L20C), and D lumen proximal TMH (V8C P9C) substituted cysteine pairs in the presence or absence of functional precursor (V-20F)tOE17 and PMF (+ nigericin/valinomycin = dissipated PMF). Oligomerization assays were carried out as described in Materials and Methods (3.3.6) and analyzed by 5-15% acrylamide Tris-Tricine SDS-PAGE under non-reducing conditions.

functional precursor (V-20F)tOE17, the 17 kDa subunit of the oxygen-evolving complex, containing a truncated transit peptide with a (V-20F) substitution in the cpTAT signal peptide region (Celedon and Cline, 2012; Gerard and Cline, 2007). In Tha4 E10 and Tha4 E10A, higher order oligomers of Tha4 (equivalent to ~14 monomers) formed by interaction between cysteines placed in the C-tail regardless of the presence of precursor or PMF (Figure 3.4B). The Tha4 E10D A65C T78C variant was also able to form oligomers but they were fainter in appearance than E10/E10A and appeared to lack some of intermediates (Figure 3.4, compare lanes 1-8 with 9-12).

Cysteines placed in the stroma proximal TMH (A18C L20C) showed increased crosslink formation in presence of precursor regardless of the presence of PMF, especially in Tha4 and Tha4 E10A (Figure 3.4C). Tha4 E10A A18C L20C showed a marked increase in band intensity and oligomer count in the presence of precursor that is not inhibited by depletion of PMF (Figure 3.4C, lanes 5-8). Tha4 A18C L20C also had a similar level of oligomer formation when compared to the E10A variant (Figure 3.4C, lanes 1-4 v. 5-8). Tha4 E10D A18C L20C presented a different pattern of oligomer formation than the E10/A variants (Figure 3.4C, lanes 9-12). These oligomers were markedly fainter in appearance and were equal to ~5 monomers in the presence of precursor (Figure 3.4C, compare lanes 1-8 with 9-12).

The final region of Tha4 that we tested for oligomerization potential was the lumen proximal TMH of Tha4 (V8C P9C). This region was of key interest to us because of its proximity to the original position of the transmembrane glutamate (Figure 3.4A). As seen in the oligomerization results for the stroma proximal TMH, each Tha4 V8C P9C variant showed an increase in oligomer formation and band intensity in the presence of precursor, regardless of the presence of PMF (Figure 3.4D). The highest observable oligomer count in each reaction containing precursor was equal to 8 monomers of Tha4 (Figure 3.4D, even lane numbers). Additionally, the Tha4 E10A V8C P9C variant showed much smaller oligomer formation (eq. to 4 monomers) in the absence of precursor regardless of the presence of PMF (Figure 3.4D, lanes 5, 7).

3.4.5 Tha4 oligomer enhancement was also tested in the presence of functional precursor with urea and non-functional precursor We determined that Tha4 oligomer formation was enhanced in the presence of functional precursor (V-20F)tOE17 (Figure 3.4). Prior work showed that Tha4 oligomer formation was enhanced in the presence of recombinant DT23 extracted from inclusion bodies with urea (Dabney-Smith and Cline, 2009) and that the presence of <500 mM urea doesn’t inhibit cpTAT function (Cline et al., 1993). So, we tested whether Tha4 organization was impacted by the presence of 300 mM urea and (V-20F)tOF17 (Figure 3.5). Our results for the C-tail showed that there was moderate enhancement of Tha4 E10 oligomerization in the presence of (V-20F)tOE17 and urea (Figure 3.5A, compare lanes 2, 4 with 1, 3). Oligomer formation in the Tha4 E10A A65C T78C variant was constant regardless of precursor or PMF in the presence of urea presenting up to a 10- mer (Figure 3.5A, lanes 5-8). Finally, interactions between the C-tail of Tha4 E10D were only present up to a dimer (Figure 3.5A, lanes 9-12). Interactions between the stroma proximate TMH were not enhanced in the presence of (V-20F)tOE17 or PMF for

each Tha4 E10/A/D variant when urea was added to reaction (Figure 3.5B). We saw similar results for oligomer formation between lumen proximate TMH of Tha4 E10/A/D variants where no enhancement oligomer formation was present in any condition tested when urea was added to the reaction (Figure 3.5C).

Figure 3.5 Effect of urea and functional precursor on organization of Tha4 E10/A/D variants in thylakoid membranes. A, B, C Oligomerization assays were conducted as before (Figure 3.4B-D) with urea added prior to precursor addition (see Materials and Methods). Crosslinking interactions formed by interactions between Tha4 E10/A/D variants in the A C-tail (A65C T78C), B stroma proximal TMH (A18C L20C), and C lumen proximal TMH (V8C P9C) by substituted cysteine pairs in the presence or absence of PMF (+ nigericin/valinomycin = dissipated PMF) and functional precursor (V- 20F)tOE17. Oligomerization assays were carried out as described in Materials and Methods and analyzed by 5-15% acrylamide Tris-Tricine SDS-PAGE under non-reducing conditions.

