Probing the Structural Dynamics, Conformational Change, And

Home , Electron crystallography

MIAMI UNIVERSITY The Graduate School

Certificate for Approving the Dissertation

We hereby approve the Dissertation

Tanbir Ahammad

Candidate for the Degree

DOCTOR OF PHILOSOPHY

______Dr. Gary A. Lorigan, Director

______Dr. Carole Dabney-Smith, Chair

______Dr. Rick Page, Reader

______Dr. David L. Tierney, Reader

______Dr. Paul Urayama, Graduate School Representative

ABSTRACT

PROBING THE STRUCTURAL DYNAMICS, CONFORMATIONAL CHANGE, AND TOPOLOGY OF PINHOLIN S21, A BACTERIOPHAGE LYTIC PROTEIN, USING ELECTRON PARAMAGNETIC RESONANCE SPECTROSCOPY

Tanbir Ahammad

Bacteriophages have evolved an efficient, protein-mediated host cell lysis mechanism that terminates the infection cycle and facilitates the release of progeny virions at an optimal time. Among the lytic proteins, holin controls the first and rate-limiting step of host cell lysis by permeabilizing the inner membrane at an allele-specific time and concentration. Recently, a prototype holin called pinholin has been reported which makes nanoscale holes that are too small for the passage of endolysin. However, the holes formed by pinholin dissipate the cytoplasmic membrane potential leading to the release and activation of membrane-tethered signal anchor release endolysins which are already exported to the periplasm. Pinholin is the evolutionary ancestor of holin, but was discovered more recently and, as such, has not been fully studied in the literature. Of all the pinholin systems, S21 from lambdoid phage 21 is one of the most well-known. Pinholin S21 consists of two holin proteins: the active pinholin S2168 and the inactive 21 antipinholin S 68IRS. Each of these proteins have a short N-terminal domain followed by two transmembrane domains connected by a short loop and terminate in a long, positively charged C-terminal region. However, the precise structural details of the proteins in this system is not well understood. The works presented in this dissertation were carried out to structurally characterize the proteins of pinholin S21 using biophysical techniques including state-of-the-art electron paramagnetic resonance (EPR) spectroscopic methods. Continuous Wave (CW) EPR line shape analysis and power saturation (PS) experiments 21 showed that both transmembrane domains (TMDs) of S 68IRS had restricted mobility meaning they were incorporated in the lipid bilayer while the termini regions had higher mobility as they were solvent exposed. However, TMD1 of S2168 was found to be partially externalized from and interacting with surface of the lipid bilayer while TMD2 remained inside of the lipid bilayer. DEER spectroscopy was used for detailed structural studies and direct comparison between the two forms of pinholin S21. These experiments validated and refined the topology and structural models of both pinholin proteins. The effects of residue mutations on the structural topology and conformational changes of pinholin S21 were also probed using CW-EPR PS and DEER spectroscopic techniques. These studies expanded the application of EPR spectroscopic techniques for the study of membrane proteins in general while also providing a deeper understanding of the structural dynamics, conformational changes, and topology of the complex pinholin S21 protein system.

PROBING THE STRUCTURAL DYNAMICS, CONFORMATIONAL CHANGE, AND TOPOLOGY OF PINHOLIN S21, A BACTERIOPHAGE LYTIC PROTEIN, USING ELECTRON PARAMAGNETIC RESONANCE SPECTROSCOPY

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

Tanbir Ahammad

The Graduate School Miami University Oxford, Ohio

2020

Dissertation Director: Dr. Gary A. Lorigan

Tanbir Ahammad

2020

TABLE OF CONTENTS

Chapter 1: Introduction 1 Part A: Membrane Proteins Structural Studies: Significance, Challenges, and Solutions 1 A.1.1 Membrane proteins and their significance…………….……………… 2 A.1.2 Challenges in MP structural characterization………..………………. 5 A.1.2.1 Inherent properties of MPs………………………………………. 5 A.1.2.2 Membrane mimetic systems………………………………………... 5 A.1.2.3 Limitations of biophysical techniques for MP structural characterization……………………………………………………………… 7 A.1.3 Promising solutions to the structural study of MPs …………………. 8 A.1.3.1 EPR Spectroscopy………………………………………………… 9 A.1.3.2 Site Directed Spin Labeling………………………………………. 11 A.1.3.3 Continuous Wave (CW) EPR spectroscopy………………………. 13 A.1.3.4 Pulsed EPR Spectroscopy Techniques…………………………… 16 A.1.3.5 Solid Phase Peptide Synthesis…………………………………….. 17 References…………………………………………………………………… 20 Part B: Pinholin S21, a prototype Holin protein from Bacteriophage Ф21…… 34 B.1.1 Bacteriophages and their significance………………………………… 34 B.1.2 Phage lytic system……………………………………………………… 34 B.1.2.1 Holin-endolysin……………………………………………………… 36 B.1.2.2 Pinholin-SAR endolysin …………………………………………… 39 B.1.3 The Pinholin S21…………………………………………………………. 41 B.1.3.1 Structure and topology of Pinholin S21……………………………… 41 B.1.3.2 Pinhole formation…………………………………………………….. 43 B.1.3.3 Lysis timing………………………………………………………… 44 References…………………………………………………………………… 45

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Chapter 2 CW-EPR Spectroscopy Reveals the Structural Topology and Dynamic 51 Properties of Active Pinholin S2168 in a Lipid Bilayer 2.1 Abstract……………………………………………………………………. 52 2.2 Introduction………………………………………………………………… 53 2.3 Experimental Methods…………………………………………………… 54 2.3.1 Solid Phase Peptide Synthesis………………………………………… 54 2.3.2 Purification and Spin Labeling………………………………………… 55 2.3.3 Peptide Incorporation into Proteoliposomes……………………………. 55 2.3.4 Circular Dichroism Spectroscopy………………………………………. 56 2.3.5 CW-EPR Spectroscopy…………………………………………………. 56 2.3.6 Spin-label Mobility Analysis…………………………………………… 56 2.3.7 CW-EPR Power Saturation Experiments……………………………… 57 2.4 Results……………………………………………………………………… 58 2.4.1 CW-EPR Line Shape Analysis of Pinholin S2168……………………… 60 2.4.2 Structural Topology with Respect to the Lipid Bilayer………………… 65 2.5 Discussion………………………………………………………………… 66 2.6 Conclusion…………………………………………………………………. 70 2.7 Supporting Information…………………………………………………… 71 Acknowledgment……………………………………………………………… 71 References……………………………………………………………………… 72

Chapter 3 Structural Dynamics and Topology of the Inactive Form of S21 Holin in a 78 Lipid Bilayer Using CW-EPR Spectroscopy 3.1 Abstract…………………………………………………………………… 79 3.2 Introduction………………………………………………………………… 80 3.3. Experimental Methods……………………………………………………. 82 3.3.1 Peptide Synthesis and Purification……………………………………… 82 3.3.2 Peptide Incorporation into Proteoliposomes……………………………. 82 3.3.3 Circular Dichroism Spectroscopy………………………………………. 83

3.3.4 CW-EPR Spectroscopy…………………………………………………. 83 3.3.5 Spin-label Mobility Analysis…………………………………………… 84 3.3.6 CW-EPR Power Saturation Experiments……………………………….. 84 3.4 Results……………………………………………………………………… 85 21 3.4.1 Structural dynamics of antipinholin S 68IRS…………………………… 88 21 3.4.2 Structural topology and interaction of S 68IRS with the lipid bilayer… 92 3.5 Discussion …………………………………………………………………. 95 3.6 Conclusion…………………………………………………………………. 97 3.7 Supporting Information…………………………………………………….. 98 Acknowledgment……………………………………………………………… 99 References…………………………………………………………………….. 100

Chapter 4 Conformational Differences are Observed for the Active and Inactive 106 Forms of Pinholin S21 Using DEER Spectroscopy 4.1 Abstract…………………………………………………………………… 107 4.2 Introduction………………………………………………………………. 108 4.3 Experimental Methods…………………………………………………… 110 4.3.1 Peptide Synthesis, Purification, and Spin Labeling…………………… 110 4.3.2 Peptide Incorporation into Proteoliposomes………………………… 111 4.3.3 EPR Spectroscopic Measurements…………………………………… 111 4.3.4 Structure Refinement of the Active and Inactive Conformations of Pinholin S21 using DEER Distance Restraints………………………………… 112 4.4 Results……………………………………………………………………… 113 4.4.1 DEER Distance measurement for the active conformation of pinholin S2168…………………………………………………………………………… 114 4.4.2 DEER Distance measurement for the inactive conformation of antiholin 117 4.4.3 Structure Refinement of the active pin holin and inactive antipinholin from MTSL DEER Distance Restraints ………………………..….………… 119 4.5 Discussion……………………………………………………..………….. 120 4.6 Conclusion………………………………………………………………… 121

4.7 Supporting Information……………………………………………….. 122 Acknowledgment………………………………………………………….. 123 References………………………………………………………………… 124 Chapter 5 Mutations of Pinholin S21 Induce Structural Topology and Conformational Changes are Observed with Electron Paramagnetic 132 Resonance Spectroscopy 5.1 Abstract………………………………………………………………... 133 5.2 Introduction……………………………………………………………. 134 5.3Experimental Methods…………………………………………………. 135 5.3.1 Peptide Synthesis, Spin Labeling, and Purification………………… 135 5.3.2 Peptide Incorporation into Proteoliposomes………………………… 136 5.3.3 CW-EPR Spectroscopy……………………………………………… 137 5.3.4 CW-EPR Power Saturation Experiments…………………………… 137 5.3.5 DEER Spectroscopic Measurements………………………………… 138 5.4 Result and Discussion…………………………………..……………… 138 5.4.1 DEER distance measurements reveal the effects of residue mutation on conformational change in the pinholin system ……………………………. 146 5.6 Conclusion………………………………….…………………………… 150 Acknowledgment………………………………….………………………… 150 References…………………………………………………………………. 151 Chapter 6 Conclusions and Future Directions 156 6.1 Summary of Dissertation Research………………………………………. 157 6.2 Future Directions………………………………………………………….. 160 6.2.1 Oligomerization of Pinholin S21 Studied Using DEER Spectroscopy…. 160 6.2.2 CW-EPR Alignment Techniques to Probe the Relative Orientation of TMD1 and TMD2……………………………………………………...... 161 6.2.3 Application of ESEEM to probe the helical length…………………… 161 6.3 Final Remarks……………………………………………………………. 162 References……………………………………………………………………. 163

LIST OF TABLES

Table 4.1: Summary of the major peak DEER distance for the active pinholin…… 117 Table 4.2: Summary of the major peak DEER distance for the inactive pinholin 21 S 68IRS…………………………………………………………………………… 119 Table 4.3: Minimum energy associated with the refined structures of active pinholin obtained from the simulated annealing molecular dynamics simulation... 122 Table 4: Minimum energy associated with the refined structures of inactive antipinholin obtained from the simulated annealing molecular dynamics simulation………………………………………………………………………… 123

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LIST OF FIGURES

Figure 1.1: Schematic drawing showing the classification of MPs based on membrane interaction…………………………………………….……………………………. 3 Figure 1.2: The relationship between protein structure, dynamics, environment, and function……………………………………………………………...... 4 Figure 1.3: Statistics of MP structure deposition in the RSCB PDB……………… 4 Figure 1.4: Representative images of different membrane mimetics incorporated with putative pinholin S21 protein………………………………….…………………… 6 Figure 1.5: Statistics of total protein structures in RSCB PDB…………………… 8 Figure 1.6: Energy level diagram for an S=1/2 unpaired electron as a function of the magnetic field………………………………..…………………………………….. 10 Figure 1.7: Structures of available nitroxide spin labels used in the SDSL EPR spectroscopy……………………………………………………………...... 12 Figure 1.8: MTSL spin labeling reaction with cysteine residue thiol yielding a nitroxide functionalized side chain (in the box) attached to the protein via a disulfide 12 bond…………………………………………………………………………………. Figure 1.9: Energy level diagram for a nitroxide spin label………………………… 15 Figure 1.10: Schematic presentation of Gram-negative bacterial cell envelope disruption by specific protein systems………………….……………………………………….. 35 Figure 1.11: A schematic representation of the lysis gene cassette for different phages………………………………………………………………………………… 36 Figure 1.12: Topological classification of holins……………..…………...... 37 Figure 1.13: The general model for holin triggering and S hole formation…………. 38 Figure 1.14: Comparison of two pathways for phage lysis in Gram-negative hosts… 40 Figure 1.15: Phage 21 lytic system and pinholin protein……………..……………… 42 Figure 1.16: Structural topology and conformational change of pinholin S21………... 43 Figure 1.17: Schematic presentation of pinhole formation…………..…..…………… 44 Figure 2.1: (A) Primary sequence and topology of S2168………………..…………… 59 Figure 2.2: CD spectrum of pinholin S2168 G40R1 in DMPC proteoliposomes……… 59

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Figure 2.3: CW-EPR spectra of R1 side chains attached at indicated positions of S2168 by replacing the native amino acid with Cys………………………………….. 61 Figure 2.4: EPR mobility analysis of R1 side chain of S2168, calculated from the inverse width of the central resonance line (δ-1)……………….……………………… 62 Figure 2.5: Scaled mobility as a function of residue position for pinholin S2168……………………………………………………………………...... 63 Figure 2.6: Rotational correlation time () of spin-labeled pinholin S2168 in DMPC proteoliposomes as a function of residue position…………………………….. 64 Figure 2.7: Representative CW-EPR power saturation curves of S2168 in DMPC proteoliposomes……………………………………………………………….. 65 Figure 2.8: Membrane depth parameter () as a function of S2168 residue positions in DMPC proteoliposomes………………………………..………………………. 67 Figure 2.9: Proposed topology of active pinholin S2168 after partial externalization of TMD1 from the lipid bilayer…………………………………………………… 69 Figure 2.10: Representative DLS spectrum of pinholin S2168 G48R1 incorporated into DMPC proteoliposomes………………………………………………………….. 71 Figure 3.1: Primary sequence and topology of S21…………………..…………… 86 21 Figure 3.2: DLS spectrum for S 68IRS F49R1 incorporated into the DMPC proteoliposomes…………………………………………………………………… 87 21 Figure 3.3: Circular Dichroism spectra for inactive pinholin S 68IRS with and without spin-label ………………………………………………………...... 88 Figure 3.4: Representative CW-EPR spectra with R1 situated at the indicated positions 21 of S 68IRS ...... 89 Figure 3.5: The relative mobility of R1 (δ-1) as a function of residue positions of the 21 primary sequence of S 68IRS……………………………………………………….. 91 Figure 3.6: The rotational correlational time (τ) as a function of residue positions of the 21 primary sequence of S 68IRS………………………………………………………… 92 21 Figure 3.7: Representative CW-EPR power saturation curves of S 68IRS in DMPC proteoliposomes……………………………………………………………………….. 93 21 Figure 3.8: Calculated depth parameter (Φ) as a function of S 68IRS residue positions in DMPC proteoliposomes……………………………………………………. 94 ix

21 Figure 3.9: The proposed structural topology of inactive pinholin S 68IRS incorporated into a lipid bilayer……………………………………………………………… 97 Figure 3.10: (A) Representative HPLC Chromatogram of the antipinholin peptide. (B) Representative MALDI-TOF mass spectrum of the antipinholin……………… 98 Figure 3.11: Characteristic depth parameter pattern for the transmembrane helix . 99 Figure 4.1: Primary sequence and tentative topology of active pinholin and inactive antipinholin………………………………………………………………………. 114 Figure 4.2: Q-band DEER data of active pinholin S2168 in DMPC Proteoliposomes 116 21 Figure 4.3: Q-band DEER data for inactive pinholin S 68IRS in DMPC proteoliposomes…………………………………………………………...... 118 Figure 4.4: Results of the structural refinement of the active pinholin and inactive antipinholin incorporating MTSL DEER distance-restraint data using an Xplor-NIH simulated annealing molecular dynamics protocol……………………………….. 120 Figure 5.1: Primary sequence and structural topology of active pinholin and inactive antipinholin…………………………………………………………………………. 139 Figure 5.2: Changes in hydrophobicity influenced the externalization of TMD1 of 141 21 S 68IRS……………………………………………………………………………… 21 Figure 5.3: Mutational effect on the externalization of S 68IRS TMD1……………. 143 21 Figure 5.4: Effect of MTSL on the externalization of S 68IRS TMD1……………… 144 21 Figure 5.5: Comparison of the depth parameter for different S 68IRS mutants with respect to the IRS_A20R1………………………………………..…………………… 145 21 Figure 5.5: Comparison of the depth parameter for different S 68IRS mutants with respect to the IRS_A20R1……………………………………………………………. 147 21 Figure 5.7: Spin label induced activation of inactive antipinholin S 68IRS. Pair wise spin label positions and probable distances are shown with arrows for active conformation of S21…………………………………………………………...... 149

DEDICATION

This dissertation is dedicated to my parents, Late grandparents, parents-in-law, and all my family members.

ACKNOWLEDGEMENTS

First, I would like to thank my advisor, Dr. Gary A. Lorigan, for giving the opportunity to work in his research group and for all the help and support he provided me along the way. I have learned so much from him, not only about chemistry and research but also about presentation, scientific communication, writing, hard work, and time management. He is my role model for teaching! I would also like to thank all the members of my committee for their guidance and advice throughout the years: Dr. Carole Dabney-Smith, Dr. Rick Page, Dr. David Tierney, and Dr. Paul Urayama. I would like to express my appreciation for the support and cooperation I received from the current and past members of the Lorigan research group. Dr. Indra D. Sahu for his unconditional help in training, discussion, data analysis, writing, and continuous mental support when my research did not work. Special thanks to Dr. Daniel Drew who started this pinholin project, trained and guided me, tolerated my endless discussion regarding both research and teaching. Rasal H. Khan for continuing this project. Rebecca Stowe, Rehani S. Perera, Dr. Andrew Morris, and Alison Bates for reviewing my papers. Emily Faul and Tianyan Li for being cheerful undergraduate researchers. Thanks to Dr. Lauren Bottorf, Dr. Rongfu Zhang, Dr. Lishan Liu, Andrew Craig, Dr. Gunjan Dixit, F. Dhilhani Mohammad, Rachel Serafin, Brandon Butcher, Benjamin Harding and so many others. I would like to further extend my appreciation to Dr. Robert McCarrick and Dr. Theresa Ramelot for their help with EPR, CD and MS instrumentation. I also want to acknowledge Dr. Paul New for his continuous support, encouragement, and feedback to finish my dissertation. I also want to extend my appreciation to the Department of Chemistry and Biochemistry, all the graduate students who appreciated me, all the TA’s who were excellent colleagues, the department office staff for their smiling faces, and Miami University for sheltering me for the last six years!

I sincerely appreciate the Bangladeshi community in Oxford who helped to make it my home. Finally, I want to thank my father, mother, brothers, and sister as well as rest of my family members and relatives who feel proud of me. Thank you for the unconditional love and support. My sincere gratitude to my wife Dr. Nusrat J. Urmi who is sacrificing her medical career to support me as well as her family members who are supporting both of us. Finally, all my love for my little scientist, Ibrahim Ahammad who pretends to be Dr. Lorigan and motivates me to do protein research.

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Chapter 1: Introduction

Part A: Membrane Proteins Structural Studies: Significance, Challenges, and Solutions

A.1.1 Membrane proteins and their significance Cell membranes are commonly occurring hydrophobic barriers that play a crucial role in living systems by compartmentalizing biological processes via separation of interior and external (sub)cellular environments.1-4 For example, the plasma membrane defines the inner boundary of animal and prokaryotic cells by surrounding the cytoplasm. Inside the cell, different organelles such as the endoplasmic reticulum, chloroplasts, and mitochondria are enclosed by their respective membrane bilayers to ensure the separation of specialized biochemical functions from the other parts of the cell.5 In addition to separation and compartmentalization, these lipid bilayers serve as the environment for several types of membrane proteins (MPs).5 MPs are polypeptide macromolecules that associate with biological membranes to perform several functions such as communication, signal transduction, selective/non-selective transportation of cargo, enzymatic reaction catalysis, and structural support.6-10 Due to their abundance and functional diversity, MPs can be classified in many different ways. One manner of understanding and grouping MPs is through the location of function as their environment is vitally linked to the specific function and stability of the protein.5 Hence, MPs can be separated into nine groups based on the organelle membrane with which they are associated: plasma, nucleus, Golgi apparatus, lysosome, mitochondria, endoplasmic reticulum, peroxisome, chloroplast, or vacuole.5 MPs are classified into six groups based on how they are associated with the membrane bilayer (Figure 1.1).5

Figure 1.1: Schematic drawing showing the classification of MPs based on membrane interaction. (a) type-I, (b) type-II single-pass transmembrane, (c) multi-pass transmembrane, (d) lipid chain- anchored, (e) glycosylphosphatidylinositol (GPI) anchored, and (f) peripheral membrane protein. (a), (b) and (c) are known as integral MPs where (d) and (e) are membrane-anchored protein. Peripheral MPs interact with other MPs but do not directly interact with the membrane bilayer. Figure 1.1 is drawn based on Chou et al. (1999).5

Approximately 20-30% of the total proteome of most cells are MPs and they perform a significant amount of specific and nonspecific cellular and physiological functions.10-15 Given their key roles in several physiological processes, many MPs have been linked to several pathophysiological conditions. Specifically, mutations in the primary sequence and misfolding events of MPs have been implicated as a direct cause or increasing susceptibility to numerous human diseases including cancer, heart disease, obesity, and cystic fibrosis.8 Owing to their functional importance and diversity, approximately 60% of available drug therapies target MPs.8, 16-19 For efficient drug design, it is crucial to know the specific structural details of the MP of interest due to the fact that stability and function are dictated by the MPs structural topology and dynamics as well as the surrounding environment (Figure 1.2).20-24

Figure 1.2: The relationship between protein structure, dynamics, environment, and function.10 As such, enormous efforts have been devoted to solving MP structures to meet the demand for high-resolution maps of potential drug targets. Overall, the progress of protein structure determination has been exponential, but the rate of MP structure determination has lagged behind soluble proteins during the equivalent time period (Figure 1.3).25 Despite their biological abundance, physiological significance, and potential pharmaceutical importance, our collective understanding of MP structures is still lacking.26 Thus, detailed MP structure and dynamics information are necessary to gain a deeper understanding of their function in normal and pathophysiological states.26, 27 Currently, there are only 1040 unique MP structures in the RCSB Protein Data Bank (PDB) which is <0.7% of the 151492 total unique deposited structures.8, 18, 27- 29

Figure 1.3: Statistics of MP structure deposition in the RSCB PDB. (A) The cumulative number of MP structures deposited since 1985. 29 (B) Pie chart representation of the difference between unique membrane protein and soluble protein structures in the RSCB PDB.