In addition to testing precursor preparation, we needed to examine Tha4 oligomer formation in the presence of non-functional precursor. Functional cpTAT precursors have a conserved twin arginine motif (RR) in their signal peptide (Peltier et al., 2002). Substitution of the RR motif with twin lysines (KK) in the signal peptide of cpTAT precursors inhibits transport by the translocase (Gerard and Cline, 2006; Henry et al., 1997). With this in mind, we tested how each Tha4 E10/A/D variant formed oligomers in the presence of KKtOE17, a non-transportable version of (V-20F)tOE17 (Figure 3.6). To begin this process, we expressed and purified KKtOE17 with a C-terminal hexahistidine tag (KKtOE17His6) using Ni-NTA affinity chromatography (Figure 3.6A). After pooling the elution fractions (Figure 3.6A, lanes 5-9), we exchanged the imidazole laden elution buffer with PBS. After a final concentration step, we quantified the amount of purified KKtOE17His6 with a BCA assay (Table 3.1). We then repeated the oligomerization assays for each double cysteine substituted Tha4 E10/A/D variant. Our

results showed that oligomer formation between adjacent Tha4 C-tails is not enhanced in the presence of non-functional precursor in each E10/A/D variant (Figure 3.6B). This was also the case for the stroma proximate (Figure 3.6C) and lumen proximate TMH (Figure 3.6D) in each Tha4 E10/A/D variant as we saw no enhancement of oligomerization in the presence of KKtOE17His6.

Table 3.1 BCA assay determination of KKtOE17His6 concentration after purification, pooling, and buffer exchange Calibration curve: y = 3.267x + 0.1670 Concentrated Absorbance Experimental Actual KKtOE17His6 562 nm concentration concentration (mg/mL) (mg/mL) 0.01 x 0.077 -0.028 Out of curve 0.1 x 0.275 0.0330 0.330 1 x 1.683 0.4641 0.4641

Figure 3.6 Purification of non-functional precursor and its effect on Tha4 E10/A/D variant organization in thylakoid membranes. A 12.5% SDS-PAGE showing our purification of KKtOE17His6 using Ni-NTA affinity chromatography (see Materials and Methods). Lanes 1-2 are the uninduced and induced expression samples. Lanes 3-4 are the flow-through and wash fractions from the column matrix. Lanes 5-9 are the elution fractions from the column matrix. B, C, D Oligomerization assays conducted as before (Figure 3.4B-D) using KKtOE17His6 as a non-transportable precursor (see Materials and Methods). Crosslinking interactions formed by interactions between Tha4 E10/A/D variants in the B C-tail (A65C T78C), C stroma proximal TMH (A18C L20C), and D lumen proximal TMH (V8C P9C) by substituted cysteine pairs in the presence or absence of PMF (+ nigericin/valinomycin = dissipated PMF) and KKtOE17His6. Oligomerization assays were carried out as described in Materials and Methods and analyzed by 5-15% acrylamide Tris-Tricine SDS-PAGE under non-reducing conditions.

3.4.6 BN-PAGE analysis revealed extensive interactions between Tha4 E10/A variants and cpTAT components To better understand how each Tha4 E10/A/D variant formed interactions with Tha4 monomers and the other cpTAT component proteins without disulfide formation, we used BN-PAGE. We integrated cys-free Tha4 E10/A/D variants into thylakoid membranes and applied the same conditions as the oligomerization assays for each (Figure 3.7A). Tha4 E10 associated in an ~250 kDa complex under each condition tested (Figure 3.7A) similar to what has been previously shown (Cline and Mori, 2001). Tha4 E10 also associated with the ~700 kDa complex (Cline and Mori, 2001) shown to previously contain cpTatC and Hcf106 (Figure 3.7A). Tha4 E10A associated with a range of different sized complexes that were only faintly present in the wild type Tha4 reactions such as the ~250 kDa and ~700 kDa complexes (Figure 3.7A). Finally, Tha4 E10D weakly associated with the ~250 kDa and ~700 kDa complexes and appeared as faint bands under each condition (Figure 3.7A). We also conducted an immunoblot analysis probing for cpTatC with the BN-PAGE samples (Figure 3.7B). In the absence of precursor, cpTatC primarily populates a lower molecular weight complex as evidenced (Figure 3.7B). cpTatC also associated with a ~700 kDa complex and a ~550 kDa complex in the absence of precursor (Figure 3.7B). Upon the addition of precursor, cpTatC shifts to be mostly in the ~700 and ~550 kDa complexes (Figure 3.7B). The increased association of cpTatC with the larger complexes was only impacted by the presence of precursor protein and not the depletion of the PMF (Figure 3.7B) in agreement with previous work that showed energy-independent binding of precursor to the receptor complex (Ma and Cline, 2000).

Figure 3.7 Interactions between cys-free Tha4 E10/A/D variants examined by Blue native PAGE. A Blue native PAGE and gel fluorography showing association of Tha4 variants following extraction from thylakoid membrane by 0.5% digitonin. B Western blot analysis of in vitro translated Tha4- integrated thylakoid membranes probed with cpTatC antibodies (see Material and Methods). 3.4.7 The stability of thylakoid membrane integrated cys-free and double cysteine Tha4 E10/A/D variants was determined by alkaline extraction The results of our oligomerization assays showed that Tha4 E10A is more likely to interact with other Tha4 E10A monomers. These observations led us to wonder how these alanine and cysteine substitutions impacted Tha4 stability in the thylakoid membrane and calculated TMH hydrophobicity. We accomplished this aim by determining how resistant each Tha4 variant was to alkaline buffers by “harsh” (0.1 M NaOH) and “mild” (0.2 M NaCO3) extraction after integration into thylakoids.

These assays allow us to correlate resistance to alkaline extraction to relative stability of Tha4 variants (Dabney-Smith et al., 2003). First, we tested cys-free Tha4 E10/A/D variants to determine how substitution of glutamate with alanine or aspartate alters Tha4 stability prior to any impact the double cysteine substitutions may have had (Figure 3.8A-C). As previously shown, Tha4 E10A was much more resistant to extraction from the thylakoid membranes by NaOH as compared to Tha4 E10 and Tha4 E10D (Figure 3.8A-C) (Dabney-Smith et al., 2003). Additionally, we quantified how these substitutions altered Tha4 TMH hydrophobicity with the ExPASy ProtScale tool and Eisenberg et al. hydropathy values (Figure 3.9) (Eisenberg et al., 1984; Wilkins et al., 1999). Substitution of glutamate with alanine in Tha4 increased the hydrophobicity of the lumen proximate TMH (residues 5-14) without changing hydrophobicity of the stroma proximate TMH (residues 15-22) (Figure 3.9A). The Tha4 E10/D V8C P9C variants (lumen proximate TMH) appeared to more readily integrate into thylakoid membranes and were more stable than their cys-free counterparts (Figure 3.8A-F). Tha4 E10A V8C P9C appeared to have a similar level of resistance as Tha4 E10A to 0.1 M NaOH extraction (Figure 3.8B, E). However, the hydrophobicity of the lumen proximate TMH in each Tha4 E10/A/D V8C P9C variant was less than was calculated

Figure 3.8 Cys-free and double cysteine Tha4 E10/A/D variant stability in thylakoid membranes as determined by alkaline extraction assays. Results of alkaline extraction of thylakoid membrane integrated Tha4 E10/A/D (A-C), Tha4 E10/A/D V8C P9C (D-F), and Tha4 E10/A/D A18C L20C (G-I). Each Tha4 variant was in vitro translated (IVT) using wheat germ extract and integrated into thylakoid membranes. An untreated, integrated sample was collected for each variant (U). Alkaline extraction assays used Tha4 integrated thylakoid membranes that were treated with either 0.1 M NaOH (OH) or 0.2 M Na2CO3 (C). Tha4 variant integrated thylakoid membranes were also treated or mock-treated with Thermolysin (T & MT). PF stands for proteolytic fragments. Alkaline extraction assays were carried out as described in Materials and Methods and analyzed by 12.5% acrylamide Tris-glycine SDS-PAGE under reducing conditions.

for their cys-free counterparts and did not decrease the hydrophobicity of the stroma proximate region of the TMH (Figure 3.9A). Tha4 E10/A/D A18C L20C variants were the least resistant to alkaline extraction and had less hydrophobic stroma proximate TMH regions than the other variants (Figure 3.8, 3.9B). The lumen proximate TMH hydrophobicity was not altered by the A18C L20C substitutions (Figure 3.9B).

Figure 3.9 Eisenberg hydrophobicity values of residues 5-30 in cys-free and double cysteine Tha4 E10/A/D variants. Comparison between Tha4 E10/A/D and A Tha4 E10A/D V8C P9C residues 5-30 or B Tha4 E10A/D A18C L20C residues 5-30. The hydrophobicity values were calculated using ExPASy ProtScale with a residue window of n = 9 and a linear weight variation model in which window edge residue weights are 10% relative to the central residue i (i = 100%, i ± 1 = 78%, i ± 2 = 55%, i ± 3 = 33%, and i ± 4 = 10%).

3.4.8 Preliminary data of crosslinking interactions between Tha4 TMH and cpTatC TM4 in the presence and absence of precursor and PMF The interactions between the Tha4 TMH and cpTatC TM4/5 are altered in a precursor transport dependent manner (Aldridge et al., 2014) (Figure 3.1B). This study showed