A.1.2 Challenges in MP structural characterization Over the last few decades, enormous efforts have been carried out in structural biology to determine MP structures as they are notoriously difficult to study.7, 30-32 The major challenges in the structural study of MPs arise due to their inherent physical and chemical natures, the complexity of the native membrane environment, and the limited applicability of available biochemical and biophysical tools.33 One of the major issues of structural characterization is the difficulty in obtaining sufficiently pure MPs that are still in their native environment. Traditional methods such as E. coli or other bacterial expression systems often only yield small quantities of functional and stable MPs.32, 34 In addition, many eukaryotic MPs require post-translational modification for proper folding and functional stability which are absent in these bacterial expression systems.6, 30, 32 In such cases, insect cell expression systems are required but these methods are more difficult to use.

A.1.2.1 Inherent properties of MPs The complex relationship between MPs and their environment must always be taken into consideration. MPs have highly hydrophobic segments that interact with the hydrophobic acyl chains of lipids in the bilayer resulting in structural rigidity. Additionally, MPs can have amphipathic and/or hydrophilic segments that are exposed in various degrees to aqueous, inter or extracellular environments. In several cases, MPs interact with all three through the hydrophobic acyl chains, the polar lipid head groups, and the aqueous environment.35 These result in very complex and dynamic structures with flexible and rigid segments in the respective polar and nonpolar environments. As a result, MPs are usually highly flexible as well as highly hydrophobic and only stable in particular conditions.36 As such, purification and reconstitution of MPs must be carried out in native or near-native mimetic membrane systems when studying their structure and function.4, 6, 10, 18, 37

A.1.2.2 Membrane mimetic systems There are several approaches available to mimic specific lipid bilayer environments although none of them are truly native.38 Additionally, there are no membrane mimetic systems that are universally compatible with every MP which necessitates costly and time-consuming optimization trials in this process. Currently, there are several commercially or synthetically available mimetic membrane systems including detergent micelles, bicelles, vesicles/proteoliposomes, nanodiscs,

5 and SMALPs (Styrene Maleic Acid Lipid Particles) (Figure 1.4). 39-44 45, 46 Each of these mimetic systems have their own strengths and limitations of use in MP structural studies.10, 47-49

Figure 1.4: Representative images of different membrane mimetics incorporated with putative pinholin S21 protein. (a) micelle, (b) bicelle, (c) liposome, (d) nanodisc, and (e) SMALP.

Detergent micelles are the most commonly used membrane mimetic system as they have excellent MP solubilizing abilities, a relatively small size, compatibility with most lipids, and have been used in several high-resolution structural analyses by common biophysical techniques such as Nuclear Magnetic Resonance (NMR) and Electron Paramagnetic Resonance (EPR) spectroscopy.48-50 The major limitation of micelles is that they are not true lipid bilayers. They poorly mimic native membranes leading to alterations in the structure and conformation of solubilized MPs. 10, 48, 49, 51-53 To avoid inaccurate structural information, the use of other lipid bilayer mimetic systems is highly recommended.54 Bicelles have been used as an alternative lipid bilayer to overcome the limitations of micelles. Bicelles are formed by mixing specific ratios of long-chain and short-chain phospholipids. They are favorable for the study of MP interactions as they allow examination of both the cytoplasmic and extracellular regions of the protein of interest.55-58 However, certain combinations of different

6 phospholipids can limit their applications for some types of MPs since lipid composition has a strong influence on structure and function relationship of membrane proteins.47, 55, 59 Vesicles, nanodiscs, and SMALP nanoparticles more closely resemble the membrane bilayer and each has its own advantages and limitations. Homogenous vesicles are a convenient and widely used mimetic due to their similarity to cellular membrane architecture. However, incorporation of proteins into vesicles can be extremely challenging especially when higher protein:lipid ratios are needed as this often leads to a heterogeneous distribution of vesicle sizes and increased MP aggregation.47, 55, 60, 61 Additionally, it is challenging to study the cytoplasmic domain of membrane proteins in vesicles due to the inaccessibility of the interior cavities of liposomes.55 Nanodiscs consist of lipids surrounded by a membrane scaffold protein (MSP) and have been shown to better retain a native-like membrane environment during MP incorporation.42, 62, 63 Nanodiscs can be formed in several sizes using a variety of lipids and have been reported to provide better stability and homogeneity than other mimetic systems.64-66 However, the formation of nanodiscs requires the use of detergents for protein incorporation leading to the need for a lengthy and extensive detergent removal process.67 Additionally, it is associates with a high cost for membrane scaffold protein (MSP) and the MSP may interfere with the protein of interest.42, 44 To overcome this limitation, a new lipid nanoparticle system was recently introduced called SMALPs. Styrene and maleic acid (SMA) co-polymer is used instead of MSPs to bind the lipid-protein complex. They provide for better incorporation of MPs with minimum interference while also providing accessibility to both sides of the protein which allows for the study of MP oligomerization.47, 48, 60, 67-73 SMALPs can be formed with any kind of lipids and in a detergent free environment to maintain the native lipid composition in solubilized nanoparticles.74-76 However, this new lipid mimetic system needs further characterization in the presence of MPs.43, 68

A.1.2.3 Limitations of biophysical techniques for MP structural characterization Regardless of which mimetic membrane system is used, the lipid environment adds another layer of complexity due to their complex, heterogeneous, and dynamic environments.77 Almost 90% of the solved protein structures in the PDB were determined by X-ray crystallography and 9% of current structures have been solved using NMR spectroscopy (Figure 1.5).78

Figure 1.5: Statistics of total protein structures in RSCB PDB. (A) The cumulative number of total protein structures deposited in PDB. (B) Comparison of the frequency of biophysical techniques used to solve protein structures.78

However, the complexity of the membrane environment and the inherent properties of MPs severely limit the application of popular biophysical techniques such as X-ray crystallography and NMR spectroscopy for use with MPs.7, 8, 30, 79-84 The generation of high resolution structural data using these techniques requires a large amount (mg scale) of properly folded, pure protein.32, 34 X- ray crystallographic studies are limited by the hydrophobicity of MPs that makes it very challenging to produce high-quality protein crystals with the appropriate size and density.18, 27, 85 For solution NMR spectroscopy, significant anisotropy is introduced as the size of the protein as well as the size of the protein-lipid complex increases which causes spectral line broadening and overlapping. This limits the application of NMR spectroscopy for the study of MPs.31, 83, 86 Furthermore, the conformation adopted by a protein in a crystal or detergent-micelle environment may not truly represent the biologically active structure or will lack the structural, conformation and dynamics of the native system.20, 87-90 So, this requires the use of biophysical techniques that offer structural dynamics and conformational related data to properly characterize functionally active MPs in their native or near-native mimetic environments.

1.3 Promising solutions to the structural study of MPs

There are several techniques commonly used to study the structural dynamics of MPs in native and mimetic environments including various spectroscopic techniques (EPR, solid-state NMR, fluorescence, and Raman Spectroscopy), mass spectrometry, and computational modeling 8 methods.2, 20, 87, 91-108. In addition, cryogenic electron microscopy (Cryo-EM) has emerged as a rapidly growing powerful structural biology tool for determining the three-dimensional structures of biomolecules at near-atomic resolutions.109, 110 Cryo-EM is applicable to a variety of sample types with a wide range of molecular weights and overcomes the major limitations of X-ray crystallography and NMR spectroscopy as it requires much smaller amounts of sample.109 Cryo- EM experiments can also be done in the presence of a lipid environment without protein crystallization.109 However, this method has a size limitation below the average molecular weight (MW) of cellular proteins (<50 kDa) as the resolution is significantly reduced below this limit.109, 111 Fortunately, EPR spectroscopy can be used to study the structural dynamics of MPs in their native environment and overcomes the limitations of X-ray crystallography, Cryo-EM, and NMR spectroscopy.

1.3.1 EPR Spectroscopy EPR spectroscopy is a magnetic resonance spectroscopic technique carried out with unpaired electrons. It is a powerful biophysical technique for studying the structure, dynamics, and electronic properties of materials with unpaired electrons.87, 91 EPR spectroscopy has been particularly useful for characterizing metal complexes, organic radicals, and materials with paramagnetic centers since it was first introduced by Zavoisky in 1944.112 Although the basic concepts of EPR are analogous to NMR, EPR spectroscopy involves electron spin transitions instead of nuclear spin transitions.33 Electrons have a relatively higher magnetic moment than nuclei making this technique more sensitive than NMR. The higher sensitivity produces spectra with a larger signal-to-noise ratio from samples with a low concentration of proteins (µmol quantities) as compared to NMR. This makes EPR spectroscopy a powerful biophysical technique to study more challenging biological samples like MPs with inherently low protein concentrations.87, 91, 113

The simplest EPR system is a single unpaired electron spin in a molecular orbital. An electron has a spin quantum number of S=1/2, so it is in one of the two possible magnetic quantum states, Ms= +1/2 or -1/2. These two states are degenerate in the absence of a static magnetic field. However, in the presence of an applied magnetic field, these two states become separated as a function of the magnetic field strength (Figure 1.6). The energy gap between these two states, E, can be quantified using the equation E = gβeB0 in which g is the electronic g-factor, βe is the Bohr 9

-1 Magnetron (joules*Gauss ), and B0 is the strength of the static applied magnetic field (Gauss). Resonance occurs when the energy gap (E) matches the microwave radiation energy (E). At resonance, electrons in the external magnetic field can transition between the ground state and the excited state by absorbing or emitting photons with energy, (E = hν where h is Planck’s constant and  is the frequency of the photon. Thus, the energy gap at the resonance condition can be calculated by the equation ∆E=hν=gβeB0.

By applying the EPR transition rule (Ms= ±1), only one spin transition will occur for a free unpaired electron. If there is no other interaction between the free electron with nearby nuclei, the EPR spectrum will be observed as a single peak (Figure 1.6).87, 114 In a typical EPR spectrum, the first derivative of the absorption spectra is recorded instead of the direct absorption.

Figure 1.6: Energy level diagram for an S=1/2 unpaired electron as a function of the magnetic field. EPR transitions due to Zeeman interactions are shown with the absorption peak at the resonance condition which can be expressed as the corresponding EPR signal spectrum.91, 114

EPR spectroscopy offers high sensitivity without macromolecular size limitations and is independent of the optical properties of the sample. EPR measurements can be made with complex mixtures including different membrane mimetic systems, suspensions, tissues, or frozen samples 10 which makes it a versatile technique to study both soluble and membrane proteins.91 Although EPR spectroscopy alone is unable to be used to solve the structure of a protein like X-ray crystallography or Cryo-EM, it is an excellent complementary technique to study the structural dynamics and conformations of proteins that are not readily accessible with other biophysical techniques.

1.3.2 Site Directed Spin Labeling Most native proteins are EPR silent as they lack inherent paramagnetic properties except for metalloproteins that contain an EPR active metal. This has restricted the application of EPR spectroscopy to metalloproteins with paramagnetic centers or enzymes with radical cofactors.91, 114 Fortunately, site-directed spin labeling (SDSL) allows the incorporation of a reporter group such as a radical spin probe at specific sites of any biological system.87, 114 Over the past few decades, the development and improvement of SDSL boost the application of EPR spectroscopic techniques to study the structure and dynamics of previously EPR silent proteins and nucleic acids.96 The combination of EPR spectroscopy with SDSL has emerged as a powerful biophysical technique for investigating the structure and dynamics of soluble and membrane proteins.20, 87, 91- 100 The most frequently used spin labels introduce a stable nitroxide free radical (known as nitroxide spin label) at specific positions in the biomolecule of interest (Figure 1.7).114

Figure 1.7: Structures of available nitroxide spin labels used in the SDSL EPR spectroscopy. (a) methanethiosulfonate spin label (MTSL), (b) maleimide spin label (MSL) N-(1-oxyl-2,2,6,6- tetramethyl-4-piperidinyl) maleimide, (c) iodoacetamide spin label (ISL), (d) bis(1-oxyl-2,2,5,5- tetramethyl-3-imidazolin-4-yl) disulfide (IDSL), (e) bifunctional spin label (BSL), (f) 2,2,6,6- tetramethyl-N-oxyl-4-amino-4-carboxylic acid (TOAC), and (g) 4-(3,3,5,5-tetramethyl-2,6-dioxo- 4-oxylpiperazin-1-yl)-l-phenylglycine (TOPP) (Figure adapted with permission).114

The most common SDSL methods rely on covalent modification of cysteine residues with thiol-reactive spin labels yielding a nitroxide functionalized side chain that acts as a reporter group in EPR (Figure 1.8).87, 114

Figure 1.8: MTSL spin labeling reaction with cysteine residue thiol yielding a nitroxide functionalized side chain (in the box) attached to the protein via a disulfide bond. 12

To achieve site-specific spin labeling, all native non-disulfide bonded cysteines in a protein of interest must be replaced with another amino acid while minimizing structural perturbations. Cysteine residues are then introduced into the primary sequence followed by reaction with a sulfhydryl-specific nitroxide reagents to generate stable spin labeled side chains (Figure 1.8).114, 115 Several studies have robustly demonstrated that many types of proteins are highly resilient to SDSL methods and retain their biological function after spin labeling.87

The most used nitroxide spin label is MTSL due to its convenience and high labeling efficiency with minimum structural perturbation. However, MTSL has a large degree of conformational flexibility due to five flexible bonds making it less reliable for distance measurements and protein backbone motion studies.116 The bifunctional spin label (BSL) which covalently binds with two adjacent Cys residues is able to overcome this drawback to some extent by forming a more rigid reporter group structure.116 However, its application is limited due to its high cost and low labeling efficiency. Other spin labels such as TOAC and TOPP (Figure 1.7F- G) are unnatural amino acid analogs that can be directly incorporated into the peptide backbone using solid-phase peptide synthesis (SPPS).114 These two spin labels have a rigid structure that is free from fluctuations in dynamics and are ideal probes to study the backbone dynamics and alignment of MPs with respect to the lipid bilayer.117, 118 However, their use has been limited to only synthetic peptides due to the difficulty of incorporating these spin labels into recombinant proteins with conventional protein expression techniques.

Another technique that is used to prepare proteins for EPR spectroscopic studies is the introduction of EPR active transition metal (e.g. Co2+, Cu2+, Gd3+) to create a paramagnetic metal center in the protein of interest.119-121 These paramagnetic metal probes can be introduced into proteins at native metal-binding sites or engineered chelating sites.119 Recently published work has shown the improvement of double Histidine-based Cu2+ labeling methods for EPR based distance measurements in solution proteins.119 However, the application of this technique has yet to be used to study MPs.

1.3.3 Continuous Wave (CW) EPR spectroscopy CW-EPR spectroscopy is the most frequently used EPR technique due to its relatively simple instrumentation requirements, shorter data collection timeframes, and the ease of data analysis as compared to pulsed EPR spectroscopic techniques. In CW-EPR, constant microwave 13 radiation is applied to the sample while the external magnetic field (B0) is swept with predefined magnetic field modulations. When the resonance conditions are met, spin transitions occur according to EPR transition rules. The collected EPR spectrum is recorded as the first derivative of the absorption spectrum against the field strength (B0). For SDSL based CW-EPR studies of peptides/proteins, the reported EPR spectrum is that of the residue bound nitroxide spin label. The nitroxide spin label has an unpaired electron which is divided into two energy states (Ms= ± ½) in the presence of an external magnetic field due to the electron Zeeman effect. The energy is further split into multiple energy states due to the hyperfine interactions with neighboring 14N nuclei following the multiplicity rule of 2nI+1 where ‘n’ is the number of magnetically equivalent nuclei and ‘I’ represent the nuclear spin. For MTSL, n=1 and I=1 (14N) which gives six energy states in the presence of Zeeman and hyperfine interactions which allows three possible transitions following the EPR transition rules (ΔMs = ±1 and ΔMI = 0) (Figure 1.9). This results in the characteristic three-line EPR spectra for a nitroxide spin label that is centered at the resonance position and separated by the hyperfine coupling as shown in Figure 1.9. These three peaks are called the low, central, and high field peaks and are denoted by MI= +1, MI= 0, and MI= -1, respectively.87, 91, 114

Figure 1.9: Energy level diagram for a nitroxide spin label. The Zeeman interactions of an unpaired electron and its hyperfine interaction with a 14N nucleus results in three-line EPR spectrum at the appropriate resonance conditions.

The collected spectrum is highly sensitive to the spin labels orientation, dynamics, and interactions which reports valuable structure, topology, and dynamics information of the protein of interest. As such, a variety of CW-EPR techniques have been developed to specifically study MP topology and dynamics including CW-EPR line shape analysis, dipolar broadening, power saturation, and alignment techniques.87, 91 CW-EPR spectral line shape analysis is a powerful biophysical technique that is used to decipher structural dynamic properties of a protein at a residue-specific level with spatial resolution.89, 112, 122-125 Central line broadening (δ) and rotational correlation time (τ) could be a

15 very useful parameter to calculate the mobility of protein residues or segments and will be explained in more detail in Chapters 2 and 3.126 The power saturation experiment is another important and convenient CW-EPR technique that is used to study the structural topology of proteins with respect to the lipid bilayer. 100 In this technique, CW-EPR spectra are collected with varying microwave power in the presence and absence of a lipid-soluble paramagnetic relaxation agent (e.g. O2 gas) and a water-soluble paramagnetic relaxation agent (e.g. NiEDDA). The relative intensity and patterns of power saturation data are then used to determine if the labeled residue is likely inside or outside of the lipid bilayer. More detail will be explained in Chapters 2, 3 and 4.126 CW-EPR alignment techniques have been proven to be a useful tool in predicting the relative orientation and calculation of helical tilt in the MP with respect to membrane normal. The hyperfine tensoral components of a rigid spin label (e.g. TOAC) depend on the relative orientation with the magnetic field and significantly change in the hyperfine splitting values at various orientations. Using this parameter, relative orientation and helical tilt angle of transmembrane peptides can be calculated for magnetically or mechanically aligned samples.117, 118 CW-EPR dipolar broadening analysis is used to quantitate short distances (8 Å - 25 Å) between two spin labels through the measurement of short-range magnetic dipole interactions based on the broadening effect.127 This method is useful to predict MP tertiary structures and oligomeric interactions.

1.3.4 Pulsed EPR Spectroscopy Techniques

Distance measurements between different segments of an individual protein or different units of a protein complex can provide valuable information about the proteins structural dynamics, conformational changes, topology, and mechanism of function.128-130 Pulsed EPR techniques can provide higher resolution distance information when compared to CW-EPR experiments.131 In pulsed EPR spectroscopy, a series of short, intense microwave pulses are applied to perturb a spin system.91, 131 Advancement in pulsed EPR spectroscopy has provided new avenues to explore structural properties of extremely difficult biological systems such as MPs. One of the pulsed EPR methods is electron spin echo envelope modulation (ESEEM) spectroscopy. This pulsed method is used to gain information about nuclei near an unpaired electron spin system and is also known as EPR-detected NMR spectroscopy.132, 133 These experiments can be carried out using different pulse sequences with two-pulse or three-pulse

16 sequences being the most utilized.133, 134 ESEEM experiments can detect the close-range interaction (8 Å) between an electron spin coupled with a nearby NMR active nucleus.91, 134 This method can be used to explore the solvent accessibility of the MP as well as its localization within the membrane.135 This technique has been heavily utilized in studying paramagnetic metal centers in metalloproteins, metal binding, and metal coordination.136, 137 Recently, a novel application of the ESEEM technique was developed by the Lorigan lab which successfully identified the local secondary structure of an MP as well as being able to differentiate between beta-sheet, alpha-helix, 132, 133, 138 and 310-helix motifs. Another pulsed EPR technique is double electron-electron resonance (DEER) spectroscopy which measures longer distance interactions between two spin labels (20 Å-80 Å).139-141 In optimized experimental conditions, it can even measure up to 160 Å.129 DEER has several advantages over other molecular distance measurement techniques including FRET as this technique uses a smaller molecular probe. DEER experiments provide average distances as well as distance distributions that are used to predict protein structural parameters and different conformations. The experimentally determined DEER distance restraints can be combined with molecular dynamics simulations to further model protein structures, conformational changes during functional activities, oligomerization, and complex formation between biomolecules. 43, 91, 130, 141-143 In this dissertation, Chapters 4, 5, and 6 will discuss the application of DEER to explore the protein structure, conformational changes induced by residue mutation, and oligomerization of the pinholin protein.