that the Tha4 TMH forms crosslinks with cpTatCaaa TM4 (endogenous cystines substituted with alanine) that are enhanced in the presence of functional signal peptide linked to the first 12 residues of (V-20F)tOE17 (Aldridge et al., 2014). As Tha4 E10D only moderately restores loss of cpTAT function and Tha4 E10A completely fails to do so (Figures 3.2C, 3.3) (Dabney-Smith et al., 2003), we decided to test whether crosslinking between the TMH of each Tha4 E10/A/D variant and cpTatCaaa TM4 is enhanced in the presence of full-length precursor and PMF (Figure 3.10). The first experiment we conducted was to test the necessity of a preincubation step used by Aldridge et al. to saturate cpTAT receptor complexes with signal peptide/precursor prior to crosslinking Tha4 and cpTatC (Figure 3.10A). Tha4 P9C formed crosslinks with cpTatC L231C in TM4 in the presence and absence of precursor and PMF regardless of preincubation in the dark (Figure 3.10A). These interactions were largely depleted after reduction by β-ME with only faint homodimers of Tha4 P9C present (Figure 3.10A). The second preliminary experiment we completed was testing for interactions between cpTatC TM4 and two additional Tha4 TMH positions, F3C and F4C, as used previously (Aldridge et al., 2014) (Figure 3.10B). Tha4 TMH F3C and P9C readily formed interactions with cpTatC L231C under each condition tested while position F4 showed slight crosslinking in the absence of precursor and minimal interaction in the presence of precursor (Figure 3.10B). Tha4-Tha4 F3C and F4C homodimers and Tha4-cpTatC crosslinks were completely depleted in reducing conditions while faint bands corresponding to Tha4 P9C homodimers remained present as seen previously (Figure 3.10A,B). The final preliminary experiment we conducted was to test for interactions between the F4C in Tha4 E10/A/D variants and cpTatC L231C (Figure 3.10C). We showed that interactions between Tha4 F4C E10/D and cpTatC L231C were much less prevalent than Tha4 F4C E10A under all conditions (Figure 3,10C). However, there was no enhancement of crosslink formation in the presence of precursor or PMF (Figure 3.10C). As before, these interactions were depleted after reduction by β-ME (Figure 3.10C). Although the data presented here in 3.4.10 is promising, much work remains to be finished to validate these preliminary observations.

3.5 Discussion In plants, the TatA component, Tha4, contains a highly conserved glutamate (E10) in the N-terminal TMH. We sought to determine if the specific position of this glutamate in TMH is required for cpTAT function. We also wanted to determine how substitution of glutamate 10 changes Tha4 oligomerization. Prior studies of Tha4 observed a topological shift upon activation (Aldridge et al., 2012). The experimentally determined topology shift of Tha4 from tilted to perpendicular to the membrane normal suggested to us that the protonation state of E10 may play a role in how Tha4 partitions into the membrane or interacts with adjacent Tha4 monomers (Figures 3.1A, 3.7) (Aldridge et al., 2012; Pal et al., 2013). Previously, molecular modelling of TatA oligomers in an E. coli plasma membrane highlighted how the membrane would be compressed by hydrophobic mismatch between the short TatA TMH and surrounding lipids (Rodriguez et al., 2013). The TatA model taken in conjunction with the Tha4 topological shift shown during transport suggested to us that E10 is exposed in the lumen (Figure 3.1A). Given that the protonation state of glutamate is determined by pKa of the side chain γ-carboxyl

Figure 3.10 Preliminary crosslinking data between Tha4 TMH and cpTatC TM4 in thylakoid membranes. The apparent MW of Tha4 is ~11 kDa, Tha4-Tha4 is ~22 kDa, and cpTatC-Tha4 is ~41 kDa A cpTatCaaa L231C-Tha4 P9C crosslinking was tested in the presence or absence of precursor and PMF after a 10 min or 0 min incubation in the dark on ice. B Crosslinking between Tha4 F3C, F4C, and P9C variants and cpTatCaaa L231C in thylakoid after 10’ incubation in the dark. C Crosslinking between Tha4 F4C E10/A/D variants and cpTatCaaa L231C after 10’ incubation in the dark. These crosslinking assays were carried out as described in Materials and Methods. Reduced and non-reduced (+/- 5% β-ME) samples were analyzed with 10% acrylamide Tris-glycine SDS- PAGE.

group and the pH of the surrounding environment, the proximity of E10 to the acidified lumen (~pH 5.8-7) during transport suggests that this residue is likely protonated (Figure 3.1A) (Fitch et al., 2002). The pKa of free glutamic acid is ~4 and experimentally determined to be ~6-9 when in synthetic peptides and model bilayers or when buried in the hydrophobic core of a protein (Panahi and Brooks, 2015; Teixeira et al., 2016). However, this is speculative and a precise measure of the Tha4 γ-carboxylate group pKa is required.

In the presence of PMF shown to be required for Tha4 recruitment to the cpTAT complex, the population of deprotonated to protonated Tha4 could change leading to a subpopulation that undergoes the observed topological shift and organizes at the precursor bound receptor complex (Figure 3.1A) (Aldridge et al., 2014; Aldridge et al., 2012). Concurrently, the charged population of Tha4 E10 would be capable of interacting with glutamine 234 in the fourth TM of cpTatC by hydrogen bonding as speculated previously (Aldridge et al., 2014) (Figure 3.1B). Similar sensing functions have been presented for voltage-gated ion channels which adopt different conformations in response to changes in the PMF (Bocharov et al., 2008; Catterall et al., 2017). Tha4 E10A would be unable to undergo the conformational shift as the replacement of glutamate with alanine caused the TMH to be more hydrophobic/more stable in thylakoids leading to tighter helix-helix interactions between adjacent Tha4 (Figure 3.8, 3.9) and insensitivity to changes in the PMF (Figure 3.7B). We showed that the Tha4 E10A variants have increased oligomerization in the presence of precursor (Figures 3.4, 3.5B) but are unable to complement loss of transport function (Figure 3.2C) as shown previously (Dabney-Smith and Cline, 2009; Dabney-Smith et al., 2003). In addition to increased stability in thylakoid membranes, Tha4 E10A variants would be incapable of forming the proposed hydrogen bond interaction with cpTatC Q234 (Aldridge et al., 2014).

To better explain how changes in the position of Tha4 E10 might be linked to function in cpTAT through sensing the PMF, we searched for restoration of transport function in a non-functional Tha4 E10A by substitution of a glutamate or aspartate into the TMH. We determined that for glutamate, proper position in the TMH is required for cpTAT complementation of function (Figure 3.2C). This is likely due to several factors. Substitution of glutamate with an alanine increases the insensitivity of Tha4 to extraction by Na2CO3, due to the increase in hydrophobicity of the TMH (Dabney-Smith et al., 2003) (Figure 3.8, 3.9). An increase in the stability of Tha4 in the membrane would then prevent the experimentally shown conformational shift of Tha4 (Aldridge et al., 2012). The stability of Tha4 in the thylakoid membrane has been shown to influence the its ability to function in protein transport by cpTAT (Dabney-Smith et al., 2003). Furthermore, increased hydrophobicity of the TMH in E10A variant may alter helix-helix packing between adjacent Tha4 preventing dissociation of monomers from the tetrameric bundles in the thylakoidal free pool ultimately preventing association with precursor bound receptor complexes (Figure 3.11A-B) (Alcock et al., 2016; Aldridge et al., 2014; Habersetzer et al., 2017; Hamsanathan and Musser, 2018). Additionally, tighter packing of adjacent Tha4 E10A would prevent translocation of substrate by preventing proper Tha4 oligomer formation in any of the current molecular models of

(cp)TAT function (Figure 3.11A-B) (Berks, 2015; Hamsanathan and Musser, 2018; New et al., 2018). Our substitution of aspartate at the 4th, 6th, 7th, and 8th residues in Tha4 E10A slightly restored function (Figure 3.3, lanes 16-17, 19-20, 22-23, 25-26, 28-29) while substitution in the 5th position failed to do so (Figure 3.3, lanes 16-17). Only moderate complementation of function was achieved when aspartate was at the 10th position in Tha4 TMH as seen previously (Figure 3.3, lanes 13-14) (Dabney-Smith et al., 2003).

We also tested whether the substitution of Tha4 E10 with alanine or aspartate is linked to Tha4 organization in the thylakoid membrane. The results of our oligomerization assays show that the C-tail of Tha4 E10/A/D variants formed consistent crosslinks with adjacent Tha4 C-tails regardless of the presence of functional precursor or PMF (Figure 3.4B). Our results are different from previous work in which Tha4 oligomerization was shown to be dependent on the presence of PMF and full-length precursor DT23 (OE23 with a dual targeting signal peptide) or the transit peptide of OE17 (tpOE17) (Dabney- Smith and Cline, 2009). In that study, they showed that depletion of PMF in the presence of tpOE17 led to fewer crosslinking interactions between adjacent Tha4 C- tails (Dabney-Smith and Cline, 2009). The Tha4 C-tail protrudes into the chloroplast stroma and is not rigid like the TMH. Thus, this region can sample more conformations in the stroma and form larger oligomers regardless of the presence or absence of precursor and PMF (Figure 3.4B). We also showed an increase in crosslink formation between adjacent stroma proximate (Tha4 A18C L20C) and lumen proximate (V8C P9C) regions of the TMH in the presence of precursor regardless of PMF in each E10/A/D variant (Figure 3.4C-D). This differs from a previous study where crosslink formation between Tha4 E10A V8C P9C was not enhanced by the addition of DT23 (Dabney-Smith and Cline, 2009). So, we considered how differences between precursor preparation might influence our oligomerization assay results. In the previous study, DT23 was recovered from E. coli inclusion bodies using 8 M urea prior to use (Dabney- Smith and Cline, 2009) and our (V-20F)tOE17 was in vitro translated with wheat germ extract and lacked urea (see Materials and Methods) (Figure 3.4B-D). The results of adding urea to our oligomerization assays showed that crosslinking was slightly enhanced between the C-tails of Tha4 E10 (Figure 3.5A, compare lanes 2,4 with 1,3); interactions between each other tested Tha4 variant were not increased (Figure 3.5A, lanes 5-12, Figure 3.5B-C). Our results agree with the previous C-tail interaction data but differ from the lumen proximate TMH data (V8C P9C) (Dabney-Smith and Cline, 2009). However, this could be due to differences in the susceptibility each precursor to urea induced unfolding. In vitro translated DT23 is transported by cpTAT (Figure 3.2) and DT23 purified with urea has been shown to stimulate Tha4 interaction with precursor bound receptor complex (Dabney-Smith et al., 2003). (V-20F)tOE17 is also transported by cpTAT but it has yet to be demonstrated if the addition of urea purified (V-20F)tOE17 leads to Tha4 coimmunoprecipitation with receptor complex. Finally, we tested if a non-functional variant of precursor, KKtOE17His6, enhanced oligomer formation in between each structural region of Tha4 E10/A/D variants. The addition of non-functional precursor didn’t enhance crosslink formation between each Tha4 variant tested in the presence or absence of PMF (Figure 3.6). Our results support the prior observations in which the presence of non-functional precursor, KK-DT23, was unable

to enhance crosslink formation between the C-tail or lumen proximate TMH in both Tha4 E10/A variants (Dabney-Smith and Cline, 2009).