1.3.5 Solid Phase Peptide Synthesis: Many MPs have poor expression and suffer from low yield which is especially common for cytotoxic proteins or those that cause host cell lysis (e.g. holins). Peptide synthesis is a good alternative to circumvent this challenge by providing flexibility for spin labeling and ease of purification methods without any additional tag (e.g. His tag). Solid-phase peptide synthesis (SPPS) is a novel approach to peptide synthesis first introduced by Robert Bruce Merrifield in 1963 that used polymer-based insoluble porous supports to synthesize a tetrapeptide.144 Prior to Merrifield’s solid support technique, peptide synthesis required rigorous isolation, purification, and characterization following each subsequent synthesis step. SPPS circumvents these time- consuming processes by allowing simple wash steps to remove reaction waste products after each synthesis step by anchoring the growing peptide to a solid support. Significant improvements to 17

SPPS have since been reported in the chemical and mechanical aspects of the process leading to its automation and adaption for use at industrial and research scales. In SPPS, the peptide is generated from N-terminus to the C-terminus of the peptide sequence which is opposite to natural protein synthesis where the protein grows from the N- to the C-terminus. The N-terminus and reactive side chains of the free amino acid to be added to the peptide must be protected using an appropriate protecting group such as acid-labile tert-butoxycarbonyl (Boc) or base-labile fluorenylmethoxycarbonyl (Fmoc).145 There are two basic strategies for SPPS depending on the amide protecting group; Boc-protected, and Fmoc-protected SPPS. Each of these methods has its own benefits and limitations. Fmoc-protected SPPS is the most common approach with more robust, efficient, and versatile commercially available reagents and synthesizers that allow single residue or fragment assembly that ultimately yields high-quality peptides at a lower cost and in less time.146 The continuous improvement in methodology, reagents, and instrumentation has significantly improved the quality of and speed of synthesis for difficult peptides.147 In the sequential coupling of amino acids, the yield and purity of the synthetic peptide is dependent on its length with a generally accepted limit of ~70 amino acids.147 However, Drew et al. reported the successful synthesis of a 73 amino acid pinholin peptide using Fmoc-SPPS and a CEM Liberty Blue Synthesizer.148 All of the peptides used in this study were synthesized by Fmoc-SPPS techniques using a CEM Liberty Blue Synthesizer coupled with a Discovery-Bio microwave unit following the previously optimized methodology.126, 148

The SPPS technique starts with a porous solid bead resin consisting of a polyethylene glycol (PEG) or polystyrene (PS) polymer at a specific particle size and contains a specific linker attached to the C-terminus of the first amino acid of the peptide sequence from the N-terminus.149 For resin preloaded with the first amino acid, the N-terminus must be protected by Fmoc that is removed before the addition of the next amino acid.150 Each amino acid is coupled with the previous amino acid of the growing peptide following a series of repeated cycles starting with the deprotection of the N-terminus, washing away of the protecting group, coupling of the new amino acid in the presence of activator and activator base, and finally washing away of any uncoupled amino acid. Piperidine, N, N′-diisopropylcarbodiimide (DIC) and oxyma are commonly used deprotection, activator, and activator base reagents, respectively. dimethylformamide (DMF) is the main solvent used to dissolve or dilute any reagents and perform the wash in every step. After a successful synthesis, the desired peptide will remain covalently bound to the resin through the 18

C-terminus and the N-terminus will be free from Fmoc by final deprotection. An optimized cleavage procedure should be followed to cleave the peptide from the resin as well as remove any sidechain protecting groups.151-153 The cleavage cocktail should contain strong volatile acid, like trifluoroacetic acid (TFA) with one or more scavengers (e.g. anisole, water, 1,2-ethanedithiol (EDT), triisopropylsilane (TIPS)) depending on the optimized condition.126, 132, 148, 154, 155 Scavengers irreversibly bind to the removed protecting groups to prevent rebinding of those groups with the peptide. Free resin is then separated by gravity filtration and the crude peptide is harvested from the filtrate by ether precipitation and centrifugation. The crude peptide is usually purified further with reverse phase high pressure liquid chromatography (RP-HPLC).

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Part B: Pinholin S21, a prototype Holin protein from Bacteriophage Ф21

B.1.1 Bacteriophages and their significance Bacteriophages (BPs) are viruses that infect and kill bacteria without a significant negative effect on human or animal cells.1 These are the most abundant organisms in the biosphere and they recycle much of the world’s biomass through ~1028 infection cycles per day.2, 3 Phages are important scientific tools used to understand fundamental molecular biology, horizontal gene transfer, and bacterial evolution.3 In 1915 William Twort discovered bacteriophages and shortly after in 1917 Felix d’Herelle realized the potential use of BPs to kill bacteria.3 During the pre- antibiotic era, BPs were used to treat a wide range of infectious diseases.1 BPs alone or in combination with antibiotics could be used to treat bacterial infections.1 However, the advent of modern antibiotics and increase in the availability of safe and effective antimicrobial drugs have contributed to the dwindling use of phage therapy.1 Phage therapy has been revived in the current era of antimicrobial resistance to combat against multidrug-resistant (MDR) bacteria.4-9 Phages and their lytic products have been suggested to be potential agents for combating MDR.7 Phages or their bacteriolytic gene cassettes can serve as effective biocontrol agents for food safety and preservation.10-12 Thus, for the judicious use of bacteriophages, understanding the bacteriophage life cycle, and its lytic system is crucial.

B.1.2 Phage lytic system The final step of bacteriophage infection is the lysis of host cell to release the viral progeny.13, 14 Phages have evolved a robust lytic system to control the length of the infection cycle for adaptation to changing environments and hosts.15 The phage lytic system evolved through immense evolutionary pressure to achieve precise timing and effectiveness.15, 16 Apparently, all phages with double-stranded nucleic acid (dsNA) genomes have at least two encoded proteins that are essential for the host cell lysis, an endolysin and a holin,.16-20 For the lysis of double membraned Gram-negative bacteria, a third lytic protein called spanin is needed to lyse the outer membrane.21- 24 The mechanism of host cell lysis is a highly regulated, three step process in which each step is regulated by at least one specific protein (Figure 1.10).15, 21, 25 17

Figure 1.10: Schematic presentation of Gram-negative bacterial cell envelope disruption by specific protein systems. Canonical holin (cyan) form larger holes to release endolysins (purple) which degrade peptidoglycan. Pinholins (orange) form small holes not large enough for folded proteins, which only disrupt the proton motive force of the inner membrane and accelerate the release and activation of SAR-endolysin. One- or two-component spanins disrupt the outer membrane, possibly involving the fusion of the outer membrane with the inner membrane. (Figure is reproduced with permission).17 In most phages, three components of the lytic system are encoded together in the lytic gene cassette. 16, 22 In lytic gene cassette, holin is followed by endolysin and spanin (Figure 1.11).16 Among these lytic genes, holin is the first gene product that controls and maintains the timing of the first and rate-limiting step of the lysis process. Endolysin is the catalytic enzyme responsible for the degradation of the peptidoglycan layer while spanin destroys the outer layer.

Figure 1.11: A schematic representation of the lysis gene cassette for different phages. The lysis genes of the coliphages λ, the Salmonella phages P22 and PS3 are shown. The lysis genes are proximal to the late promoter, PR’ (indicated by arrow). The size of each reading frame in codons is shown above each ORF and unambiguously orthologous relationships are indicated by identical 16 colors. (PR’ of PS3 has not been mapped.)

B.1.2.1 Holin-endolysin Holin is a group of proteins that form holes (pores). This term was initially coined to refer only the bacteriophage proteins which permeabilized the cytoplasmic membranes of bacteria to initiate the host cell lysis and control the phage infection cycle.26 However, subsequent studies have indicated that holins or holin-like proteins are ubiquitous and present across all three domains of life including bacteria, eukaryotes, and archaea and serve a variety of functions.4 Within bacterial genomes, holins serve diverse functions including gene transfer, biofilm formation, promotion of responses to stress conditions, differentiation, and the release of toxins and other proteins.4 In animals, programmed cell death involves holin-like proteins such as Bax and Bak that may have evolved from bacterial holins.4, 27 Holin homologs have also been identified in archaea.4 Holins are diverse group of proteins that consist of ~900 proteins.28 Currently, there are seven super-families with 58 recognized families of holins containing 1-4 transmembrane domain(s) (TMD), but many more families have yet to be discovered.4 Based on structural topology, holins are classified in three classes: Class I, II, and III.29 Class I holins have three TMDs with over 95 residues where the N-terminus is in the periplasm and the C-terminus is in the

36 cytoplasm. Well-characterized Class I holins are Sλ holin of phage λ and holin15 of phage P68.30 Class II holins have two TMDs, with 65 to 95 residues where both the N- and the C-terminus are located in the cytoplasm e.g. S21 holin from phage Ф21, Hol3626 from phage Ф3626.30, 31 Class III holins only have a single, highly hydrophilic TMD with the N-terminus in the cytoplasm and the C-terminus in the periplasm and a well-characterized example is T4 holin from phage T4.29, 32

Figure 1.12: Topological classification of holins.29

Bacteriophage holins play two crucial roles in the phage infection cycle. Their primary function is membrane permeation to release endolysin and their secondary function is determination of the end of the infection cycle timing. Due to their precise timing capabilities, they are known as a molecular clock.4, 16 The holin proteins accumulate benignly in the inner cytoplasmic membrane during the vegetative cycle until the protein “triggers” at an allele-specific time.19 Triggering is the term used to denote when the holin reaches a critical concentration in the membrane and attains the functionality to permeabilize the membrane. Figure 1.13 shows the general mechanism of hole formation by canonical holins.33 Holins start oligomerization and raft nucleation at a critical concentration which is further accelerated by the reduced proton motive force (pmf) ultimately forming large microscale holes.33, 34 The hole formation is inherently saltatory, where the formation of one hole accelerates the process and leads other rafts to be converted to holes27,30.16, 35 These holes are large enough for fully folded functional endolysin to be released to hydrolyze peptidoglycan within seconds.20, 36, 37 Lesions formed by the holin are 37 non-specific and have been termed “holes” to show distinction from channels and other such membrane permeabilization pathways.33

Figure 1.13: The general model for holin triggering and S hole formation.33, 34 Holin accumulate as dimer and proton motive force (pmf) remains -180 mV. At critical concentration holin start to form raft which reduce the pmf. This further accelerate the hole formation and form large hole by conformational change.33

The actual mechanism of holin triggering time is not well understood, although it is believed that this time is governed by the primary sequence of holins.31 Many holins have counter proteins known as antiholins which act antagonistically to prevent or delay hole formation by holin. In many holin systems, holin and antiholin are encoded by the same gene by the virtue of dual start motif gene.17, 31, 35, 38-42 For example, canonical holin Sλ, which is one of the well-studied holin systems has two holin proteins; Sλ105 holin and Sλ107 antiholin.17, 35, 39, 40, 43-49 At normal conditions, the holin:antiholin ratio is 2:1.16 Antiholin Sλ107 prevents Sλ105, from hole formation until the holin critical concentration is reached such that it plays a vital role to achieve precise timing.45 Holin and antiholin have similar primary sequences except antiholin has an extra positive charge in the N-terminus which favors a different structural topology and conformation than that of holin and prevent raft formation. When membrane depolarization occurs due to initial membrane permeabilization, antiholin losses its ability to retain the opposing conformation and both holin and antiholin adopt similar conformations such that all the dimers are converted to active forms, thus active holin dimers will be tripled soon after triggering and large holes form instantaneously.21, 22

B.1.2.2 Pinholin-SAR endolysin Until recently, it was believed that all holins form non-selective, micron-scale holes in the inner cytoplasmic membrane and endolysin passes through these holes to reach the peptidoglycan layer.36 However, more recently, a prototype lytic system was uncovered in some phages which have a significantly different lytic mechanism than the canonical holin-endolysin system.2, 31, 50 The holin in the prototype system is not able to form microscale holes to allow pre-folded endolysin to pass through the cytoplasmic membrane to reach the peptidoglycan layer. Hence, it is not able to induce cell lysis when it is co-expressed with canonical endolysin.31 Instead of making microscale holes, it makes large numbers of nanoscale holes which are large enough only for dissipation of the proton motive force.31, 41 This class of holin is called ‘pinholin” to indicate the smaller holes formed by it. In this class of lytic system, endolysin must be in the periplasm and must remain inactive to prevent immature host cell lysis until the proper assembly and optimum numbers of progeny viruses have been attained. This type of endolysin is called signal-anchor release (SAR) endolysin and has novel N-terminal secretory signals. SAR sequences initially act as signal anchor domains and promote the integration of proteins into the cytoplasmic membrane with type II, N-in, C-out topology.41, 50, 51 SAR endolysins are exported by the host Secretory system and accumulate in the periplasm.51 Although the catalytic domain of the SAR endolysin is secreted to the periplasm, these enzymes are catalytically inactive to prevent premature lysis in their membrane-tethered state until the pinholin trigger event. Pinholin triggering dissipates the pmf which accelerates the release of SAR endolysin. Once untethered from the membrane, the periplasmic catalytic domain of SAR endolysin refolds to the active form and begins to hydrolyze the muralytic layer within seconds.31, 51-53 Thus, for phages encoding SAR endolysins, holins do not have to make large holes as only small holes are needed to depolarize the membrane in order to fulfill their role in controlling the timing of lysis.31

Although the pinholin-SAR endolysin system was discovered more recently, it is the evolutionary ancestor of the holin-endolysin system. Approximately 25% of phages have this prototype lytic system.31 Pinholin-SAR endolysin systems are an inferior lytic system than canonical holing-endolysin system since SAR-endolysin has a 1000 fold less lytic capability than canonical endolysin and can’t be overexpressed as endolysin to prevent premature host cell lysis.31 However, SAR-endolysin coupled with pinholin is able to achieve the intended lytic function at a predefined time with a different mechanism.31, 41 Pinholins are always associated with SAR- 39 endolysin where canonical holins can be associated with both endolysin or SAR-endolysin.31 49, 50 Figure 1.14 depicts the comparison of holin-endolysin vs pinholin-SAR endolysin systems.22

Figure 1.14: Comparison of two pathways for phage lysis in Gram-negative hosts. Schematic views of the holin–endolysin (left) and pinholin–SAR endolysin (right) pathways to lysis. The inner (IM) and outer membranes (OM) of cells are shown with holin or pinholin (blue ovals) accumulating in the IM. During phage morphogenesis, holin or pinholin accumulate harmlessly. In holin-endolysin, endolysin accumulate in the cytoplasm (The orange symbols represent the enzymatically active canonical endolysin). At holin triggering, holin makes micron-scale hole(s) to release active endolysin (In the bottom, left). In contrast, SAR endolysin accumulate as inactive membrane-tethered until pinholin triggering. The pinholin triggers to from many nanoscale pinholes (represented by double ovals with a channel). The red symbols with the closed and open ‘active sites’ represent the inactive and active SAR endolysin, respectively. (Figure are reproduced with permission).22

B.1.3 The Pinholin S21 Holin of lambdoid phage 21 forms small membrane holes and is known as pinholin S21.31, 41 Pinholins represent a large group of holins which are about one-fourth of the overall holin systems.31 Pinholin is the evolutionary ancestor of holin. However, this group of holins was discovered more recently and is underrepresented in the literature. Among the pinholin proteins, pinholin S21, encoded by the S21 lysis gene is one of the most well-studied pinholin systems.41, 42, 54-58 However, the structural details are not well understood. The work in this dissertation will focus on the structural characterization of this under characterized pinholin protein using pinholin S21 as a model pinholin.

B.1.3.1 Structure and topology of Pinholin S21 Like many other phage lysis cassettes, the phage 21 lysis cassette has three components. 21 21 21 21 The S pinholin gene is followed by the R endolysin and spanin (Rz and Rz1 ) as shown in Figure 1.15 (A).41, 42 The S21 is a dual-start motif gene which encodes both pinholin and antiholin from the same reading frame. S2168 is the 68 amino acid long pinholin starting from Met4 and S2171 is the 71 amino acid long antipinholin starting from Met1.

Figure 1.15: Phage 21 lytic system and pinholin protein. (A) Lytic gene cassette. Lysis genes S21, 21 21 21 R , Rz and Rz1 are pinholin, SAR endolysin, outer and inner spanin, respectively, transcribed 21 from late promoter PR’. The length of gene products is given on the top of the gene. S can be transcribed as 71 amino acid long antiholin (S2171) or 68 amino acid active pinholin (S2168). The primary sequence of natural antipinholin (S2171) (B), active pinholin (S2168) (C), Modified 21 41, 42 dominant antipinholin (S 68IRS) (D).

Both pinholin proteins have two transmembrane domains (TMDs) with a short N-terminus and a long positively charged C-terminus.38 For pinhole formation, TMD1 must be externalized from the lipid bilayer, a spontaneous process that quickly occurs for S2168. TMD1 of S2171 externalizes slowly due to the presence of an additional Lys positive charge in the N-terminus which delays pinhole formation. S2171 is a weak antipinholin which can only delay but can’t completely block the externalization of TMD1.42 Ry Young and his lab have reported a dominant antipinholin by inserting five extra amino acids (RYIRS) in the N-terminus of S2168 which completely blocks the externalization of TMD1 due to the presence of two extra positive charges 21 42 21 and bulky side chains, and denoted by S 68IRS (Figure 1.15D). S 68IRS is a convenient structural analog of S2171 and was used to study the structural topology of antipinholin. Figure 1.16 shows the structural topology and conformational changes for all three S21 pinholin proteins.

Figure 1.16: Structural topology and conformational change of pinholin S21. (A) Active pinholin S2168. TMD1 externalizes quickly and adopts one of two possible orientations. (B) Natural antipinholin S2171. Due to the extra positive charge in the N-terminus, TMD1 externalizes slowly. 21 21 After externalization, it adopts a similar orientation to S 68. (C) Dominant antiholin S 68IRS in which TMD1 cannot externalize due to two extra positive charges.

B.1.3.2 Pinhole formation During the vegetative cycle, pinholin is incorporated in the cytoplasmic membrane as inactive monomers that accumulate benignly as an inactive dimer with both TMDs in the lipid bilayer. Since the S2168:S2171 production ratio is ~2:1, the dominant dimer formations are that of the homodimers S2168:S2168 and heterodimers S2168:S2171.54 TMD1 of pinholin S21 is the inhibitory domain and TMD2 is the functional domain which makes the pinhole by oligomerization. TMD1 inhibits TMD2 both in cis and in trans if it is inside of the lipid bilayer. It keeps the dimer in one of two possible inactive forms and prevents the subsequent oligomerization and pinhole formation. Once holin triggers, it accelerates the externalization of TMD1 and converts all inactive dimer to active dimer.42, 54 When a critical concentration of active dimer is reached, it starts to form 2D aggregates which in turn form heptameric pinholes with an average size of 13-16 Å.41 43

Figure 1.17: Schematic presentation of pinhole formation.41, 42

B.1.3.3 Lysis timing Every holin triggers at a specific time that is programmed into the structure of the holin by natural selection.16, 59 Phages use the combination of pinholin and antipinholin to adjust this lysis timing.16, 31 In addition, phages tune their lytic function by the mutation of endolysin and/or holin to adjust with the changing environments.16, 31 However, the mutation of SAR-endolysin alone is not a good alternative way to advance or slow lysis timing because there are very few mutational 31 options in the membrane-tethered N-terminus without changing the effective Kcat. Hence, mutations of pinholin could be a more viable alternative to adjust lysis timing with changing environments. Pang et al. reported an extensive mutational study of S21 and showed a wide range of phenotypes including absolute lysis defectives, delayed, or accelerated lysis triggering by pinholin S21.42 To evaluate the mutational effect on pinholin function, they monitored the cell lysis timing which itself is not enough to study the pinholin activity due to the fact the SAR endolysin itself can cause lysis at some point after induction, irrespective of the pinholin allele.31, 42

In this study, EPR spectroscopy was used to probe mutational effects on the structural topology of pinholin S21. As pinholin S21 is a membrane protein (MP) and its functions are strictly associated with its membrane association and topological change, the structural characterization must be done in the presence of lipid bilayer to maintain its structural-function integrity. In this project, EPR spectroscopy coupled with site-directed spin-labeling (SDSL) was used to study the structural topology and dynamics of pinholin S21, which is a suitable biophysical technique to study the MPs in the presence of a lipid bilayer.60-67 This protein is associated with bacterial cell lysis which makes it very challenging to get enough pure protein using bacterial expression techniques. Hence, solid-phase peptide synthesis (SPPS) technique was used to generate the pinholin S21 protein for the subsequent studies. 44

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38. Barenboim, M.; Chang, C. Y.; Hajj, F. D.; Young, R., Characterization of the dual start motif of a class II holin gene. Molecular Microbiology 1999, 32 (4), 715-727. 39. Chang, C. Y.; Nam, K.; Young, R. Y., S-GENE EXPRESSION AND THE TIMING OF LYSIS BY BACTERIOPHAGE-LAMBDA. Journal of Bacteriology 1995, 177 (11), 3283-3294. 40. Graschopf, A.; Blasi, U., Functional assembly of the lambda S holin requires periplasmic localization of its N-terminus. Archives of Microbiology 1999, 172 (1), 31-39. 41. Pang, T.; Savva, C. G.; Fleming, K. G.; Struck, D. K.; Young, R., Structure of the lethal phage pinhole. Proceedings of the National Academy of Sciences of the United States of America 2009, 106 (45), 18966-18971. 42. Pang, T.; Park, T.; Young, R., Mutational analysis of the S21 pinholin. Molecular Microbiology 2010, 76 (1), 68-77. 43. Blasi, U.; Fraisl, P.; Chang, C. Y.; Zhang, N.; Young, R., The C-terminal sequence of the lambda holin constitutes a cytoplasmic regulatory domain. Journal of Bacteriology 1999, 181 (9), 2922-2929. 44. Deaton, J.; Savva, C. G.; Sun, J. C.; Holzenburg, A.; Berry, J.; Young, R., Solubilization and delivery by GroEL of megadalton complexes of the lambda holin. Protein Science 2004, 13 (7), 1778-1786. 45. Gruendling, A.; Smith, D. L.; Blaesi, U.; Young, R., Dimerization between the holin and holin inhibitor of phage lambda. Journal of Bacteriology 2000, 182 (21), 6075-6081. 46. Gruendling, A.; Blaesi, U.; Young, R., Genetic and biochemical analysis of dimer and oligomer interactions of the lambda S holin. Journal of Bacteriology 2000, 182 (21), 6082-6090. 47. Grundling, A.; Blasi, U.; Young, R., Biochemical and genetic evidence for three transmembrane domains in the class I holin, lambda S. Journal of Biological Chemistry 2000, 275 (2), 769-776. 48. Grundling, A.; Smith, D. L.; Blasi, U.; Young, R., Dimerization between the holin and holin inhibitor of phage lambda. Journal of Bacteriology 2000, 182 (21), 6075-6081. 49. Young, R.; Blasi, U., Holins - form and function in bacteriophage lysis. Fems Microbiology Reviews 1995, 17 (1-2), 191-205. 50. Xu, M.; Struck, D. K.; Deaton, J.; Wang, I. N.; Young, R., A signal-arrest-release sequence mediates export and control of the phage P1 endolysin. Proceedings of the National Academy of Sciences of the United States of America 2004, 101 (17), 6415-6420.