An additional aspect of the Tha4 conformational shift yet to be defined is how Tha4 tetramers (or larger bundles) in the free pool organize into larger oligomers prior to transport (Aldridge et al., 2012; Dabney-Smith and Cline, 2009). The first piece of the puzzle that we inspected was the C-tail. However, previous results suggest that the C- tail appears to only minimally impact Tha4 TMH oligomerization (Dabney-Smith and Cline, 2009). Our results suggest that precursor binding with receptor complex has a much larger impact on Tha4 TMH organization than the presence of the PMF (Figure 3.4). Additionally, Tha4 V8C P9C formed higher order oligomers than Tha4 A18C L20C in the presence of precursor and PMF (Figure 3.4C-D, lane 2 in each). Tha4 E10A TMH interactions were also enhanced in the presence of precursor but were indifferent to the presence of PMF (Figure 3.4C-D, lanes 2, 4 in each). An interpretation of this data is that the helix-helix interactions between adjacent Tha4 E10 rearrange from a “crossed point” interaction to a parallel, side by side organization as Tha4 bundles amass at precursor bound receptor complexes (Figure 3.11C). Our model also shows the shift in preference for interactions between the stroma proximate TMH to the lumen proximate TMH in the presence of precursor (Figure 3.4C-D, 3.11C). However, it remains to be shown if this reorganization occurs prior to or during the Tha4 interaction with the precursor bound receptor complex.

Figure 3.11 Models of Tha4 E10/A/D helix packing and reorganization in the presence of functional precursor. Models of A Tha4 E10/D and B Tha4 E10A packing interactions in thylakoid membrane when functional precursor is present. Tighter packing interactions between Tha4 E10A are shown by increased proximity between adjacent TMH. C Model of Tha4 TMH reorganization in response to the presence of precursor. Tetrameric Tha4 rearrangment from a “crossed point” interaction to parallel bundle interaction. Stroma proximate cysteine residues A18 and L20 are depicted in purple, lumen proximate residues V8 and P9 are depicted in green, and E10 is depicted in orange.

Our data showing how changes in Tha4 TMH hydrophobicity impacts interactions between Tha4 monomers and other cpTAT component proteins are also supported by BN-PAGE analysis (Figure 3.7). Tha4 E10A formed several associations in a range of sizes that were resistant to dissolution by the detergent digitonin while Tha4 and Tha4 E10D associated complexes were much less resistant to detergent extraction than the alanine variant (Figure 3.7A). Despite these differences, each variant was associated with cpTatC in a ~700 kDa complex (Figure 3.7). In support of the oligomerization assay data and the BN-PAGE results, changes in Tha4 stability in the thylakoids were also observed during alkaline extraction of Tha4 variants in the membrane (Figure 3.8). Substitution of glutamate for alanine increased overall Tha4 resistance to alkaline extraction from thylakoid membranes while aspartate substitution decreased this resistance (Figure 3.8A-C). However double cysteine substitutions had different effects on Tha4 E10/A/D stability in thylakoids (Figure 3.8). Tha4 E10/A/D V8C P9C variants were more resistant to alkaline extraction by NaOH and thus had a more stable interaction with the membrane lipids and/or adjacent Tha4 monomers (Figure 3.8D-F). The Tha4 E10/A/D A18C L20C variants were much more susceptible to extraction from the membrane by the “harsh” NaOH and “mild” NaCO3 buffers (Figure 3.8G-I). This is likely due to either changes in Tha4 TMH membrane partitioning depth or stronger/tighter packing interactions between adjacent Tha4 in the thylakoid membrane. The oligomerization assay data suggests that this is more likely due to the latter as Tha4 V8C P9C variants formed higher order oligomer associations with adjacent monomers than Tha4 A18C L20C (Figure 3.4). This scenario is supported by a previously published comparison between the transport complementation efficiency of these two Tha4 double cysteine variants (Dabney-Smith and Cline, 2009). Tha4 V8C P9C was less efficient at complementing loss of transport function in αTha4 IgG treated thylakoid membranes than wild type; Tha4 A18C L20C was nearly twice as efficient (Dabney-Smith and Cline, 2009). If adjacent Tha4 TMH are more tightly packed with each other, then they might be unable to properly organize into the protein conducting element during cpTAT function preventing transport (Alcock et al., 2016; Blummel et al., 2015; Hamsanathan and Musser, 2018; New et al., 2018). Our alkaline extraction assay data might also point to the significance of the Tha4 TMH proline at position 9. Proline residues introduce a kink and twist into transmembrane helices which are necessary for proper function of g-coupled protein receptor complexes (Law et al., 2016) and connexin32 gap junction channels (Brennan et al., 2015). Through replacement of P9 with a cysteine residue, the Tha4 TMH V8C P9C might adopt a straighter conformation leading to tighter packing between adjacent monomers and the observed loss of transport efficiency in complementation assays and increased oligomer formation. This hypothesis needs to be confirmed experimentally.

3.6 Conclusions and Future Directions Through our exploration of amino acid substitutions in the TMH of Tha4, we were able to show that the transmembrane glutamate at position 10 appears to balance TMH hydrophobicity for proper function and organization in cpTAT. When the TMH is more hydrophobic (E10A), Tha4 resistance to alkaline and detergent extraction is markedly increased, transport of precursor is prevented, and enhancement of oligomer formation in the presence of precursor is decreased relative to wild type Tha4 E10. What remains

to be confirmed is how these increased Tha4 TMH packing interactions prevent transport of precursor, specifically, if tighter packing prevents formation of the protein conducting element or changes how Tha4 interacts with other cpTAT component proteins. Although the data presented in 3.4.8 is promising, much work remains to be finished. First, each preliminary experiment presented needs to be repeated for biological replicates. We also need to repeat these experiments with radiolabeled (3H)cpTatC L231C and unlabeled Tha4 to verify the formation of crosslinks between these proteins. Additionally, these interactions would need to be confirmed by more sensitive and direct experimental means such as co-immunoprecipitation assays after crosslink formation. Finally, we need to determine if interactions between the TMH of Tha4 E10/A/D variants and cpTatC TM5 (V270C) occur in the presence of full-length precursor. Tha4 TMH crosslinks do form with cpTatC TM4 and TM5 in the presence of signal peptides linked to truncated mature domain peptides but these constructs are not transported nor are they able to dissociate after binding with the receptor complex (Aldridge et al., 2014). We also used Tha4 P9C in these preliminary studies. As mentioned above, the TMH proline likely puts a kink in the helix so another route of study is to structurally characterize Tha4 variants lacking the proline and then link its presence to cpTAT function.

A second avenue of investigation is to better define how the Tha4 C-tail contributes to Tha4/cpTAT function. Oligomerization of adjacent Tha4 TMH and Tha4/TatA function occurs without the C-tail present (Dabney-Smith and Cline, 2009; Lee et al., 2002). Although this motif is not required for precursor transport, crosslinking interactions have been discovered between the Tha4 C-tail (A65C) and a solvent exposed OE17 residue (S84C) (Pal et al., 2013). Prior work has also determined that Tha4 C-tail oligomer formation requires functional receptor complexes (Dabney-Smith and Cline, 2009). Currently, we speculate that the C-tails of adjacent Tha4 might form an unstructured, proteinaceous mesh that prevents the precursor mature domain from diffusing back into the stroma ultimately directing transport across the membrane (Pal et al., 2013). Any formation of a Tha4 C-tail meshwork will need to be confirmed.

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Chapter 4: Conclusions

4.1 Conclusions For proper function, chloroplasts require expression of both nuclear and organellar genes which encode for nearly 3000 different proteins, most of which are translated in the cytosol of the plant cell (Peltier et al., 2002; Peltier et al., 2000). Any protein destined for the chloroplast has an N-terminal, cleavable transit peptide which is responsible for proper targeting and additional routing to various compartments within the organelle (Leister and Schneider, 2003). These proteins find initial entry into the chloroplast by the action of translocons on the outer and inner chloroplast membranes (TOC and TIC, respectively) (Paila et al., 2015; Thomson et al., 2020). After being imported by TOC/TIC, some of these proteins destined for the lumen or thylakoid membrane are directed to and actively transported by one of two pathways, the chloroplast secretory system (cpSec) (Albiniak et al., 2012) or the chloroplast twin arginine transport (cpTAT) system (New et al., 2018). cpTAT is so named because of the conserved twin arginine (RR) motif located in cpTAT targeted transit peptides (Peltier et al., 2002). It is important to understand cpTAT function because its substrate proteins are required for thylakoid biogenesis and maintenance of functional photosystems (Jarvi et al., 2013; Leister and Schneider, 2003). cpTAT is unique in that it transports fully folded substrate proteins using only the energy stored in the proton motive force (PMF) through the concerted efforts of three membrane bound component proteins: cpTatC, Hcf106, and Tha4 (Alder and Theg, 2003; Braun et al., 2007; Cline and Mori, 2001; New et al., 2018). Several prokaryotes also have TAT systems in their plasma membranes that are homologous to cpTAT in function (Berks, 2015; New et al., 2018; Palmer and Berks, 2012). However, these prokaryotic TAT systems differ from cpTAT primarily in their component protein primary sequences, stoichiometry, evolutionary history, and functional requirement (some TAT systems lack component proteins that are present in others) (Berks, 2015; New et al., 2018; Palmer and Berks, 2012). Although great strides have been made in the study of (cp)TAT, there are questions that remain unanswered including 1) how does (cp)TAT harness the energy of the PMF, 2) how does Tha4/TatA facilitate precursor transport, and 3) what, if any, are the differences between the experimentally determined structures of TAT and cpTAT component proteins (Hamsanathan and Musser, 2018; New et al., 2018). The answers to these questions will help us to more accurately model the specific mechanistic details of cpTAT function and better define the differences between the chloroplast and prokaryote systems. I approached answering these questions by designing separate studies of the cpTAT component proteins Hcf106 and Tha4.