51. Xu, M.; Arulandu, A.; Struck, D. K.; Swanson, S.; Sacchettini, J. C.; Young, R., Disulfide isomerization after membrane release of its SAR domain activates P1 lysozyme. Science 2005, 307 (5706), 113-117. 52. Kuty, G. F.; Xu, M.; Struck, D. K.; Summer, E.; Young, R., Making and breaking disulfides in Sar-endolysins: Lessons in enzyme regulation. Abstracts of the General Meeting of the American Society for Microbiology 2007, 107, 416-416. 53. Sun, Q. G.; Kuty, G. F.; Arockiasamy, A.; Xu, M.; Young, R.; Sacchettini, J. C., Regulation of a muralytic enzyme by dynamic membrane topology. Nature Structural & Molecular Biology 2009, 16 (11), 1192-1194. 54. Pang, T.; Park, T.; Young, R., Mapping the pinhole formation pathway of S21. Molecular Microbiology 2010, 78 (3), 710-719. 55. Ahammad, T.; Drew, D. L.; Sahu, I. D.; Serafin, R. A.; Clowes, K. R.; Lorigan, G. A., Continuous Wave Electron Paramagnetic Resonance Spectroscopy Reveals the Structural Topology and Dynamic Properties of Active Pinholin S(21)68 in a Lipid Bilayer. Journal of Physical Chemistry B 2019, 123 (38), 8048-8056. 56. Drew, D. L., Jr.; Ahammad, T.; Serafin, R. A.; Butcher, B. J.; Clowes, K. R.; Drake, Z.; Sahu, I. D.; McCarrick, R. M.; Lorigan, G. A., Solid phase synthesis and spectroscopic characterization of the active and inactive forms of bacteriophage S21 pinholin protein. Analytical biochemistry 2018, 567, 14-20. 57. Drew, D. L., Jr.; Butcher, B.; Sahu, I. D.; Ahammad, T.; Dixit, G.; Lorigan, G. A., Active S2168 and inactive S21IRS pinholin interact differently with the lipid bilayer: A 31P and 2H solid state NMR study. Biochimica et biophysica acta. Biomembranes 2020, 183257-183257. 58. Pang, T.; Fleming, T. C.; Pogliano, K.; Young, R., Visualization of pinholin lesions in vivo. Proceedings of the National Academy of Sciences of the United States of America 2013, 110 (22), E2054-E2063. 59. Wang, I. N.; Dykhuizen, D. E.; Slobodkin, L. B., The evolution of phage lysis timing. Evolutionary Ecology 1996, 10 (5), 545-558. 60. Hubbell, W. L.; Gross, A.; Langen, R.; Lietzow, M. A., Recent advances in site-directed spin labeling of proteins. Current Opinion in Structural Biology 1998, 8 (5), 649-656. 61. Klug, C. S.; Feix, J. B., Methods and applications of site-directed spin Labeling EPR Spectroscopy. Biophysical Tools for Biologists: Vol 1 in Vitro Techniques 2008, 84, 617-658.

62. Sahu, I. D.; McCarrick, R. M.; Lorigan, G. A., Use of Electron Paramagnetic Resonance To Solve Biochemical Problems. Biochemistry 2013, 52 (35), 5967-5984. 63. Altenbach, C.; Froncisz, W.; Hemker, R.; McHaourab, H.; Hubbell, W. L., Accessibility of nitroxide side chains: Absolute Heisenberg exchange rates from power saturation EPR. Biophysical Journal 2005, 89 (3), 2103-2112. 64. Pyka, J.; Ilnicki, J.; Altenbach, C.; Hubbell, W. L.; Froncisz, W., Accessibility and dynamics of nitroxide side chains in T4 lysozyme measured by saturation recovery EPR. Biophysical Journal 2005, 89 (3), 2059-2068. 65. Hubbell, W. L.; Cafiso, D. S.; Altenbach, C., Identifying conformational changes with site-directed spin labeling. Nature Structural Biology 2000, 7 (9), 735-739. 66. Altenbach, C.; Marti, T.; Khorana, H. G.; Hubbell, W. L., Transmembrane protein- structure - spin labeling of bacteriorhodopsin mutants. Science 1990, 248 (4959), 1088-1092. 67. Columbus, L.; Hubbell, W. L., A new spin on protein dynamics. Trends in Biochemical Sciences 2002, 27 (6), 288-295.

Chapter 2

CW-EPR Spectroscopy Reveals the Structural Topology and Dynamic Properties of Active Pinholin S2168 in a Lipid Bilayer

Tanbir Ahammad, Daniel L. Drew Jr., Indra D. Sahu, Rachel A. Serafin, Katherine R. Clowes, Gary A. Lorigan

Department of Chemistry and Biochemistry, Miami University, Oxford, OH, 45056, USA.

Published in Journal of Physical Chemistry B 2019, 123, 38, 8048-8056

DOI: 10.1021/acs.jpcb.9b06480

I carried out most of the experiments, data analysis, and wrote the manuscript. Dr. Sahu, and Dr. Drew trained me on the instrumentation and data analysis. Serafin and Clowes helped with the sample preparation.

2.1 Abstract Pinholin S2168 is an essential part of the phage 21 lytic protein system to release the virus progeny at the end of the infection cycle. It is known as the simplest natural timing system for its precise control of hole formation in the inner cytoplasmic membrane. Pinholin S2168 is a 68 amino acid integral membrane protein consisting of two transmembrane domains (TMDs) called TMD1 and TMD2. Despite its biological importance, structural and dynamic information of the S2168 protein in a membrane environment is not well understood. Systematic site-directed spin labeling (SDSL) and continuous wave electron paramagnetic resonance (CW-EPR) spectroscopic studies of pinholin S2168 in DMPC (1,2-Dimyristoyl -sn-Glycero-3-Phosphocholine) proteoliposomes are used to reveal the structural topology and dynamic properties in a native-like environment. CW- EPR spectral line shape analysis of the R1 side chain for 39 residue positions of S2168 indicates that the TMDs have more restricted mobility when compared to the N and C-termini. CW-EPR power saturation data indicate that TMD1 partially externalizes from the lipid bilayer and interacts with the membrane surface, whereas TMD2 remains buried in the lipid bilayer in the active conformation of pinholin S2168. A tentative structural topology model of pinholin S2168 is also suggested based on EPR spectroscopic data reported in this study.

2.2 Introduction The most frequent cytocidal event in the biosphere is the bacteriophage infection cycle with the last stage of this cycle being bacterial cell lysis to release the mature neonate virus 1-3. Cell lysis is precisely controlled and synchronized by at least three groups of proteins 4-6. The first step of this process is the hole formation in the inner cytoplasmic membrane by the Holin, followed by the murein layer (peptidoglycan) degradation by the Endolysin, and outer membrane disruption by the Spanin complex 6-8. The entire process happens within seconds of holin triggering which occurs at an allele-specific time and concentration 1, 9. The canonical holins form a nonspecific, microscale hole in the inner cytoplasmic membrane to allow the diffusion of functionally folded endolysin to the peptidoglycan for degradation. 2, 10-13 However, some phages (e.g. phage P1 or 21) represent a significantly different and alternative class of holin which make smaller holes in comparison to the canonical holins. These nanoscale holes are large enough for depolarization of membrane potential 2, 13-15. This group of holin is responsible for the release of signal-anchor release (SAR) endolysin and is known as pinholin for the small-scale pinholes it creates 13, 14. Pinholin S21 is encoded by the S21 gene of phage 21. The S21 gene contains a dual start motif gene which encodes two proteins, the 68 amino acid long active pinholin (S2168) and the 71 amino acid long antiholin (S2171). Antiholin (S2171) is transcribed from the first codon of the S21 gene where active pinholin (S2168) is transcribed from the 4th codon with the first amino acid of S2168 is denoted as Met4. Pinholin S2168 has two TMDs connected via a short periplasmic loop, and cytoplasmic N and C-termini. Pinholin progressively accumulates in the bacterial inner cytoplasmic membrane as an inactive dimer with both TMDs remaining in the lipid bilayer. TMD1 of the active form of pinholin S2168 externalizes very quickly to the periplasm resulting in the active dimer 1, 2, 12. Within seconds of pinholin triggering, it forms heptametric holes by rapid oligomerization and reorientation of TMD2. A study of pinholin S2168 will describes the functionally and structurally unique group of holin which consists of ~900 proteins 16. The structure and functional model of pinholin S2168 has been reported by the Young group using biomolecular and functional studies 1, 2, 12, 17-19. They have also used a computational approach to predict the number of monomers and the size of the pinhole using a truncated form of pinholin S2168 (TMD1 was deleted) 1. However, dynamic information, as well as relative orientations and interactions of TMD1 and TMD2 of S2168 with lipid or other residues, were not

53 extensively studied. For the confirmation of the proposed model, further biophysical studies were recommended 2. It has been widely recognized that the function and stability of proteins are interrelated with their structural topology and dynamic properties 20-24. Hence, it is important to know the structural topology and dynamic properties of membrane proteins in their native-like environment to understand their biological functions and mechanisms. EPR spectroscopy is a unique biophysical technique to study the structural topology and dynamic properties of proteins with high sensitivity in membrane mimic conditions 20, 25-35. This study is focused on the structural topology and dynamic properties of full-length active pinholin (S2168) using electron paramagnetic resonance (EPR) spectroscopy coupled with site-directed spin labeling (SDSL). Fmoc solid phase peptide synthesis (Fmoc-SPPS) was used for sample preparation with high yield and flexibility of spin-label incorporation at any residue position of the peptide 36. In this EPR spectroscopic study of pinholin S2168, nitroxide spin label, MTSL (S-(1-oxyl-2,2,5,5-tetramethyl-2,5-dihydro-1H- pyrrol-3-yl) methyl methanesulfonothioate) was attached to a Cys side chain at specific sites using SDSL 37. This is the first biophysical study to probe the structural topology and dynamic properties of the full-length pinholin S2168. Using the combination of CW-EPR spectroscopic line shape analysis and EPR power saturation data, we propose a tentative model of the active form of S2168, where TMD2 remains incorporated in lipid bilayers while TMD1 partially externalizes or interacts with the headgroup of the lipid bilayers.

2.3 Experimental Methods 2.3.1 Solid Phase Peptide Synthesis All peptides were synthesized via optimized Fmoc solid phase peptide synthesis (SPPS) 38. A 0.1 millimole peptide synthesis was conducted on an automated CEM Liberty Blue peptide synthesizer equipped with the Discovery Bio microwave system. The synthesis was started with a preloaded TGA resin in Dimethylformamide (DMF) base solvent system where 20% (v/v) Piperidine, 20% (v/v) N, N′-Diisopropylcarbodiimide (DIC) and 10% (w/v) Oxyma in DMF were used as deprotecting, activator, and activator base, respectively. The cleavage reaction was run for at least three hours under optimized cocktail conditions (Trifluoroacetic acid (TFA) 94%, water 2.5%, 1,2-Ethanedithiol (EDT) 2.5%, triisoproylsilane (TIPS) 1%) or (TFA 85%, water 5%, Anisole 5%, TIPS 5%) to remove the resin and side chain protecting groups 38-41. 54

2.3.2 Purification and Spin Labeling The crude peptide was purified by reverse phase high-pressure liquid chromatography (RP- HPLC) using a GE HPLC system. The sample was loaded onto a C4 (10 µm) preparative column (Vydac 214TP, 250 x 22 mm), and was run with a two solvent gradient system, where the polar solvent was 100% water and the nonpolar solvent was 90% acetonitrile with 10% water. 0.1 % TFA was added to both solvents to acidify it. The gradient was started from 5% nonpolar and increased up to 98% nonpolar by optimized run time 38. The purified pinholin peptide collected from HPLC elution was lyophilized. To attach the spin label, the lyophilized pure peptide was dissolved in dimethyl sulfoxide (DMSO) with 5-fold excess of MTSL (1:5 molar ratio) and stirred for 24 hours in a dark environment. The spin-labeled (SL) peptide was lyophilized again and purified with a C4 semi-preparative column (Vydac 214TP, 250 x 10 mm), using the same solvent and gradient system to remove free MTSL and further purify the peptide sample. After each purification, the purity of the target peptide was confirmed by MALDI-TOF MS. Spin labeling efficiency was calculated ~85 to 90% using CW-EPR 38.

2.3.3 Peptide Incorporation into Proteoliposomes To mimic the membrane environment spin-labeled pinholin peptides were incorporated into DMPC (1,2-Dimyristoyl -sn-Glycero-3-Phosphocholine) proteoliposomes following the thin film method 38. Pure spin-labeled peptides were dissolved in 2,2,2-Trifluoroethanol (TFE) and mixed with pre-dissolved DMPC lipid solution in a pear-shaped flask. The organic solvent was gently evaporated by N2(g) purging to get a uniform thin film inside the pear-shaped flask. It was left under a vacuum desiccator overnight to remove any residual organic solvent. 20 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) buffer (pH ~7.0) was used to rehydrate the thin film to get the final concentration of 200 mM lipid and 200 µM peptide into proteoliposomes at a 1:1000 protein to lipid ratio to minimize the formation of oligomers. Before rehydration, both HEPES buffer and the sample flask were kept in a warm water bath for a short period of time to bring the temperature above the phase transition temperature of DMPC 42, 43. The lipid film was dispersed from the side wall by several vortexes followed by warming in a water bath. Three more freeze-thaw cycles were performed before adding glycerol. 10% glycerol was added to the sample and mixed properly. Glycerol helps the sample to remain suspended for a longer duration at room temperature without phase separation. Sample homogeneity and size of the proteoliposomes were

55 confirmed by dynamic light scattering (DLS) spectroscopy using ZETASIZER NANO Series (Malvern Instruments) at 25 °C in a disposable 40 μL micro cuvette.

2.3.4 Circular Dichroism Spectroscopy Circular Dichroism (CD) spectra for pinholin S2168 incorporated into proteoliposomes were collected using an Aviv Circular Dichroism Spectrometer (Model 435) in a quartz cuvette with a 1.0 mm path length. Pinholin S2168 incorporated proteoliposomes samples were diluted 10 times with 10 mM phosphate buffer to reduce the final HEPES buffer concentration (which gives absorbance at lower wavelength) as well as protein concentration (20 µM). Data were collected from 260 to 190 nm with an average of 10 scans per sample and 1 nm bandwidth at 25°C 38.

2.3.5 CW-EPR Spectroscopy CW-EPR spectra were collected at X-band (~9.34 GHz) using a Bruker EMX spectrometer equipped with ER041xG microwave bridge and ER4119-HS cavity at Ohio Advanced EPR Laboratory at Miami University. Each spectrum was acquired by the signal averaging of 10 scans with 3315 G central field, sweep width 150 G, 42 s field sweep, 100 kHz modulation frequency, 1 G modulation amplitude, and 10 mW microwave power. Each experiment was repeated at least three times. All experiments were carried out at room temperature.

2.3.6 Spin-label Mobility Analysis -1 The inverse line width (δ ) of the first derivative central resonance line (mI = 0) was calculated to determine the side chain mobility. The mobility parameter, δ-1 was normalized to 20, 31, 44 “scaled mobility” factor (Ms), using equation (1) .

−1 −1 δ −δi Ms = −1 −1 (1) δm −δi where δi and δm are central line widths for the most immobilized and the most mobile side chains, respectively. The mean scaled mobility (M̅̅̅̅s) was calculated by taking the average of all studied spin-labeled residues. To further explore the dynamic properties, the rotational correlation time (τ) was calculated using equation (2) 32, 33, 45-47.

h τ = K δ [√( 0 − 1) − 1] (2) ℎ−1

−10 where, K = 6.5 × 10 s, δ is the width of the center-line, and h0 and h−1 are the heights of the center and high field lines, respectively 46, 47.

2.3.7 CW-EPR Power Saturation Experiments CW-EPR power saturation experiments were performed on a Bruker EMX X-band spectrometer coupled with ER 041XG microwave bridge and ER 4123D CW-Resonator (Bruker BioSpin). Experimental setups were optimized following the published literature 28, 34, 35, 45, 48. Samples were loaded into gas permeable TPX capillary tubes with a total volume of 3-4 μL at a concentration of 100-150 μM 48-51. EPR spectroscopic data were collected using a modulation amplitude of 1.0 G, a modulation frequency of 100 kHz, field sweep 42 s, and a sweep width 90 G. Incident microwave powers were varied from 0.06 mW to 159 mW. 3 to 5 scans were signal averaged at each microwave power. For each spin-labeled site, the spectra were recorded under three equilibrium conditions. At first, the samples were equilibrated with a lipid-soluble paramagnetic relaxant (21% oxygen) followed by the equilibration with nitrogen gas (as control), and equilibration with a water-soluble paramagnetic relaxant (2 mM NiEDDA) with a continuous purge of nitrogen gas 51. Each set of experiment was repeated at least three times. NiEDDA was synthesized according to the published protocol 35, 51. The samples were purged with nitrogen gas for at least one hour, at a rate of 10 mL per minute before starting nitrogen or NiEDDA data acquisition. The resonator was connected to the gas supply (air or nitrogen gas) during all measurements and all the experiments were performed at room temperature. The peak-to-peak amplitudes (A) of the first derivative mI = 0 resonance lines were extracted and plotted against the square root of the incident microwave power. These data points were then fitted according to Equation (3) 35, 50.

1 – ε (2ε− 1)P A = I√P [1 + ] (3) P1 2

where I is a scaling factor, P1 is the power where the first derivative amplitude is reduced 2 to half of its unsaturated value, and ε is a measure of the homogeneity of saturation of the resonance line. For the homogeneous and inhomogeneous saturation limits, ε = 1.5 and ε = 0.5, respectively 35 . In equation (3), I, ε, and P1⁄2 are adjustable parameters and yield a characteristic P1⁄2 value for

57 each equilibrium condition. Data analysis was performed using a MATLAB software script. The corresponding depth parameter () was calculated using equation (4) 35.

ΔP1(푂2)  = ln [ 2 ] (4) ΔP1(NiEDDA) 2 where ∆P1/2(O2) is the difference in the P1/2 values for oxygen and nitrogen equilibriums, and

∆P1/2(NiEDDA) is the difference in the P1/2 values for NiEDDA and nitrogen equilibriums.

2.4 Results Recently, the successful synthesis and spectroscopic characterization of the full-length active and inactive forms of pinholin S21 were reported by the Lorigan research group 38. For this study, the dynamic properties of pinholin S2168 were investigated by analyzing the CW-EPR spectra obtained from 39 spin-labeled positions of S2168 peptides incorporated into DMPC proteoliposomes at a mole ratio of 1:1000 to minimize the formation of oligomers. The representative DLS data is shown in supporting information Figure 2.10 for pinholin S2168 G48R1 incorporated into DMPC proteoliposomes which confirmed the homogeneity of the proteoliposomes samples. The primary amino acid sequence of the full-length active pinholin S2168 is shown in Figure 2.1(A). Residues studied with SDSL are indicated in blue. Predicted TMD1 and TMD2 are indicated by the two boxes. The predicted topology of Pinholin S2168 is adapted from the literature and shown in Figure 2.1(B) 1, 2, 12, 38. The R1 side chain attachment to the protein via a disulfide bond is shown in Figure 2.1(C) 26.

Figure 2.1: (A) Primary sequence of S2168, boxes indicate TMD1 and TMD2. The amino acid positions studied by EPR spectroscopy are shown in blue. (B) The predicted topology of S2168 is adapted from the literature 1, 2, 12, 38. TMD1 completely externalizes from the lipid bilayer and remains in the periplasm or partially externalizes and stays on the surface of the lipid bilayer, where TMD2 remains in the lipid bilayer. (C) R1 side chain shown in the dotted box which is attached to the protein through the disulfide bond of a Cys residue.

Figure 2.2 shows the representative CD spectrum of pinholin S2168 G40R1 in DMPC proteoliposomes. Two minima around 222 nm and 208 nm, and large positive peak close to 195 nm confirmed the α-helical secondary structure of pinholin S2168 with the R1 side chain.

Figure 2.2: CD spectrum of pinholin S2168 G40R1 in DMPC proteoliposomes was collected at pH 7 and 298 K. Mean Residue Ellipticity (MRE) is plotted against incident radiation wavelength. 59

2.4.1 CW-EPR Line Shape Analysis of Pinholin S2168 CW-EPR spectral analysis for a set of spin-labeled protein allows probing the structural and dynamic properties of the protein with a spatial resolution at the residue-specific level 45, 52-56. Figure 2.3 shows 39 CW-EPR spectra collected for R1 side chains placed at strategic locations of S2168 incorporated into DMPC proteoliposomes.

Figure 2.3: CW-EPR spectra of R1 side chains attached at indicated positions of S2168 by replacing the native amino acid with Cys. All spectra were normalized to the highest spectral intensity. (A) EPR spectra from the N-terminal and TMD1 of pinholin S2168. (B) EPR spectra from the loop to the C-terminal of pinholin S2168 including TMD2. CW-EPR spectra composed of multiple components were marked with (*).

All CW-EPR spectra show the conventional three 14N hyperfine peaks of the R1 side chain attached to pinholin S2168. EPR spectra from the N and C-termini show a relatively sharper central peak than the TMD regions which indicate higher mobility of the R1 side chain or protein 61 backbone in the terminal regions when compared to the TMDs residues. EPR spectra of residues within TMD1 and TMD2 show line broadening indicative of restricted mobility in those regions. Most of the spectra show a single motional component, which indicates mono-dispersion of conformational and motional properties of S2168. However, CW-EPR spectra for W23R1, Q26R1, N55R1, L56R1 (indicated by (*) in Figure 2.3) show two spectral components (rigid and fast components), which are indicative of heterogeneous dynamics at those sites 52.

Beside this qualitative information, CW-EPR line shape analysis can be used to derive quantitative dynamic information. Inversion of the width of the central resonance line (δ-1) has been used as a semiquantitative measurement of nitroxide mobility 20, 29, 57, 58. The mobility of individual residues is shown in Figure 2.4, by plotting δ-1 as a function of residue positions for pinholin S2168.

Figure 2.4: EPR mobility analysis of R1 side chain of S2168, calculated from the inverse width of the central resonance line (δ-1). Larger δ-1 value indicates increasing motion.

Both N and C-termini residues had higher mobility than residues inside the TMDs as speculated qualitatively from the line shape analysis. The loop region had intermediate or restricted mobility which can be explained by the small loop size and the junction between the two slow motional segments (TMD1 and TMD2) of S2168. The highest mobility was observed for D5R1, 62 present in the N-terminal with the δ-1 of 0.62 G-1 while the lowest mobility was 0.22 G-1 for W23R1, present in TMD1. Many more fluctuations of the δ-1 values were observed for the TMD1 residues, when compared to the δ-1 values of TMD2 residues. The δ-1 of the TMD1 varies from 0.22 G-1 to 0.59 G-1, while the δ-1 of the TMD2 varies between 0.22 G-1 to 0.36 G-1.

Scaled mobility (Ms) is another convenient method to compare the R1 dynamics for different sites relative to the most mobile and immobile sites. The Ms values were calculated using equation (1). The central line width was observed among the entire 39 residue set with the most mobile (D5R1) and immobile (W23R1) sites being 1.6 G (δm) and 4.5 G (δi), respectively. The individual Ms value as a function of residue position is shown in Figure 2.5.

Figure 2.5: Scaled mobility as a function of residue position for pinholin S2168.

From the individual Ms values, the average scaled mobility, 푀̅̅̅̅푠 was calculated which reflect the compactness of protein system, helical packing density, and local backbone motions of 20 21 a protein or segment of a protein . An overall 푀̅̅̅̅푠 value of 0.27 was determined for pinholin S 68 20 which indicates moderate to tight helical packing . For the individual TMDs, 푀̅̅̅̅푠 values were 0.34 (TMD1; 8 to 27) and 0.17 (TMD2; 36 to 56), which indicates TMD2 has tighter helical packing when compared to TMD1. 63

To further explore the dynamic properties of pinholin S2168 in a membrane, the rotational correlation time () was calculated. () is the time required for the spin-label to rotate one radian angle 45. The rotational correlation time of the R1 side chain is the result of three different rotational correlation times including the rotational correlation time of the entire protein (τR), local protein backbone fluctuations, and the flexibility of the spin-labeled side chain relative to the protein 20, 29, 59 . For membrane proteins, τR value greater than 60 ns, which is out of EPR sensitivity, can be ignored 44, 59. Therefore, the calculated τ values are the reflection of a combination of backbone fluctuations, side chains dynamic, and interactions with the surroundings. Figure 2.6 shows the calculated () values for the corresponding residues of spin-labeled pinholin S2168.

Figure 2.6: Rotational correlation time () of spin-labeled pinholin S2168 in DMPC proteoliposomes as a function of residue position.

The minimum and maximum () values were calculated to be 0.73 ns (D5R1) and 9.75 ns (W23R1), respectively. Shorter () values (below 2 ns) for N and C-termini residues suggest higher spin label motion due to the lipid-free environments, where longer () for TMDs residues suggest their restricted motion due to lipid environment and/or restricted backbone motions. Dynamic patterns observed from the τ calculation are consistent with the side chains mobility data extracted from the central line broadening shown in Figure 2.4 and 2.5. 64

2.4.2 Structural Topology with Respect to the Lipid Bilayer CW-EPR power saturation is a powerful and convenient biophysical technique to study the structural topology of proteins with respect to the lipid bilayer 35. In this study, CW-EPR power saturation experiments were performed on 31 spin-labeled pinholin S2168 samples incorporated into DMPC proteoliposomes. Representative CW-EPR power saturation curves are shown in Figure 2.7.

Figure 2.7: Representative CW-EPR power saturation curves of S2168 in DMPC proteoliposomes. (A) A12R1 and (B) Q26R1 are in TMD1 and (C) L50R1 is in TMD2. Red triangle NiEDDA, Green circle oxygen, Blue square nitrogen spectra with their fitted line from equation (3). The amplitudes of first derivative mI = 0 peak were plotted against the square root of the incident microwave power. (D) The color-coded primary sequence of S2168, where green residues are buried in the lipid bilayer and red residues are solvent exposed based on CW-EPR power saturation data. Black residues are not studied by CW-EPR power saturation experiment.

The relative power saturation profile of the oxygen and NiEDDA spectra indicate that A12R1 was more accessible to the polar relaxing reagent NiEDDA, whereas Q26R1 and L50R1 were more accessible to nonpolar oxygen. This trend implies that residue A12R1 falls outside of the lipid bilayer, whereas Q26R1 and L50R1 are buried inside the lipid bilayer. For the quantitative analysis of membrane insertion, the depth parameter (Φ) was calculated for individual residues

65 using equation (4). A positive Φ value indicates the corresponding side chain is buried inside the lipid bilayer where the negative Φ value indicates side chain is solvent exposed or outside of the lipid bilayer. Figure 2.7D shows residues buried in the lipid bilayer (green residues) or solvent exposed (red residues) as suggested by Φ value measurements. Individual depth parameter (Φ) will be examined in detail in the discussion section.

2.5 Discussion The qualitative and quantitative data reported in this study will provide a better understanding of side chain dynamics and structural topology of pinholin S2168 in DMPC proteoliposomes. The prominent appearance of two motional components for W23R1, Q26R1, N55R1, L56R1 (indicated by an * in Figure 3) can be attributed to their positions in TMD1 or TMD2, and the bulky side chains on the same side of the helix 53, 57. The R1 side chain shows different spectral components when its motion is restricted by a heterogenous interaction with the local environment 57. This heterogeneous interaction may arise from their position near the end of the helix or the hole opening 53. Interestingly, a similar trend was found for the bacterial K+ channel where residues at the beginning and the end of the helix (opening of the channel) show rigid and fast components 53. Another possible explanation for two components could be the bulky side chains on the same side of the helix (3 or 4 residues away from specific R1) which give characteristic anisotropic motion 57. W23, Q26 are present in the vicinity of Y22, W23, Q26, and Q30, which can also contribute to the multi-components observed in the W23R1 and Q26R1 spectra. Similar reasoning is applicable for the N55R1, L56R1 EPR spectra, which are located at the interface of nonpolar lipid and polar solvent environments and bulky side chains were present on the same side of the helix (Y52, 59K). In the active form of pinholin S2168, TMD1 was predicted to be externalized from the lipid bilayer, whereas TMD2 remains incorporated into the lipid bilayer 1, 2, 12, 18. It was expected that the mobility of TMD1’s residues should be higher than the mobility of TMD2’s residues due to the protein segmental motion and lipid-free environment. The EPR data collected in this work indicated that only the N-terminal of TMD1 (S8 to Y13) had higher mobility when compared to TMD2, but residues G14 to W27 of TMD1 had no significant difference in mobility when compared to TMD2 residues. Although, the average scaled mobility (푀̅̅̅̅푠) of TMD1 (0.34) was significantly higher than TMD2 (0.17). Again, the 푀̅̅̅̅푠 values allow comparing the protein of

20, 31, 44 interest with other protein systems studied independently . The 푀̅̅̅̅푠 value for TMD2 of pinholin S2168 is comparable with other channel-forming proteins, like Bacteriorhodopsin (BR) and Annexin XII, which had tight helical packing with the 푀̅̅̅̅푠 values of 0.12 and 0.19, respectively, 60 for the channel forming region . The 푀̅̅̅̅푠 value of TMD2 is harmonic with the predicted small 21 pore size of S 68. This 푀̅̅̅̅푠 value implies tight, but somewhat flexible helical packing to allow interconversion of homotypic TMD2-TMD2 interaction to heterotypic TMD2- TMD2 interaction as reported by Young Lab 1, 20.

To resolve the relative orientation and interaction of TMD1 and TMD2 with respect to the lipid bilayer, we calculated the depth parameter (Φ) values and plotted them as a function of residue positions in Figure 2.8.

Figure 2.8: Membrane depth parameter () as a function of S2168 residue positions in DMPC proteoliposomes. Positive  values (green) indicate that the R1 side chains are embedded inside the lipid bilayer and negative  values (red) indicate that the R1 side chains are solvent exposed.

The higher negative Φ values suggested that the N and C- termini residues (e.g. D5R1, S8R1, E62R1, A67R1) were highly solvent accessible and not buried in the lipid bilayer. It was

67 predicted that all the residues from TMD1 would show negative Φ values based on the assumption that TMD1 is externalized from the lipid bilayer as proposed by Park et. al 2. Negative Φ values for residues 5 to 14 indicate that the N-terminal domain of TMD1 was solvent exposed and externalized from the lipid bilayer. However, residues 24 to 27 of TMD1 showed positive Φ values, which implies these residues were embedded in the lipid bilayer. Residues from the middle portion of TMD1 were partially solvent exposed (G14R1, A17R1, G18R1, A20R1) or buried in the lipid bilayer (T15R1, S16R1, S19R1). It is worth mentioning here that residues T15, S16, S19 are on the opposite side of the helix than G14, A17, G18, A20 1. This indicates that the middle portion of TMD1 might be remained on the surface of the lipid bilayer, whereas some side chains were pointing out of the lipid bilayer and others were buried in the phospholipid membrane. All residues of TMD2 (36 to 56) of pinholin had positive  values, which confirmed the membrane immersion of this segment. The first residue of TMD2 (W36) had a Φ value 0.60, which gradually increased and reached to the maximum 2.56 (F49), then gradually decreased to the end of the TMD2 (L56, Φ = 0.23). This is the characteristic pattern for a membrane-spanning segment of a protein or a membrane-embedded channel forming-segment of a protein 53. Although V41, S44, G48, N55 are located inside of the lipid bilayer, they have shown relatively lower Φ values than their neighboring residues. This can be explained by the fact that all these residues have been identified as hole lining residues or on the same side of the hole as proposed by the Young lab 1.

Based on the  values and mobility data, it can be inferred that instead of complete externalization of TMD1, this helical domain is partially externalized from the lipid bilayer where certain residues of TMD1 were exposed to solvent and others were buried in the lipid bilayer. This partial externalization can be explained by the presence of Trp (W27) at the C-terminal of TMD1. The smaller  value for W27R1 implies that it was in the lipid-water interface. The anchoring effect of Trp residue has been widely recognized when it is located at the polar-nonpolar interface of a membrane 61, 62. The partial externalization and orientation of TMD1 can be further explained by the amphipathic nature of TMD1. Thermodynamically it may be more favorable for an amphipathic helix like TMD1, initially buried in the membrane, to stay in a polar and nonpolar interface instead of in a fully polar environment. Again, partial externalization of the TMD1 may be more favorable with less steric hindrance at the hole opening. In the active dimer of S2168, the hydrophilic faces

68 of two TMD2s interact with each other by a homotypic TMD2-TMD2 interaction, which is facilitated by a homotypic TMD1-TMD1 interaction 2. However, the homotypic TMD2-TMD2 interaction is converted to heterotypic interaction in their heptameric form 1. This heterotypic interaction is a common phenomenon for channel proteins having a glycine-zipper 1, 63. Similar glycine zipper (G10xxxG14xxxG18) present in TMD1 may facilitate heterotypic interaction with other TMD1 via oligomerization. Heterotypic TMD1-TMD1 interaction will be more favorable if TMD1 is lying on the surface and interacting with neighboring TMD1(s). Park et al. proposed the complete externalization of TMD1 based on their disulfide-linked dimer and protease sensitivity for S16C mutant which can be still possible at partial externalization condition where the S16 residue will be outside or surface of the lipid bilayer 2.

Based on the EPR spectroscopic data presented in this study and considering the literature model for S2168, we are proposing a tentative structural topology model of pinholin S2168 as shown in Figure 2.9.

Figure 2.9: Proposed topology of active pinholin S2168 after partial externalization of TMD1 from the lipid bilayer. The red amino acids represent solvent exposed and the green amino acids represent the lipid buried residues based on CW-EPR power saturation data. Black letters are for those residues which were not studied via EPR power saturation. 69

In the active conformation of S2168, the N-terminal remains in the periplasm and TMD1 lays on the surface of the lipid bilayer with some residues pointing out of the lipid bilayer and other residues buried in the lipid environment. TMD2 remains incorporated in the lipid bilayer where one side of the helix is facing towards the pore and another side towards the lipid environment. The C-terminal of the S2168 remains in the cytoplasm. Future biophysical experiments are needed to confirm the proposed topology model. Double electron-electron resonance (DEER) EPR spectroscopic technique can be used to measure the distance between different segments of the S2168 to fine-tune the proposed topology.

2.6 Conclusion This study reported on the structural topology and dynamic properties of the phage 21 lytic protein, pinholin S2168, in a lipid bilayer using EPR spectroscopic techniques. The CD data confirmed that pinholin S2168 maintains a predicted native α-helical secondary structure in DMPC proteoliposomes. R1 SDSL scanning of pinholin S2168 suggested that the N and C-termini have higher mobility when compared to the two TMDs. The result for EPR power saturation indicates the partial externalization of TMD1 from the lipid bilayer, whereas TMD2 remains in the lipid bilayer. The structural topology model of S2168 presented in this study will be useful for future structural studies of pinholin as well as other holin systems using biophysical techniques.

2.7 Supporting Information

Figure 2.10: Representative DLS spectrum of pinholin S2168 G48R1 incorporated into DMPC proteoliposomes. The size distribution by intensity is shown on a log scale using Igor Pro which demonstrates the homogenous size distribution of proteoliposomes sample.

Acknowledgment

We would like to thank Dr. Robert McCarrick for his continuous support for EPR instruments and data analysis. We are also grateful to the members of the Ry Young group at Texas A&M University for their experimental suggestions. This work was generously supported by the NIGMS/NIH Maximizing Investigator’s Research Award (MIRA) R35 GM126935, the NSF CHE-1807131 grant, the NSF (MRI-1725502) grant, the Ohio Board of Regents, and Miami University. Gary A. Lorigan would also like to acknowledge support from the John W. Steube Professorship.

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Chapter 3

Structural Dynamics and Topology of the Inactive Form of S21 Holin in a Lipid Bilayer Using CW-EPR Spectroscopy

Tanbir Ahammad†, Daniel L. Drew Jr. †, Rasal H. Khan†, Indra D. Sahu†‡, Emily Faul†, Tianyan Li†, Gary A. Lorigan†

†Department of Chemistry and Biochemistry, Miami University, Oxford, OH, 45056, USA.

‡Natural Science Division, Campbellsville University, Campbellsville, KY, 42718, USA.

Published in Journal of Physical Chemistry B 2020, 124, 26, 5370-5379

DOI: 10.1021/acs.jpcb.0c03575

I carried out most of the experiments, data analysis, and wrote the manuscript. Dr. Sahu, and Dr. Drew helped me on the instrumentation and data analysis. Khan, Faul, and Li helped with the sample preparation.

3.1 Abstract The bacteriophage infection cycle plays a crucial role in recycling the world’s biomass. Bacteriophage devised various cell lysis systems to strictly control the length of the infection cycle for the efficient phage life cycle. Phages evolved with lysis protein systems which can control and fine-tune the length of this infection cycle depending on the host and growing environment. Among these lysis proteins, holin controls the first and rate-limiting step of host cell lysis by permeabilizing the inner membrane at an allele-specific time and concentration hence known as the simplest molecular clock. Pinholin S21 is the holin from phage Φ21 which defines the cell lysis time through a predefined ratio of active pinholin and antipinholin (inactive form of pinholin). Active pinholin and antipinholin fine-tune the lysis timing through structural dynamics and conformational changes. Previously we reported the structural dynamics and topology of active pinholin S2168. Currently, there is no detailed structural study of the antipinholin using biophysical 21 techniques. In this study, the structural dynamics and topology of antipinholin S 68IRS in DMPC proteoliposomes is investigated using Electron paramagnetic resonance (EPR) spectroscopic techniques. Continuous-wave (CW) EPR line shape analysis experiments of 35 different R1 side 21 chains of S 68IRS indicated restricted mobility of the transmembrane domains (TMDs) which were predicted to be inside the lipid bilayer when compared to the N- and C-termini R1 side chains. In addition, R1 accessibility test performed on 24 residues using CW-EPR power saturation 21 experiment indicated that TMD1 and TMD2 of S 68IRS were incorporated into the lipid bilayer where N- and C-termini were located outside of the lipid bilayer. Based on this study, a tentative 21 model of S 68IRS is proposed where both TMDs remain incorporated in the lipid bilayer, N- and C-termini were located outside of the lipid bilayer. This work will pave the way for the further studies of other holins using biophysical techniques and will give structural insight for these biological clocks in molecular detail.

3.2 Introduction Much of the world’s biomass is recycled by the bacteriophage infection cycle which is repeating ~1028 times per day and must be precisely controlled to maintain the phage’s ability to continue this cycle.1 Bacteriophage infected gram-negative bacterial cell lysis is accomplished in a controlled way by at least three groups of phage lysis proteins. These lysis proteins include holin, endolysin, and spanin. They are responsible for the permeabilization of the inner membrane, degradation of the peptidoglycan layer and outer membrane, respectively.2-4 The proper timing of infected host cell lysis is crucial for the phage life cycle.5 Holin plays significant roles to define the length of the phage infection cycles.5, 6 Consequently, it is subjected to immense evolutionary pressure to achieve the optimum lysis time.6 Holins are the gatekeeper of the infected cell lysis process and regulated by several kinds of protein inhibitors.6 Most of the holins are co-expressed with an inhibitory holin called antiholin.5, 7 Although some of the holins maintain their regulatory function without known antiholin or in the absence of complementary antiholin, it is well accepted that the precise timing of these simplest biological/molecular clocks is attributed to their corresponding antiholins.6 Some groups of holin (e.g. S105, S21) were extensively studied with an emphasis on the active forms of holin due to its direct biological significance.1, 7-14 However, structural, dynamic, and topology studies of antiholins are less studied despite their significant difference in structure and topology. Pinholin S21 encoded by the S21 gene of phage Φ21 is significantly different from canonical holins. It makes very small holes in comparison to the canonical holins and destroys the proton motive force to release and activate the signal-anchor release (SAR) endolysin at an allele- specific time and concentration.1, 15 The S21 is a dual start motif gene which encodes the 71 amino acids long antipinholin (S2171) or the 68 amino acids long pinholin (S2168), each containing two putative transmembrane domains (TMDs). Both TMDs are incorporated into the inner cytoplasmic membrane, keeping the N and C-termini in the cytoplasm. However, TMD1 of S2168 externalizes very quickly from the inner membrane, exposing itself to the periplasm. The extra three amino acids in the N terminal of S2171 add an extra positive charge which delays the externalization of TMD1 from the lipid bilayer. Some studies have indicated that pinholin makes homo/hetero dimers and remains inactive unless both TMD1s of a dimer are externalized from the lipid bilayer to make active dimers.10 Hence S2171 delays the formation of the active dimer which is a prerequisite for

80 the pinhole formation.10, 12 A proper combination of active pinholin and antipinholin gives precise timing of host cell lysis.6 Although antipinholin S2171 delays the holin triggering time, TMD1 still externalizes after an initial delay and behaves like active pinholin which makes it difficult to study the structural topology of antipinholin S2171. Ry Young’s group reported a modified form of antipinholin by adding five extra amino acids (RYIRS) after the Methionine-4 (M4) of the active pinholin, referred 21 12 to as S 68IRS. This ‘RYIRS’ tag prevents the externalization of TMD1 from the lipid bilayer due to the bulky side chains and two extra positive charges. It is reported as a dominant inhibitor of pinholin which makes it more feasible for the study of the inactive form of pinholin S21.12 In 21 this study, this inactive analog of pinholin (S 68IRS) is used to study the structural dynamics and topology of antipinholin of phage Φ21. Most of the studies reported on holin and pinholin were conducted using biomolecular and functional techniques.1, 7-14 However, structural dynamics, conformational changes, and topology of antipinholin have not been extensively studied using a variety of biophysical techniques. It is well recognized that proteins’ stability and functions are coined in its structural topology and dynamics relationship.16-20 As a membrane protein, it is important to know the structure and dynamics of this protein in a lipid environment. The study of membrane proteins in the presence of a lipid environment is very challenging using conventional biophysical techniques such as X- ray crystallography and NMR spectroscopy.21-24 Electron paramagnetic resonance (EPR) spectroscopy is a powerful biophysical technique used to study the structural dynamics and topology of membrane proteins in lipid environments with higher sensitivity and without any size limitation.16, 25-35 21 In this study, the structural dynamics and topology of antipinholin (S 68IRS) in proteoliposomes were investigated using EPR spectroscopic techniques. The full-length 21 antipinholin (S 68IRS) was synthesized using Fmoc solid-phase peptide synthesis (Fmoc-SPPS) for convenient spin labeling and high yield protein sample preparation. Nitroxide spin-label, MTSL (S-(1-oxyl-2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrol-3-yl) methyl ethanesulfonothioate) was attached site-specifically using site-directed spin labeling (SDSL) to make an EPR active antipinholin construct.36, 37 Based on our CW-EPR spectroscopic line shape analysis and EPR 21 power saturation data, a tentative structural topology model of the antipinholin S 68IRS is

81 proposed, where both TMD1 and TMD2 remains incorporated into the lipid bilayers while the N- terminal, C-terminal, and loop regions remained solvent-exposed.