The aim of my first project was to develop a purification scheme for full-length Hcf106 and use biophysical techniques to characterize its structure in and interactions with the lipids of a mimetic membrane system. Prior structural characterization studies of TAT and cpTAT proteins were carried out with truncated or small fragmented sections of TatA and TatB/Hcf106 (Hu et al., 2010; Zhang et al., 2013; Zhang et al., 2014a; Zhang et al., 2014b). So, to characterize full-length Hcf106, I needed to develop a purification scheme capable of producing adequate quantities of homogenously pure protein (Chapter 2). To overcome limitations in Hcf106 solubility and for ease of purification, we created a fusion of an N-terminal maltose binding protein (MBP) linked to full-length Hcf106 with a tobacco etch virus protease (TEVp) recognition sequence (Lebendiker

and Danieli, 2011; Tropea et al., 2009). We chose TEVp because it is readily purified from E. coli (Tropea et al., 2009). Following expression and purification of protease and fusion proteins, we screened several reaction conditions to optimize the proteolytic efficiency of TEVp with MBP-Hcf106. We were able to coax the proteolysis of our fusion protein further towards completion by increasing duration and temperature while also increasing the concentration of TEVp in the reaction. However, we were unable to optimize the reaction for complete cleavage of Hcf106 from MBP. Although the addition of detergents or urea wasn’t enough to push the reaction to completion, some additives further improved the efficiency of TEVp. After optimizing the reaction as much as able, we were able to remove TEVp from the reaction mixture with magnetized Ni-NTA beads. Additionally, we were able to separate free MBP from free Hcf106 and noncleaved fusion protein by size exclusion chromatography. While this was a promising result, Hcf106 and MBP-Hcf106 coeluted after FPLC in multiple trials with different detergents and urea. This was likely due to Hcf106 TMH (and APH) domains preferentially forming tight inter-helix packing or hydrophobic interactions instead of being solubilized by detergent. Our work did show that TEVp efficiency is severely impacted by the detergents we used in this study with only one exception, CHAPS. It is my opinion that future work with MBP-TEVp-Hcf106 should retest purification with CHAPS. It was the only detergent that improved TEVp reaction efficiency relative to without. In addition to CHAPS, future studies could use other proteases such as enterokinase or thrombin as they are both compatible with the detergents previously used to purify TatB (Vergis and Wiener, 2011; Zhang et al., 2014b). Alternatively, newly developed TEVp variants that have increased proteolytic efficiency for soluble and affinity matrix bound substrates are now available (Sanchez and Ting, 2020; Zhu et al., 2017). While developing a purification scheme for full-length Hcf106, I also studied the role of the membrane embedded, Tha4 transmembrane helix glutamate in cpTAT function.

The aim of my second project was to better understand how a transmembrane helix glutamate impacts Tha4 function and organization in the cpTAT system. Previously, substitution of this residue with an aspartate (Tha4 E10D) restored transport of precursor while substitution with alanine (Tha4 E10A) was unable to restore loss of transport function in isolated thylakoids (Dabney-Smith et al., 2003). It was also shown previously that the E10A substitution disrupted Tha4 oligomer formation (Dabney-Smith and Cline, 2009) but Tha4 E10D organization wasn’t examined. So, we used the same complementation of function and oligomer formation assays used previously with Tha4 E10/A/D variants (Chapter 3). We showed that aspartate substitutions in the TMH of Tha4 variant E10A were weakly able to restore loss of function and glutamate substitutions were unable to restore any cpTAT function. We also examined and compared how self-interactions between three structural motifs in each Tha4 E10/A/D variant changed in the presence or absence of precursor and PMF. Ultimately, we showed that the transmembrane glutamate at position 10 is required for proper Tha4 function and optimal organization in the presence of precursor. We also found that when the TMH is more hydrophobic (E10A), Tha4 stability in the thylakoid membrane is increased, its ability to facilitate precursor transport is diminished, and its organization into higher order oligomers in the presence of precursor is decreased. One piece of the

puzzle that remains to be solved is how Tha4 E10/A/D variant interactions with the individual receptor complex proteins change in absence or presence precursor and PMF. Specifically, it is of key interest to determine if Tha4 E10A/D variant interactions with cpTatC TM4 are enhanced during active transport as described previously (Aldridge et al., 2014). If Tha4 E10A is able to consistently form this interaction with cpTatC TM4 in the presence and absence of precursor and PMF, then one can envision a model of cpTAT function where Tha4 E10A enters an annulus of heterotrimeric cpTAT receptor complexes, binds with cpTatC TM4 (Figure 1.4, 1.5), and acts as a nucleation point for additional Tha4 E10A that pack too tightly to facilitate transport. If this interaction is only enhanced in the presence of precursor, the precursor binding step would be implicated as the trigger for Tha4 E10A entry into the active translocase where the increased packing prevents transport. Finally, a lack of cpTatC TM4-Tha4 E10A TMH interactions that are not enhanced in the presence of precursor suggests a model in which Tha4 E10A are packed too tightly to reorganize at and enter a precursor bound receptor complex. In addition to clarifying how Tha4 E10A disrupts transport, our proposed model of Tha4 TMH rearrangement (Figure 3.11C) in presence of precursor needs to be confirmed by biophysical characterization experiments and molecular modeling simulations.

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