3.3. Experimental Methods 3.3.1 Peptide Synthesis and Purification All peptides were synthesized on an automated CEM Liberty Blue peptide synthesizer equipped with the Discovery Bio microwave system via optimized Fmoc solid-phase peptide synthesis (SPPS) reported in previous studies.37, 38 Each synthesis was started with a Glutamate preloaded TGA resin in the dimethylformamide (DMF) based solvent system. 20% (v/v) Piperidine in DMF was used as a deprotector to remove the Fmoc protecting group before each coupling cycle. During each coupling cycle, 0.2 M amino acid was added to the reaction vessel in the presence of 15.6% (v/v) N, N′-diisopropylcarbodiimide (DIC), and 14.2% (w/v) oxyma as an activator, and activator base, respectively. After successful synthesis, the cleavage reaction was run for at least three hours under optimized cocktail conditions followed by N2 (g) evaporation and ether precipitation.37-40 Peptide pellets were washed three more times with ice-cool ether followed by centrifugation and precipitated peptide was lyophilized to get fluffy and easy to solubilize powder peptide. The crude peptide was purified by reverse-phase high-performance liquid chromatography (RP-HPLC) using a GE HPLC system coupled with a C4 (10 µm) preparative column (Vydac 214TP, 250 x 22 mm). A two solvent gradient system was used, where the polar solvent was 100% water and the nonpolar solvent was 90% acetonitrile with 10% of water. 0.1 % TFA was added to both solvents to acidify it. To attach the spin-label to the peptide, the lyophilized pure peptide was dissolved in dimethyl sulfoxide (DMSO) with a 5-fold excess of MTSL (1:5 molar ratio) and stirred for 24 hours in a dark environment. The spin-labeled (SL) peptide was lyophilized again and purified with a C4 semi-preparative column (Vydac 214TP, 250 x 10 mm), using the same solvent and gradient system to remove free MTSL and further purify the peptide sample. After each purification, the purity of the target peptide was confirmed by MALDI-TOF MS. Spin labeling efficiency was calculated ~85 to 90% using CW-EPR.38

3.3.2 Peptide Incorporation into Proteoliposomes To mimic the membrane environment spin-labeled antipinholin peptides were incorporated into DMPC (1,2-Dimyristoyl -sn-Glycero-3-Phosphocholine) proteoliposomes following the thin

82 film method.37, 38 In brief, pure spin-labeled peptide was dissolved in 2,2,2-Trifluoroethanol (TFE) and mixed with pre-dissolved DMPC lipid solution in a pear-shaped flask. The organic solvent was gently evaporated by N2(g) purging to get a uniformly thin film inside the pear-shaped flask followed be overnight vacuum desiccation to remove any residual organic solvent. A 10 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) buffer (pH ~7.0) was used to rehydrate the thin film to get the final concentration of 200 mM lipid and 200 µM peptide within the proteoliposomes sample. Before rehydration, both HEPES buffer and the sample flask were kept in a warm water bath for a short period of time to bring the temperature above the phase transition temperature of DMPC.37, 41, 42 The lipid film was dispersed from the flask sidewall by vortexing several times followed by warming in a hot water bath. At least three freeze-thaw cycles were performed before adding glycerol. Glycerol was added to give a final concentration of 10% and mixed thoroughly which helps the sample remain suspended for a longer duration at room temperature without phase separation. Sample homogeneity and proteoliposomes size were confirmed by using dynamic light scattering (DLS) spectroscopy (ZETASIZER NANO Series; Malvern Instruments) at 25 °C in a disposable 40 μL micro cuvette.

3.3.3 Circular Dichroism Spectroscopy 21 Circular Dichroism (CD) data for antipinholin S 68IRS were collected using an Aviv Circular Dichroism Spectrometer (Model 435) in a quartz cuvette with a 1.0 mm path length. The pure peptide sample was dissolved in TFE to obtain the CD spectra in the solution form. Data were collected from 260 to 200 nm with an average of 3 scans per sample and 1 nm bandwidth at 25°C.

3.3.4 CW-EPR Spectroscopy CW-EPR spectra were collected at X-band (~9.34 GHz) with a Bruker EMX spectrometer equipped with ER041xG microwave bridge and ER4119-HS cavity at Ohio Advanced EPR Laboratory at Miami University. Each spectrum was acquired by signal averaging 10 scans with 3315 G central field, sweep width 150 G, 42 s field sweep, 100 kHz modulation frequency, 1 G modulation amplitude, and 10 mW microwave power.37 Each experiment was repeated at least three times at room temperature.

3.3.5 Spin-label Mobility Analysis -1 The inverse line width (δ ) of the first derivative central resonance line (mI = 0) was calculated to determine the side chain mobility. To further explore the dynamic properties, the rotational correlation time (τ) was calculated using equation (1):32, 33, 43-45

h τ = K δ [√( 0 − 1) − 1] (1) ℎ−1

−10 where K = 6.5 × 10 s, δ is the width of the central line, and h0 and h−1 are the heights of the center and high field lines, respectively.44, 45

3.3.6 CW-EPR Power Saturation Experiments CW-EPR power saturation experiments were performed on a Bruker EMX X-band spectrometer coupled with ER 041XG microwave bridge and ER 4123D CW-Resonator (Bruker BioSpin). Experimental setups were optimized following published literature articles.28, 34, 46 Samples were loaded into gas permeable TPX capillary tubes with a total volume of 3-4 μL at a concentration of 100-150 μM.46-49 EPR spectroscopic data was collected using a modulation amplitude of 1.0 G, a modulation frequency of 100 kHz, 42 sec field sweep, and 90 G sweep width. Incident microwave powers were varied from 0.06 mW to 159 mW. At each microwave power, three to five scans were taken for signal-averaging. For each spin-labeled site, the spectra were recorded under three equilibrium conditions and at each condition, experiments were repeated at least three times. At first, the samples were equilibrated with a lipid-soluble paramagnetic relaxant (21% oxygen) followed by equilibration with nitrogen gas (as a control), and equilibration with a water-soluble paramagnetic relaxant (2 mM NiEDDA) with a continuous purge of nitrogen gas.37, 49 NiEDDA was synthesized according to the published protocol.35, 49 The samples were purged with nitrogen gas for at least one hour, at a rate of 10 mL per minute before starting nitrogen or NiEDDA data acquisition. The resonator was connected to the gas supply (air (as a source of 21 % oxygen) or nitrogen gas) during all measurements and all the experiments were performed at room temperature. The peak-to-peak amplitudes (A) of the first derivative mI = 0 resonance lines were extracted and plotted against the square root of the incident microwave power. These data points were then fitted according to Equation (2):35, 48

1 – ε (2ε− 1)P A = I√P [1 + ] (2) P1 2

where I is a scaling factor, P1⁄2 is the power where the first derivative amplitude is reduced to half of its unsaturated value, and ε is a measure of the homogeneity of saturation of the resonance line. For the homogeneous and inhomogeneous saturation limits, ε = 1.5 and ε = 0.5, respectively.35 In equation (2), I, ε, and P1⁄2 are adjustable parameters and yield a characteristic P1⁄2 value for each equilibrium condition. Data analysis was performed using a MATLAB software script. The corresponding depth parameter () was calculated using equation (3):35

ΔP1(푂2)  = ln [ 2 ] (3) ΔP1(NiEDDA) 2 where ∆P1/2(O2) is the difference in the P1/2 values for oxygen and nitrogen equilibriums, and

∆P1/2(NiEDDA) is the difference in the P1/2 values for NiEDDA and nitrogen equilibriums.

3.4 Results Recently, the structural topology and dynamic properties of active pinholin S2168 was reported using CW-EPR spectroscopic techniques.37 In this study, the structural dynamics and topology of 21 the antipinholin S 68IRS peptide incorporated into DMPC proteoliposomes were investigated using CW-EPR spectral line shape analysis and power saturation experiments . The spin-labeled positions were judiciously selected based on published literature and initial experimental data to minimize structural perturbations.10 Figures 3.1A and 3.1B are the primary sequence of the S2171and S2168, which are wild type antipinholin and active pinholin respectively as a quick reference and comparison with reporting antipinholin allele. Figure 3.1C shows the primary 21 21 sequence of the inactive analog of pinholin S (S 68IRS) where the spin-label positions are indicated in blue. MTSL was attached to the Cys side chain where Cys replaced the native amino acid in the site-specific position. MTSL attached to Cys side chain, known as R1 is shown in Figure 21 1, 3.1D. The predicted topology of S 68IRS is adapted from the literature and shown in Figure 3.1E. 10, 12, 38, 50

Figure 3.1: Primary sequence and topology of S21. Primary sequences of wild type antipinholin 21 21 21 21 S 71 (A), wild type active pinholin S 68 (B) and the inactive analog of pinholin S (S 68IRS) (C). The amino acid positions studied by EPR spectroscopy are shown in blue. The inactive tag is color- coded red and indicating its incorporation site by the arrow. The helical region of TMD1 and TMD2 are indicated by blue and green boxes, respectively. (D) MTSL attached to Cys side chain 21 of the protein via a disulfide bond (also known as R1). (E) The predicted topology of S 68IRS where both TMDs remain incorporated in the lipid bilayer and TMD1 has not been externalized from the lipid bilayer.10, 12, 15, 37 Pure spin-labeled peptide samples were obtained by double purification using RP-HPLC and confirmed by MALDI-TOF MS. A representative HPLC and MS spectra are shown in supporting information (SI) Figure 3.10. Proteoliposomes sample size and homogeneity were investigated using DLS. A representative DLS spectrum is shown in Figure 3.2. It shows homogenous proteoliposomes with maximum intensity around 124 nm.

21 Figure 3.2: DLS spectrum for S 68IRS F49R1 incorporated into the DMPC proteoliposomes. Signal intensity is plotted as a log function of particle diameter.

Proper folding and alpha-helical secondary structure of the peptide samples were examined by 21 CD measurements. CD spectra were obtained for the antipinholin S 68IRS with and without spin- label in TFE solution to confirm that the spin-label incorporation has no significant effects on the 21 secondary structure. Figure 3 shows representative CD spectra for antipinholin S 68IRS without 21 spin-label (black solid line) and antipinholin S 68IRS F24R1, spin labeled at F24 position (blue solid line). In Figure 3.3, both spectra exhibit two minima at 222 nm and 208 nm which suggest that both samples have -helical secondary structures in the TFE solution.51 These CD data confirmed that nitroxide spin labeling has no significant effect on the global secondary structure.

21 Figure 3.3: Circular Dichroism spectra for inactive pinholin S 68IRS without spin-label (black) 21 and S 68IRS F24R1, with spin-label (blue). Spectra were signal averaged for three scans. Mean residue molar ellipticity (MRE) is plotted against incident radiation wavelength. 21 3.4.1 Structural dynamics of antipinholin S 68IRS CW-EPR spectral analysis is a powerful biophysical technique used to decipher the structural dynamic properties of a protein at a residue-specific level with spatial resolution.43, 52-56 21 To probe the structural dynamics of S 68IRS, a total of 35 CW-EPR spectra of the R1 side chain 21 were analyzed which were placed at strategic locations of S 68IRS indicated in Figure 3.1C. Figure 3.4 shows representative CW-EPR spectra collected for R1 side chains placed at the 21 N-terminal, TMD1, TMD2, and C-terminal regions of S 68IRS in DMPC proteoliposomes.

Figure 3.4: Representative CW-EPR spectra with R1 situated at the indicated positions. Here, blue EPR spectra represent TMD1 residues, green for TMD2, and black indicating loop and terminal regions. All spectra were normalized to the highest spectral intensity. CW-EPR spectra composed of multiple components were marked with (*).

All CW-EPR spectra show the conventional three peaks of the nitroxide spin label due to the 14N hyperfine splitting. Higher mobility of the R1 side chain in the terminal regions was confirmed by the sharper peaks. TMD located R1 spectra, indicated restricted mobility by virtue of the broader central peak. Most of the EPR spectra showed a single motional component. However, some spectra (e.g. W23R1, G40R1, G48R1, T51R1, N55R1, Y57R1) showed two spectral components (rigid/slower and fast/higher motional components), which are indicative of heterogeneous dynamics of the R1 spin label.52 CW-EPR spectra with a significant population of rigid component (W23R1, N55R1, Y57R1 ) are indicated by (*) in Figure 3.4. Different motional components can be observed on the R1 spectra when the motion of spin label is restricted by a heterogeneous interaction with the local environment.57 These residues are close to the end of TMD1/TMD2 and near the lipid/solvent interface which could induce heterogeneous interaction.53 Another reason may be the presence of the bulky side chains on the same side of the helix.57 Here Q26 and W27 have bulky side chains which are present on the same side of W23 may cause multiple motional components for the W23R1 spectrum. Similarly, Y52 with the bulky side chain present on the same side of N55 causing multiple motional components for N55R1.

21 To quantify the dynamic properties of antipinholin S 68IRS, the width of the central resonance line (δ) was measured and the inverse of the width of the central resonance line (δ-1) was calculated. Inverse width of the central resonance line (δ-1) is known as the relative mobility of the spin-label and has been used as a semiquantitative measurement of nitroxide mobility.16, 29, 57, 58 The mobility of individual residues is shown in Figure 3.5, by plotting δ-1 as a function of 21 residue positions for pinholin S 68IRS.

Figure 3.5: The relative mobility of R1 (δ-1) as a function of residue positions of the primary 21 -1 sequence of S 68IRS. Higher the value of (δ ) indicates the higher mobility of the nitroxide spin- label at that corresponding position. Here, blue closed circles represent TMD1 residues, green for TMD2, and black indicating loop and terminal regions.

Both TMDs had restricted mobility when compared to the N and C-termini residues which imply that TMDs are incorporated into the lipid bilayer where N and C-termini residues are solvent exposed and had less interaction with the surrounding environments. The highest mobility was observed for S8R1 (0.62 G-1), present in the N-terminal while lowest mobility was observed for T51R1(0.22 G-1), present in the TMD2. A greater range of values for δ-1 are observed for TMD1, when compared to that observed in TMD2, specifically in the N-terminal side of TMD1. The N- terminal of TMD1 (first 3~4 amino acids) had significantly higher mobility in comparison with rest of the helical region implying that this segment was outside of the lipid bilayer and had less interaction with surrounding environments.

21 To further quantify the relative motion of different side chains of S 68IRS, the rotational correlational time (τ) was calculated for the nitroxide spin label placed in corresponding positions 21 of S 68IRS. The rotational correlational time (τ) is the time required for the spin-label to rotate 1 radian and calculated by using the empirical formula shown in equation (1).43 The calculated τ values are indicative of backbone fluctuations, side chain dynamics, and interactions of side chain 91 with the surroundings.37, 59, 60 Figure 3.6 represents the calculated τ values for the corresponding 21 residues of spin-labeled inactive pinholin S 68IRS.

Figure 3.6: The rotational correlational time (τ) as a function of residue positions of the 21 primary sequence of S 68IRS. The same color code is used as in Figure 3.5.

Shorter τ values for N and C-termini residues suggesting higher spin label motion and imply these segments were outside of the lipid bilayer. Conversely, longer τ for TMDs residues suggest their restricted motion due to the lipid environment and/or restricted backbone motions. The minimum and maximum τ values were calculated to be 1.0 ns (D5R1, S8R1) and 8.2 ns (T51R1), respectively. Dynamic patterns observed from the τ value calculation are consistent with the side chain mobility data extracted from the central line broadening shown in Figure 3.5.

21 3.4.2 Structural topology and interaction of S 68IRS with the lipid bilayer 21 To explore the interaction and incorporation of antipinholin S 68IRS into DMPC proteoliposomes, CW-EPR power saturation experiments was carried out in the presence of lipid- soluble (oxygen) and water-soluble (NiEDDA) paramagnetic relaxants. Oxygen and NiEDDA primarily probe the relative abundance and accessibility of nitroxide spin-label in the lipid and aqueous phase, respectively.61 Relative accessibility of the particular spin-label to the lipid-soluble and water-soluble paramagnetic relaxing agents provides information on whether this spin-label is within the lipid environment or exposed to the aqueous environment. Figure 3.7 represents CW- 92

EPR power saturation curves showing the relative signal intensity and saturation pattern as a 21 function of square root of the microwave power for S 68IRS A20R1 (Figure 3.7A) and 21 A67R1(Figure 3.7B) spin-label positions. S 68IRS A20R1 required higher microwave power to reach the saturation in the presence of oxygen when compared to that in the presence of nitrogen and NiEDDA which qualitatively indicates that this position of spin-label was incorporated into the lipid bilayer and was more accessible to oxygen and less accessible to NiEDDA. Conversely, 21 S 68IRS A67R1 was more accessible to NiEDDA than oxygen indicating that this position was outside of the lipid bilayer.

(A) (B)

NiEDDA Oxygen Nitrogen

(C)

21 Figure 3.7: Representative CW-EPR power saturation curves of S 68IRS in DMPC proteoliposomes. (A) A20R1 and (B) A67R1. Red triangle represents NiEDDA, Green circle represents oxygen, Blue square represents nitrogen spectra with their fitted line from equation (2).

The amplitudes of the first derivative mI = 0 peak were plotted against the square root (SQRT) of 21 the incident microwave (MW) power. (C) The color-coded primary sequence of S 68IRS, where green residues are buried in the lipid bilayer and red residues are solvent-exposed based on CW- EPR power saturation data.

21 A total of 24 spin-labeled S 68IRS positions were studied via the CW-EPR power saturation technique. The positions are color-coded in the primary sequence of antipinholin 21 (S 68IRS) where red are the positions found outside of the lipid bilayer and green are the positions found inside the lipid bilayer. CW-EPR power saturation data clearly demonstrate that N-terminal, C-terminal, and loop regions were solvent-exposed and outside of the lipid bilayer where most of the parts of TMD1 and TMD2 were inside of the lipid bilayer.

To predict the relative orientation of TMD1 and TMD2 with respect to the lipid bilayer, the depth parameter (Φ) for individual residues was calculated using equation (3). The depth parameter (Φ) gives a quantitative analysis of the insertion of spin label into the lipid bilayer which is derived from relative biomolecular collision rates with the molecular oxygen and NiEDDA. The positive (Φ) value is proportional to the membrane insertion where the negative (Φ) value is proportional to solvent accessibility.35 Figure 3.8 shows the depth parameter (Φ) as a function of 21 the spin-label position for the S 68IRS.

21 Figure 3.8: Calculated depth parameter (Φ) as a function of S 68IRS residue positions in DMPC proteoliposomes. Positive (Φ) values (green) indicate that the R1 side chains are embedded inside the lipid bilayer and negative (Φ) values (red) indicate that the R1 side chains are solvent exposed.

Negative (Φ) values for the terminal regions confirm that N and C-termini were solvent- exposed and outside of the lipid bilayer. All the spin-label positions in TMD2 showed positive (Φ) values confirming the membrane insertion of TMD2 consistent with the previously proposed model.12 Close observation of the selected region of TMD2 shown in SI Figure 3.11 revealed a characteristic pattern (inverted ‘V’) of membrane-embedded segments of a protein.53 The positive (Φ) values of TMD2 gradually increased from a smaller value to the maximum around the center of the lipid bilayer, followed by gradually decreased to the minimum at the end of the lipid bilayer. The highest depth parameter was observed for G48R1 of TMD2 with Φ value of 1.15, suggesting this site was close to the central position in the lipid bilayer. A similar trend was observed for TMD1, although there were less data points when compare to TMD2. The highest depth parameter was 1.0 for A20R1 of TMD1. However, Some other residues located at the N- terminal side (A12) or C-terminal side (Q26, W27) of the TMD1 showing negative (Φ) values which imply that starting and end regions of the helix remain outside of the lipid bilayer. This behavior agrees with the previously proposed topology from the Ry Young Lab.10, 12

3.5 Discussion This is the first biophysical study of the inactive form of pinholin S21 using EPR spectroscopy and illustrated the relative difference in the dynamics of different segments and 21 residues of S 68IRS. Terminal regions have significantly higher mobility due to its structural flexibility and a lipid-free environment. Conversely, both TMD regions have restricted mobility, especially the residues which are predicted to be incorporated into the lipid bilayer. CW-EPR power saturation data were consistent with mobility data derived from CW-EPR line shape analysis and confirmed the membrane insertion of TMD1 and TMD2 where the N and C termini, and both ends of the helices were solvent exposed. 21 For this EPR spectroscopic study of S 68IRS, the spin-labeled positions were judiciously selected based on published literature, and initial experimental data to minimize structural perturbations.10 This selective spin labeling led to fewer data points for TMD1 when compared with TMD2. Moreover, the TMD1 is prone to mutational effects and has a natural tendency to externalize from the lipid bilayer. Some spin-labeled residues of TMD1 were found outside of the membrane which were expected to be inside of lipid bilayer (e.g. A17R1, G21R1). This may be due to the change of hydrophobicity or interruption of the TMD1-TMD2 interaction or both when the native amino acids were replaced with the R1 side chain.10 Those residues were excluded from 95 the topological analysis of antipinholin due to their structural perturbations. It is worthy to mention 21 here that previous study reported by the Young lab found activation of antipinholin S 68IRS when 21 10 they performed the mutations at some of these positions e.g. S 68IRS A17Q, G21Q. Based on the dynamic information and accessibility data reported in this study, a tentative 21 structural topology model of S 68IRS is proposed as shown in Figure 3.9. In Figure 3.9, residues are color-coded based on power saturation data where red is solvent-exposed, green is located inside of the lipid bilayer and black is not studied by power saturation. The ‘RYIRS’ tag is indicated by red filled circles. Several residues are labeled with residue positions as visualization guide. The helical tilt of TMD2 is proposed based on a computation study previously reported by the Young lab and possible positive hydrophobic mismatch for TMD2.12, 62 Further structural 21 studies are needed to confirm the proposed topology of S 68IRS, such as application of double electron-electron resonance (DEER) experiments, which could be performed to refine the structure and relative orientation of TMD1 and TMD2 in the presence of a lipid bilayer.

21 Figure 3.9: The proposed structural topology of inactive pinholin S 68IRS incorporated into a lipid bilayer. The red amino acids represent solvent exposed and the green amino acids represent the lipid buried residues based on CW-EPR power saturation data. Black letters were not studied by the EPR power saturation experiment. Red filled circles are the RYIRS tag used for the inactive conformation of pinholin S21.

3.6 Conclusion This study reported on the structural dynamics and topology of the inactive analog of 21 21 pinholin S (S 68IRS) incorporated into a DMPC lipid bilayer using EPR spectroscopic techniques. Before EPR experiments, the purity and spin labeling efficiency of all peptide samples were confirmed. It was confirmed that protein samples maintained their native secondary structure in all experimental conditions using CD spectroscopy. Spin label positions were selected 97 judiciously to minimize structural perturbation, while gaining maximum information of structural 21 21 dynamics and topology of the antipinholin S . The R1 scanning of antipinholin S 68IRS suggested that the N and C-termini have higher mobility when compared to the two TMDs. The EPR power saturation data indicate that both TMD1 and TMD2 remain in the lipid bilayer. The structural 21 topology model of S 68IRS presented in this study will be useful for future structural studies of antipinholin as well as other holin systems using EPR spectroscopic techniques. Further 21 experiments are recommended to confirm the proposed topology of S 68IRS

3.7 Supporting Information

Figure 3.10: (A) Representative HPLC Chromatogram of the antipinholin peptide. (B) Representative MALDI-TOF mass spectrum of the antipinholin with a target m/z of 8386.

Figure 3.11: Characteristic depth parameter pattern for the transmembrane helix. Calculated 21 depth parameter (Φ) as a function of S 68IRS residue positions in DMPC proteoliposomes. In TMD2, positive (Φ) values gradually increased to the maximum followed by gradually decreased to the minimum at the other end of the lipid bilayer which fit for characteristic (^) patter indicative of TM helix. TMD1 also shows similar pattern.

Acknowledgment We are grateful to the members of the Ry Young group at Texas A&M University for their experimental suggestions. We would like to thank Dr. Robert McCarrick for his continuous support for EPR instruments and data analysis. This work was generously supported by the NSF CHE-1807131 grant, the NSF (MRI-1725502) grant, the Ohio Board of Regents, and Miami University. Gary A. Lorigan would also like to acknowledge support from the John W. Steube Professorship.

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Chapter 4

Conformational Differences are Observed for the Active and Inactive Forms of Pinholin S21 Using DEER Spectroscopy

Tanbir Ahammad†, Daniel L. Drew Jr. †, Indra D. Sahu‡†, Rasal H. Khan†, Brandon J. Butcher†, Rachel A. Serafin†, Alberto P. Galende‡, Robert M. McCarrick†, Gary A. Lorigan†

†Department of Chemistry and Biochemistry, Miami University, Oxford, OH, 45056, USA.

‡Natural Science Division, Campbellsville University, Campbellsville, KY, 42718, USA.

“Manuscript is ready to submit for publication”

I carried out most of the experiments, data analysis, and wrote the manuscript. Drew carried out initial experiments. Dr. Sahu helped in data analysis and did the computational works. Khan, Butcher, and Serafin helped with the sample preparation. Galende helped in computational works.

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4.1 Abstract Bacteriophage evolved with an efficient host cell lysis mechanism to terminate the infection cycle and release the new progeny virion at the optimum time, allowing adaptation with the changing host and environment. Among the lytic proteins, holin controls the first and rate- limiting step of host cell lysis by permeabilizing the inner membrane at an allele-specific time know as ‘holin triggering’. Pinholin S21 is a prototype holin of the phage Φ21 which makes many nanoscale holes and destroys the proton motive force, which in turn activates the SAR-endolysin to degrade the peptidoglycan layer of host cell and destruction of the outer membrane by the spanin complex. Like many others holin, phage Φ21 has two holin proteins; active pinholin and antipinholin. The antipinholin form differs only by three extra amino acids at the N-termini but has a different structural topology and conformation with respect to the membrane. Predefined combinations of active pinholin and antipinholin fine-tune the lysis timing through structural dynamics and conformational changes. Previously, we reported the structural dynamics and topology of active pinholin and antipinholin (Ahammad et al. JPCB 2019, 2020). However, detailed structural studies and direct comparison of these two forms of pinholin S21 are absent in the literature. In this study, the structural topology of active pinholin (S2168) and antipinholin 21 (S 68IRS) in DMPC proteoliposomes were investigated using four-pulsed double electron-electron resonance (DEER) pulse EPR spectroscopic technique to measure the distances between TMD1 and TMD2. Using DEER spectroscopy, five inter-label distances were measured for both the active and inactive forms of pinholin. A model for the most probable structure of the active pinholin and inactive antipinholin in DMPC proteoliposomes were obtained using the experimental DEER distances coupled with the simulated annealing software package Xplor-NIH. In the proposed model, TMD2 of active pinholin remains in the lipid bilayer and TMD1 is partially externalized from the bilayer with some residues located on the surface. However, both TMDs remain 21 incorporated in the lipid bilayer for the inactive S 68IRS form. This study demonstrates for the first time, clear structural topology, and conformational difference between the two forms of pinholin S21. This work will pave the way for the further studies of other holin system using DEER spectroscopic technique and will give structural insight for these biological clocks in molecular detail.

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4.2 Introduction

Phage therapy has been revived in the current era of antimicrobial resistance to combat multidrug-resistant (MDR) bacteria.1-6 Phages and their bacteriolytic gene cassettes can serve as effective biocontrol agents for food safety and preservation.7-9 The holin-endolysin system at the center of these bacteriolytic gene cassettes has potential therapeutic and pathophysiological application in higher animals.1-5, 10-13 It is a prerequisite to understand the bacteriophage life cycle and its lytic system for the judicious use of bacteriophages. Phages devised a robust lytic system to control the length of the infection cycle in response to a changing environment and host.14 Most large, dsDNA bacteriophages evolved the holin-endolysin system coupled with the spanin, which are an efficient host lysis system instrumental to lyse gram-negative bacterial cells at an optimum time.15-18 Endolysin is the key component of this lysis machineries which degrade the peptidoglycan layer within a few seconds of reaching the target site.14, 19 However, holin is the molecular clock responsible for the precise timing of host cell lysis.15, 20-24 Holin triggers at an allele specific time and concentration to make micron-scale holes.21, 25-29 These holes are large enough to allow the fully folded functional endolysin to cross the inner membrane to reach the peptidoglycan layer.14, 30, 31 More recently, a prototype holin system was discovered in some phages, which makes smaller holes in compare to the canonical holin, and denoted as pinholin.32 Pinholins are coupled with the signal anchor release (SAR) endolysin system which maintains a membrane-tethered inactive conformation during the vegetative phase of the phage infection cycle.32 Like canonical holin, pinholin triggers at an allele-specific time and concentration to permeabilize the cytoplasmic membrane. However, in contrast to canonical holin, pinholin makes large numbers of nanoscale holes and destroys the proton motive force which accelerates the release and activation of SAR endolysin to degrade the peptidoglycan layer.32, 33 Pinholin S21 is a class-II holin, encoded by S21 gene of phage Φ21.18, 34 The S21 is a dual start motif gene; it encodes two holin proteins, a lysis effector (pinholin S2168) and lysis inhibitor (antipinholin S2171).34 Both of these proteins have two transmembrane domains (TMDs), initially which remain incorporated in the bacterial inner cytoplasmic membrane (ICM) and accumulate harmlessly as inactive dimers.30, 35 However, TMD1 of S2168 externalizes very quickly from the ICM which is the prerequisite for the active dimer and subsequent oligomerization for pinhole formation. Conversely, TMD1 of S2171externalizes slowly which in turn delays the initial

108 oligomerization and pinhole formation.30 A proper combination of active pinholin and antipinholin gives precise timing of host cell lysis.36 The functional difference between pinholin and antipinholin has been attributed to the presence of an extra positively charged amino acid in S2171.30, 34 However, after the initial delay of externalization, TMD1 of S2171 is exported to the periplasm and achieves a similar topology to S2168 which makes it inconvenient for the topological study of antipinholin using conventional spectroscopic techniques. Instead of S2171, Young’s group reported a structural analog of antipinholin by adding five extra amino acids (RYIRS) after the Met of the N terminal of active 21 30, 37 pinholin, referred to as S 68IRS. This ‘RYIRS’ tag prevents the externalization of TMD1 from the lipid bilayer and is reported as a dominant inhibitor of pinholin which makes it a more feasible structural analog of antipinholin S21 for the structural dynamics and topological study using biophysical techniques.30, 35, 37 It is well accepted that proteins’ stability and functions are dictated by its structural topology and dynamic properties.38-42 Hence, it is vital to know the structural details and topological differences between pinholin and antipinholin to understand the structure-activity relationship of this molecular clock mechanism. Pinholin systems were extensively studied by the Young lab using biomolecular and functional techniques.27, 29, 30, 32, 34, 35, 37, 43 However, the structural dynamics, topology, and conformational changes of pinholin have not been extensively investigated using biophysical techniques. There are no detailed structural studies of this system using conventional biophysical techniques such as X-ray crystallography, Cryo-EM, and NMR spectroscopy due to the fact that the study of membrane proteins in the presence of a lipid environment is very challenging using these techniques.44-47 CD experiments have indicated that both forms of pinholin are mostly -helical.48-50 Electron paramagnetic resonance (EPR) spectroscopy is a powerful biophysical technique to study membrane protein structure and dynamics in the presence of lipid environments with high sensitivity and without any size limitations.38, 51-61 Recently, the Lorigan lab reported a tentative model of pinholin S21 using CW- EPR power saturation and solid-state NMR spectroscopic techniques.48, 49, 62 It was reported that TMD1 and TMD2 of inactive antiholin were incorporated into the lipid bilayer.49 However, TMD1 was externalized from the lipid bilayer and remained on the surface of the lipid bilayer for the active pinholin.48 However, higher quality structural data is needed to clearly describe the conformation and topology of the different forms of pinholin. 33, 48, 49, 62 109

To study the membrane topology and structural differenced of pinholin (S2168) and 21 antipinholin (S 68IRS) in a lipid bilayer, four-pulsed double electron-electron resonance (DEER) spectroscopy was used which measures the distance between two spin labels strategically placed in TMD1 and TMD2. DEER spectroscopy coupled with site-directed spin labeling (SDSL) is an powerful biophysical tool for long-range distance (20-80 Å) measurements in biomolecules.51, 63- 68 For this study, full-length pinholin and antipinholin were synthesized using Fmoc solid-phase peptide synthesis (Fmoc-SPPS). Nitroxide spin-label, MTSL (S-(1-oxyl-2,2,5,5-tetramethyl-2,5- dihydro-1H-pyrrol-3-yl) methyl ethanesulfonothioate) was attached by site-directed spin labeling (SDSL) to make an EPR active antipinholin construct.48, 69, 70 DEER measurements were carried 21 21 out on several dual spin labeled active pinholin (S 68) and antipinholin (S 68IRS) proteins in a lipid bilayer. Using these distances restraints, the structural topology of active pinholin and inactive antipinholin were simulated using molecular dynamic simulations (MDs). Data generated in this study and the model generated using MDs showing distinct topological and conformational differences between active and inactive pinholin. The data and the corresponding model clearly indicate that TMD2 of active pinholin remains incorporated in the lipid bilayer, while TMD1 lies on or near the surface of the lipid bilayer perpendicular to TMD2. However, the inactive form of antipinholin adopts a different conformation in which the TMDs have an approximate parallel alignment between TMD1 and TMD2 and both helices remain incorporated into the lipid bilayer. The data agrees well with previous CW-EPR power saturation studies.48, 49

4.3 Experimental Methods 4.3.1 Peptide Synthesis, Purification, and Spin Labeling All peptides were synthesized on an automated CEM Liberty Blue peptide synthesizer equipped with the Discovery Bio microwave system via optimized Fmoc solid-phase peptide synthesis (SPPS) reported in previous studies.48-50 After successful synthesis, the cleavage reaction ran for at least three hours under optimized cocktail conditions to remove the resin and protecting groups form the peptide chain followed by filtration, N2 (g) evaporation, ether precipitation and lyophilization.48-50, 71, 72 The crude peptide was first purified by reverse-phase high-performance liquid chromatography (RP-HPLC) using a GE HPLC system coupled with a C4 (10 µm) preparative column (Vydac 214TP, 250 x 22 mm) as described previously.48 The purified peptide was further purified using C4 semi-preparative column (Vydac 214TP, 250 x 10 mm). The lyophilized pure peptide was dissolved in dimethyl sulfoxide (DMSO) with a 10-fold excess of 110

MTSL (1:5 molar ratio for each spin label site) and stirred for 24 hours in a dark environment to obtain double spin labeled peptide. The spin-labeled (SL) peptide was purified again with a C4 semi-preparative column to remove free MTSL. After each purification, the purity of the target peptide was confirmed by MALDI-TOF MS. Spin labeling efficiency was calculated ~90 to 95% using CW-EPR spectroscopy.

4.3.2 Peptide Incorporation into Proteoliposomes For the DEER measurements, each spin-labeled pinholin or antipinholin peptide was incorporated into DMPC (1,2-Dimyristoyl -sn-Glycero-3-Phosphocholine) proteoliposomes following the thin film method as described previously.48, 50 A 20 mM HEPES (4-(2- hydroxyethyl)-1-piperazineethanesulfonic acid) buffer in D20 (pH ~7.0) was used to rehydrate the thin film. D2O was used instead of H2O to improve the phase time memory of the EPR sample. The final concentrations of lipid and peptide were 50 mM and 50 µM, respectively in the proteoliposomes sample to get a 1000:1 lipid:peptide ratio unless specified. This ratio was chosen to minimize the effect of intermolecular interactions of pinholin which has been shown to oligomerize in the penultimate step of the lysis mechanism.35, 43 A 30% (v/v) glycerol was added to each DEER samples as a cryoprotectant. Homogeneity and size of the proteoliposomes samples were confirmed by using dynamic light scattering (DLS) spectroscopy (ZETASIZER NANO Series; Malvern Instruments) at 25 °C in a disposable 40 μL micro cuvette.48

4.3.3 EPR Spectroscopic Measurements All EPR experiments were conducted at the Ohio Advance EPR Laboratory of Miami University. All DEER samples were initially scanned with CW-EPR spectroscopy using an X- band (~9.34 GHz) Bruker EMX spectrometer equipped with ER041xG microwave bridge and ER4119-HS cavity to confirm the quality of the samples. Experimental setups for CW-EPR spectroscopy were described previously.48 The four-pulsed DEER, experiments were conducted using a Bruker ELEXSYS E580 spectrometer with a SUPERQ-FT pulse Q-band system. The system first used a 10 W amplifier, but then was upgraded to a more powerful 300 W amplifier, with an EN5107D2 resonator. Approximately 70 µL of the sample was loaded into 3 mm quartz EPR tubes and flash frozen with liquid nitrogen just before loading into the resonator cavity. Experimental data was collected with 16-step phase cycling at a temperature of 80 K. An optimized the four-pulsed sequence [(π/2)ν1 – τ1 – (π)ν1 – t - (π)ν2 – (τ1 + τ2 – t) - (π)ν1 - τ2 – echo] was used

111 for dead time free DEER experimental data collection.67, 73 The probe pulse width was 8/16 ns, and pump (π)ν2 pulse width was of 24 ns. 120 MHz of frequency difference was used between the pump and probe pulses. In the upgraded instrumental set up, the pump (π)ν2 pulse was a 70 ns frequency-swept chirp pulse spanning 85 MHz. The shot repetition time was 1000 µs with 100 shots/point. Data acquisition time was 2~3 µs depending on the samples’ phase memory time (T2) and S/N ratio. Data acquisition was done overnight for signal averaging. The DEER data analysis was conducted using the MATLAB DEER Analysis 2015 Program.74 DEER distance distributions, P(r), were obtained using Tikhonov regularization in the distance domain with a minimum distance constraint P(r) > 0 under DEER Analysis 2015.75 The background correction was performed using a two-dimensional homogeneous model for proteoliposomes. The best fit of the time domain data was used for optimizing the regularization parameter in the L-curve.

4.3.4 Structure Refinement of the Active and Inactive Conformations of Pinholin S21 using DEER Distance Restraints The structural refinement of the active and inactive forms of pinholin S21 was carried out using an Xplor-NIH (version 2.46) simulated annealing protocol 76, 77 in a similar manner to that described previously.78-80 A starting structure was obtained by modeling the peptide standard α- helical dihedral angles (Φ =-57.0°, ψ = -47.0°, and ω = 180.0°) for the backbone of the TMDs of 21 81 the active conformations of pinholin S using VEGA ZZ 3.0.5 simulation toolkit. Positions 16, 17, 20, 24, 27, 38, 40, 44 and 46 along the active form of the pinholin S21 sequence were mutated to Cys and MTSL side chains were attached with the Xplor-NIH addAtoms.py script using NIH- Xplor-2.46.76, 77 Experimental DEER data for five inter-label distances (16/46, 17/38, 20/44, 24/40, 27/38) on the active version of pinholin were used to define restraints for an Xplor-NIH simulated annealing protocol for active pinholin. A starting structure was obtained by using the amino acid sequence of the inactive conformation of pinholin S21 on the template structure of the helix 3 and helix 4 (S3-S4) segment of the KCNQ1-VSD (PDB ID: 5VMS) under Xplor-NIH using Xplor- NIH addAtoms.py script. This template was used because the structural conformation was similar to that of the inactive antiholin based on earlier study by Ry Young group.30, 37 Positions 8, 14, 15, 24, 27, 38, 40, and 53 along the inactive form of pinholin S21 sequence were mutated to Cys and MTSL side chains were attached with the Xplor-NIH addAtoms.py script. Experimental DEER data for five inter-label distances (8/53, 14/53, 15/53, 24/40, 27/38) on the inactive version of pinholin were used to define restraints for an Xplor-NIH simulated annealing protocol for inactive 112 pinholin. The allowable ranges used for inter-label distances were established through a series of preliminary simulated annealing molecular dynamics calculations in which these ranges were varied. 80 One hundred structures were generated using the Xplor-NIH simulated annealing routine with the Xplor-NIH_foldB.py script. The ten lowest energy structures were further refined using simulated annealing protocol using Xplor-NIH refine_EEF.py using the above-mentioned DEER distance restraints using eefxPotTools included in the refine_EEF.py script. The simulated annealing procedure used 3500K as the high temperature and 25 K as the cooling temperature with temperature steps of 12.5 K. The 10 lowest energy structures were kept with energies < -1575.71 Kcal/mol for active conformation of pinholin. Similarly, the 10 lowest energy structures were kept with energies < 89.54 Kcal/mol for the inactive conformation of pinholin. These lowest energy structures were further validated against DEER distance restraints to make sure the spin labels distances are within experimental errors. Further analysis and visualization was done using VMD- Xplor software.82 The final structures were generated by replacing MTSL spin labeled side chains with native side chains using Xplor-NIH addAtoms.py script under the Xplor-NIH similar to a previously reported method.80 The further details of the simulated annealing outputs are given in Supporting Information (Table S1 and S2). All MD simulations were run at the Miami Redhawk cluster computing facilities at Miami University.

4.4 Results Recently, we reported the structural dynamics and topology of active pinholin and inactive antipinholin incorporated into DMPC proteoliposomes using solid state nuclear magnetic resonance (SSNMR) spectroscopy, CW-EPR line-shape analysis, and power saturation spectroscopy.48, 50, 62 In this study, DEER spectroscopy is used to investigate the structural topology of active pinholin and inactive antipinholin, and to demonstrate the topological differences between these two forms of pinholin S21. Figures 4.1(A) and (B) are the primary amino acid 21 21 sequence for the active pinholin (S 68) and the inactive analog of antipinholin (S 68IRS) as a reference. Figures 4.1(C) and 4.1(D) are the tentative topology of active and inactive conformation of pinholin S21, respectively. To apply DEER spectroscopy to study the structural parameters of pinholin S21, we replaced the specific native amino acids with cysteine to introduce nitroxide spin labels at site directed positions. In total, we studied five dual spin labeled full-length active 21 21 pinholin (S 68) and five dual spin-labeled inactive antipinholin (S 68IRS). Spin label positions

113 were judiciously selected to abate structural perturbation based on published mutational analysis and our initial studies.35, 37, 43, 49

Figure 4.1: Primary sequence and tentative topology of active pinholin and inactive antipinholin. TMD1 and TMD2 are indicated by red and green boxes, respectively. (A) The primary sequence 21 21 of pinholin S 68, (B) Primary sequence of antipinholin S 68IRS. ‘RYIRS’ tag are incorporated between Methionine and Aspartate in the N-terminal. (C) Two possible orientations of active pinholin where TMD1 is completely externalized from the lipid bilayer or stay on the surface of 21 the lipid-solvent interface. (D) The tentative topology of inactive antipinholin S 68IRS.

4.4.1 DEER Distance measurement for the active conformation of pinholin S2168 The structural and functional biochemical literature on pinholin S2168 suggest that TMD1 of pinholin S2168 externalizes from the lipid bilayer, while TMD2 remains incorporated in the lipid bilayer to achieve the active conformation of pinholin S2168.30, 32, 37, 43, 48, 62 However, there is no concrete experimental evidence to show that TMD1 remains in the periplasm after externalization and the relative orientation of TMD1 and TMD2. Based on Cys cross-linking and accessibility of

114 disulfide bonds, Ry Young’s lab proposed the complete externalization of TMD1 from the lipid bilayer. However, recent biophysical studies have indicated a partial externalization of TMD1 which lay on the surface of the lipid bilayer to adapt a topology where TMD1 is almost perpendicular to TMD2, based on CW-EPR line-shape analysis, power saturation, and solid state NMR experiments.48, 62 Both proposed conformations are shown in Figure 4.1(C). To investigate the relative orientation and distance between TMD1 and TMD2, DEER spectroscopy was used in which one spin label was placed on TMD1 and another spin label on TMD2. The positions of the spin labels were judiciously selected based on the published literature data and our initial experimental results.37, 48 Spin pairs were designed to show the relative distance between TMD1 and TMD2 and positions of the spin labels as a pair are shown in Figure 4.2(A). The extreme ends of N- and C-termini were avoided considering the fact that the distance distribution measured above 50 Å is not quantitative for membrane protein in the presence of lipid bilayer due to the short data acquisition time for DEER trace.83 Again, distances longer than 50 Å are usually suppressed by background correction and can generate artifact peaks at long distances for a data-trace length of 2 µs.83 Corresponding DEER spectra of active pinholin in DMPC proteoliposomes are shown in Figures 4.2 (B)-(F).

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Figure 4.2: Q-band DEER data of active pinholin S2168 in DMPC Proteoliposomes. (A) Spin label positions in structural model, (B-E) are the corresponding DEER distances with time domain data.

The baseline-corrected time-domain traces (left) are shown with the corresponding distance probability distributions from Tikhonov regularization (right). Table 4.1 presents the final distance summary obtained for pinholin S2168 with the error limit of ±3Å.

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Table 4.1: Summary of the major peak DEER distance for the active pinholin.

Spin labels positions on Pinholin S2168 Distance (±3Å) W27R1_A38R1 24 F24R1_G40R1 25 A17R1_A38R1 35 A20R1_S44R1 44 S16R1_V46R1 49

DEER distance clearly demonstrates that distance between two spin labels increases gradually when spin pairs move from the loop region to the termini regions which indicates that the N-terminus of TMD1 is going apart from the C-terminus of TMD2. The shortest distance observed was 24Å for W27_A38 and the longest distance was 49Å for S16_V46. Table 4.1 presents the final distance summary obtained for pinholin S2168. These distance restraints were used for the MDS of the structural topology of active pinholin S2168.

4.4.2 DEER Distance measurement for the inactive conformation of antiholin 21 To probe the relative orientation and distance between TMD1 and TMD2 of S 68IRS, one spin label was placed on TMD1 and another on TMD2 of antiholin. Figure 4.3 shows the spin label positions in the structural model of antipinholin. Spin label pairs were designed to minimize structural perturbations based upon previous studies in the literature.37, 48 This was critical for TMD1 which has a natural tendency to externalize and certain single mutations can (e.g. G21Q) cause activation of antipinholin (implies externalization of TMD1) as reported by Pang et.al .37 The DEER data and corresponding distance probability distributions for each pair of spin labels 21 placed on antipinholin S 68IRS are shown in Figure 4.3.

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21 Figure 4.3: Q-band DEER data for inactive pinholin S 68IRS in DMPC proteoliposomes. (A) Spin label positions are shown in the structural model, (B-E) are the corresponding DEER time domain data and distances probability distributions.

Table 4.2 presents the final distance summary obtained for the inactive antipinholin 21 S 68IRS. These distance restraints were used for the MDs of the structural topology of the inactive 21 antipinholin S 68IRS.

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21 Table 4.2: Summary of the major peak DEER distance for the inactive pinholin S 68IRS.

21 Spin labels positions on antipinholin S 68IRS Distance (±3Å) W27R1_A38R1 23 F24R1_G40R1 25 T15R1_L53R1 26 G14R1_L53R1 25 S8R1_L53R1 26

The shortest distance was 23 Å for W27R1_A38R1 (close to the loop region) which remain similar for the rest of the parallel spin pair distances. The distance between terminal regions (S8R1_L53R1) was found 26Å which is unlikely if TMD1 is externalized and moves far from TMD2 like active pinholin. Other distances were 25, 26, 25 Å for F24R1_G40R1, T15R1_L53R1, and G14R1_L53R1, respectively.

4.4.3 Structure Refinement of the active pin holin and inactive antipinholin from MTSL DEER Distance Restraints A model for the most probable structure of the active pinholin and inactive antipinholin in DMPC proteoliposomes was obtained using the experimental DEER distances coupled with the simulated annealing software package Xplor-NIH (version 2.33).76, 77 The DEER distance data obtained for five pairs of MTSL spin labeled sites (see Materials and Methods) in proteoliposomes independently for both active pinholin (Table 4.1) and inactive antipinholin (Table 4.2) were converted into Xplor distance restraints and utilized in a simulated annealing protocol. The MTSL DEER distance restraints were used to make the structure calculation procedure simpler within Xplor-NIH. Since MTSL is a widely used spin probe for EPR spectroscopic studies, the method of structure refinement used in this study can be widely applied to many other membrane proteins. A series of simulated annealing calculations were performed using distance-restraint uncertainties of ± 3 Å independently for both active pinholin and inactive antipinholin. Figure 4.4 displays the 10 lowest energy structures from a family of 100 calculated structures of the active pinholin and inactive antipinholin in proteoliposomes obtained from the simulated annealing calculation using MTSL-derived distance restraints. These structures satisfied Experimental DEER distances within 119 experimental errors. To generate the final structures, the MTSL spin-labeled side chains were replaced by the native side chains. The output energies corresponding to ten minimum energy structures obtained from Xplor-NIH refinement are given in the Supporting Information (Table 4.3 and 4.4).

Figure 4.4: Results of the structural refinement of the active pinholin and inactive antipinholin incorporating MTSL DEER distance-restraint data using an Xplor-NIH simulated annealing molecular dynamics protocol. Overlay of the ribbon representation of the 10 DEER structures with lowest energy obtained from restrained simulated annealing calculations using the amino acids of active pinholin (A) and inactive pinholin (B). The final structures were generated by replacing the MTSL labeled side chains by the native amino acid side chains with retention of the 82 Cβ position in the label. Image was created using VMD-Xplor program.

4.5 Discussion Q-band DEER data and the corresponding distance distributions coupled MDs clearly demonstrate two different structural conformations and topology for active pinholin vs inactive antipinholin. Distances between TMD1 and TMD2 for inactive antipinholin clearly indicated that both helices remain in close proximity with a parallel conformation maintaining an average distance of 23 to 26 Å. Our previous studies using CW-EPR, power saturation, and solid-state NMR indicated that both helices remain in the lipid bilayer for inactive antipinholin which was consistent with the model proposed by the Young group.30, 62 The structural model obtained using DEER distance restrains under simulated annealing molecular dynamics simulation (Figure 4.4) is consistent with the results found in this study as well as the previous model.

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For the active form of pinholin, DEER distances between TMD1 and TMD2 clearly indicate that the N-terminal of TMD1 is moving away from the C-terminal of TMD2 and support the notion that TMD1 externalizes from the lipid bilayer.30, 48, 62 Considering our previous CW- EPR power saturation studies and proposed model by Young’s lab, it was confirmed that TMD2 remains inside the lipid bilayer, which implies that TMD1 moves away from TMD2 and is externalized from the lipid bilayer.30, 33, 48, 62 However, it was not certain what the orientation of TMD1 is relative to TMD2 after externalization and two probable orientations were proposed in the literature as shown in Figure 4.1(C). The distance between S16R and V46R1 was found to be 49 Å, which would be a much longer distance if TMD1 completely externalized from the lipid bilayer as shown in Figure 4.1C. This clearly supports the orientation where TMD1 remains about perpendicular with TMD2 instead of completely externalized. Although, we are not completely excluding the probability of complete externalization considering the fact that a dynamic protein like pinholin can adopt multiple conformations, where the majority of the population remains in the perpendicular orientation on the surface and some population remains in completely externalized orientations. This may be one of the possible reasons for the broadening of the DEER distribution for most of the DEER data obtained in this study. Longer DEER distance above 50 Å can get suppressed for membrane proteins incorporated into a lipid bilayer due to the short data acquisition time and background subtraction which becomes undetectable or nontrivial for very small population.83 At the same time, shorter distance populations around 25 Å probable arose from a small population of active pinholin in which TMD1 did not externalize from the lipid bilayer and oriented parallel with TMD2 and/or intermolecular distances between adjacent TMD2- TMD2 due to the natural propensity of oligomerization for active pinholin S2168 in the presence of lipid bilayer. Oligomerization was minimized in the study by conducting the experiments at a low (1000:1) lipid:peptide ratio. The DEER structural models obtained for both active pinholin and inactive antiholin are consistent with our earlier biophysical studies and biological experiments conducted by the Ry Young group.30, 37, 48, 62

4.6 Conclusion This study reported structural models and conformations of both the active pinholin and antipinholin of the phage 21 lytic proteins, in a lipid bilayer using DEER spectroscopy coupled with the simulated annealing molecular dynamics (MD) simulation. DEER distance distribution

121 and derived structural model using simulated annealing MD simulation demonstrated clear conformational differences between two forms of pinholin protein. Furthermore, this study reported refined structural models of these two proteins and resolved existing conflict structural models of active pinholin in the field. The structural model of S21 presented in this study will be useful for understanding of structural and functional relationship of pinholin S21 as well as other holin systems using biophysical techniques. This study will help researchers to apply this powerful ESP spectroscopic approaches to investigate conformational changes in more complicated membrane protein system and hence will move the structural biology field forward.

4.7 Supporting Information

Structures Energy (Kcal/mol) 1 -1955.02 2 -1949.99 3 -1939.41 4 -1930.00 5 -1923.97 6 -1920.49 7 -1875.80 8 -1824.46 9 -234.00 10 -203.96 Average -1575.71

Table 4.3: Minimum energy associated with the refined structures of active pinholin obtained from the simulated annealing molecular dynamics simulation.

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Structures Energy (Kcal/mol) 1 52.43 2 75.15 3 78.47 4 80.87 5 92.62 6 94.47 7 96.73 8 107.60 9 107.71 10 109.31 Average 89.54

Table 4.4: Minimum energy associated with the refined structures of inactive antipinholin obtained from the simulated annealing molecular dynamics simulation.

Acknowledgment We are grateful to the members of the Ry Young group at Texas A&M University for their experimental suggestions. We would also like to appreciate Dr. Jens Mueller, the Facility Manager Redhawk Cluster Miami University, for assistance with the computational work. This work was generously supported by the NIGMS/NIH Maximizing Investigator’s Research Award (MIRA) R35 GM126935, the NSF CHE-1807131 grant, the NSF (MRI-1725502) grant, the Ohio Board of Regents, and Miami University. Gary A. Lorigan would also like to acknowledge support from the John W. Steube Professorship.

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Chapter 5

Mutations of Pinholin S21 Induce Structural Topology and Conformational Changes are Observed with Electron Paramagnetic Resonance Spectroscopy

Tanbir Ahammad†, Rasal H. Khan†, Indra D. Sahu‡†, Daniel L. Drew Jr. †, Emily Faul†, Tianyan Li†, Robert M. McCarrick†, Gary A. Lorigan†*

†Department of Chemistry and Biochemistry, Miami University, Oxford, OH, 45056, USA.

‡Natural Science Division, Campbellsville University, Campbellsville, KY, 42718, USA.

“Preparing the manuscript for publication”

I carried out most of the experiments, data analysis, and wrote the manuscript. Khan and Drew carried out some experiments. Dr. Sahu helped in data analysis. Faul, and Li helped with the sample preparation.

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5.1 Abstract The bacteriophage infection cycle is terminated at a predefined time to release the progeny virions via a robust lytic system composed of holin, endolysin, and spanin proteins. Holin is the timekeeper of this process. Pinholin S21 is a prototype holin of phage 21, which determines the timing of host cell lysis through the coordinated efforts of pinholin and antipinholin. However, mutations in pinholin and antiholin play a significant role to slow or accelerate the timing of lysis depending on adverse or favorable growth conditions. Earlier studies have shown that single point mutations of pinholin S21 alter the cell lysis timing (Pang et al 2010). However, mutational analysis of pinholin using cell lysis timing is an indirect measurement of pinholin function as lysis is dependent on other lytic proteins as well. In this study, continuous wave electron paramagnetic resonance (CW-EPR) power saturation and double electron-electron resonance (DEER) spectroscopic techniques were used to directly probe the effects of mutations on the structure and conformational changes of pinholin S21 that correlate with pinholin function. The collected DEER and CW-EPR power saturation data clearly demonstrates that residue mutations that increase hydrophilicity accelerate the externalization of antipinholin transmembrane domain 1 (TMD1), while increased hydrophobicity prevents the externalization of TMD1 thus possibly accelerating or delaying the activation of pinholin S21, respectively. It was also found that mutations can influence intra- or intermolecular interactions in this system which contribute to the activation and oligomerization to form a pinhole and modulate the cell lysis timing. This could be a novel approach to analyze the mutational effects on other holin systems as well as any other membrane protein in which mutation directly leads to structural and conformational changes.

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5.2 Introduction Bacteriophages are the most abundant organism in the biosphere and they recycle much of the world’s biomass thorough ~1028 infection cycles per day.1, 2 Phages evolved a robust lytic system to control the length of the infection cycle so that it can be adjusted for different environments and host populations.3, 4 Most large, dsDNA bacteriophages use at least three different proteins (holin, endolysin, and spanin) to lyse gram-negative bacteria by permeabilizing and degrading the cytoplasmic membrane, peptidoglycan layer, and cell wall, respectively.5-9 The holin protein functions as an allele-specific molecular timer that triggers the formation of microscale holes for the release of fully-folded functional endolysin4, 5, 10-21 Although the holin- endolysin system is a robust and efficient lytic system, ~25% of phages employ a prototype evolutionary intermediate lytic system which is known as the pinholin-SAR (signal anchor release) endolysin system.22 Pinholins make nanoscale holes which are not large enough to accommodate folded endolysin but allow the passage of protons to dissipate the proton motive force (PMF) which in turn leads to the release and activation of the membrane-tethered SAR-endolysin for its muralytic action.10, 22 Pinholin S21 from lambdoid phage 21 is one of the most well-studied pinholin systems.10, 23-28 This system has two holin proteins with an active pinholin (S2168) and an antipinholin (S2171). Both pinholin proteins have two TMDs and they accumulate benignly in the cytoplasmic membrane as an inactive dimer where both TMDs reside in the bilayer. For pinhole formation to occur, TMD1 of S21 must be externalized from the lipid bilayer. For S2168, TMD1 externalization is a rapid and spontaneous process. The externalization of S2171 TMD1 is much slower relative to S2168 due to the presence of an additional positively charged Lys residue in the N-terminal region. The difference in the rate of externalization between these two proteins leads to delayed pinhole formation.10, 24 S2171 is a weak antipinholin as it can only delay, but can’t completely block the externalization of TMD1.24 The Ry Young lab has reported the generation of a strong antipinholin by inserting five amino acids (RYIRS) into the N-terminal region of S2168. The externalization of TMD1 is blocked because of the addition of more positive charges and bulky side chains, and this 21 24 21 21 antipinholin variant is denoted by S 68IRS. In this study, S 68IRS was used instead of S 71 to prove the mutation effect on inactive antipinholin. Phages use a combination of holin and antiholin to adjust the timing of lysis.3, 22 In addition, phages tune their lytic function by the mutation of endolysin and/or holin to adjust with 134 the changing host and environment.3, 22 In the case of phage 21, the mutation of SAR-endolysin alone is not a viable way for the phage to change the lysis timing, since there are very few mutational options in the membrane-tethered N-terminal region without changing the effective

22 Kcat. Hence, mutation of pinholin instead could be the mechanism by which phage 21 adjusts lysis timing for changing environments. Pang et al. (2010) reported an extensive mutational study of S21 with a wide range of phenotypes, including absolute lysis defective variants, as well as those 24 21 with delayed or accelerated lysis triggering. Although S 68IRS has been shown to be the dominant antipinholin that prevents cell lysis, some mutations rendered it active and abolished its antiholin properties.24 In that study, more emphasis was given to S2168 and fewer mutants of 21 S 68IRS were examined. To prove the mutational effects, the cell lysis time was monitored which is not enough to define pinholin activity as SAR endolysin alone can cause lysis after induction, regardless of expression of the pinholin allele.22, 24 This study reports the effects of various mutations on the structural and topological 21 properties of antipinholin S 68IRS using CW-EPR power saturation and DEER spectroscopic 21 techniques. This work directly evaluated the structural and conformational changes of S 68IRS and correlated those observed with activation or inactivation of the protein from a previous biological study.24 This mutational study clearly demonstrates that the relative hydrophobicity of TMD1 impacts its externalization, which further controls the activation of inactive dimers, and finally the formation of pinholes. In addition, mutations also changed how TMD1 and TMD2 interact which ultimately impacted the externalization of TMD1, leading to alteration of triggering time. These results provide a much more comprehensive picture of the mutational effects on the structural 21 topology and conformational changes of S 68IRS and explain how these conformational changes influence the functionality of pinholin system.

5.3 Experimental Methods 5.3.1 Peptide Synthesis, Spin Labeling, and Purification All peptides were synthesized on an automated CEM Liberty Blue peptide synthesizer equipped with the Discovery Bio microwave system via optimized Fmoc solid-phase peptide synthesis (SPPS) reported in previous studies.25, 26 In brief, each synthesis was started with 0.1 mM glutamate preloaded TGA resin. The synthesis was done in the dimethylformamide (DMF) based solvent system. Piperidine (20% v/v), N, N′-diisopropylcarbodiimide (DIC) (15.6% v/v),

135 and oxyma (14.2% w/v) in DMF were used as a deprotecting agent, activator, and activator base, respectively. During each coupling cycle, 0.2 M amino acid was added to the reaction vessel in the presence of activator and activator base. After successful synthesis, the crude peptide was obtained following a previously published optimized cleavage procedure.25, 26, 29-31 The crude peptide was purified by reverse-phase high-performance liquid chromatography (RP-HPLC) using a GE HPLC system coupled with a C4 (10 µm) preparative column (Vydac 214TP, 250 x 22 mm) following the optimized method.25, 26 To attach the spin label (SL) to the peptide, the lyophilized pure peptide was dissolved in dimethyl sulfoxide (DMSO) with a 5-fold excess of MTSL (S-(1-oxyl-2,2,5,5-tetramethyl-2,5- dihydro-1H-pyrrol-3-yl) methyl ethanesulfonothioate) per Cys residue (1:5 molar ratio for the single spin label and 1:10 molar ratio for the dual spin label) and stirred for 24 hours in the dark. The spin labeled peptide was further purified using a C4 semi-preparative column (Vydac 214TP, 250 x 10 mm) to remove unbound MTSL and other contaminants.25, 26 After each purification, the purity and identity of the target peptide was confirmed by MALDI-TOF MS. Spin labeling efficiency was ~85-90% as calculated by CW-EPR measurements.26

5.3.2 Peptide Incorporation into Proteoliposomes To mimic the membrane environment, spin labeled antipinholin peptides were incorporated into DMPC (1,2-Dimyristoyl-sn-Glycero-3-Phosphocholine) proteoliposomes using the thin film method.25, 26 To prepare the samples for CW-EPR and power saturation experiments, a 10 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) buffer (pH ~7.0) was used to rehydrate the thin film. The peptide:lipid ratio was 1:1000 in the final proteoliposome sample. Glycerol was added to a 10% final concentration to help the sample remain suspended for a longer duration at room temperature without phase separation.25 For DEER samples, 20 mM HEPES (4-

(2-hydroxyethyl)-1-piperazineethanesulfonic acid) buffer in D2O (pH ~7.0) was used to rehydrate the thin film. The final concentrations of lipid and peptide in the proteoliposome samples were 50 mM and 50 µM, respectively, to get a 1:1000 ratio of peptide:lipid unless otherwise specified. This ratio was chosen to minimize the effect of intermolecular interactions between monomers of pinholin which have been shown to oligomerize in the penultimate step of the lysis mechanism.23, 28 Glycerol was added to each DEER samples at a 30% (v/v) final concentration as a cryoprotectant. Before rehydration of any proteoliposome sample, both HEPES buffer and the sample flask were kept in a warm water bath for a short period of time to bring the temperature 136 above the phase transition temperature of DMPC.25, 32, 33 Homogeneity and size of the proteoliposomes in each sample were confirmed with dynamic light scattering (DLS) spectroscopy (ZETASIZER NANO Series; Malvern Instruments) at 25°C in a disposable 40 μL micro cuvette.25

5.3.3 CW-EPR Spectroscopy All EPR experiments were conducted at the Ohio Advanced EPR Laboratory at Miami University. CW-EPR spectra were collected at X-band (~9.34 GHz) with a Bruker EMX spectrometer equipped with ER041xG microwave bridge and ER4119-HS cavity. Each spectrum was acquired by signal averaging 10 scans with 3315 G central field, 150 G sweep width, 42 s field sweep, 100 kHz modulation frequency, 1 G modulation amplitude, and 10 mW microwave power.25

5.3.4 CW-EPR Power Saturation Experiments CW-EPR power saturation experiments were performed on a Bruker EMX X-band spectrometer coupled with ER 041XG microwave bridge and ER 4123D CW-Resonator (Bruker BioSpin). Experimental setups were optimized following previously published literature.34-36 3-4 μL samples were loaded into a gas permeable TPX capillary tube at a concentration of 100-150 μM.34, 37-39 EPR spectroscopic data was collected using a modulation amplitude of 1.0 G, a modulation frequency of 100 kHz, 42 sec field sweep, and 90 G sweep width.25 Incident microwave power was varied from 0.05 mW to 126 mW. For each spin labeled site, the spectra were recorded under three equilibrium conditions; oxygen, nitrogen, and NiEDDA equilibriums as described in previous literature.25, 39 CW-EPR power saturation data were extracted and analyzed using a MATLAB software script. The peak-to-peak amplitudes of the first derivative central resonance lines (A) were extracted and plotted against the square root of the incident microwave power (P). These data points were then fitted according to Equation (1):38, 40

1 – ε (2ε− 1)P A = I√P [1 + ] (1) P1 2

where I is a scaling factor, ε is the homogeneity of saturation of the resonance line, and P1⁄2 is the power where the first derivative amplitude is reduced to half of its unsaturated value. ε values varied between 1.5 to 0.5 for the homogeneous to inhomogeneous saturation, respectively.40 In

137 equation (1), I, ε, and P1⁄2 are adjustable parameters and yield a characteristic P1⁄2 value for each equilibrium condition. The corresponding depth parameter () was calculated using equation (2):40

ΔP1(푂2)  = ln [ 2 ] (2) ΔP1(NiEDDA) 2 where ∆P1/2(NiEDDA) is the difference in the P1/2 values for NiEDDA and nitrogen equilibriums, and∆P1/2(O2) is the difference in the P1/2 values for oxygen and nitrogen equilibriums.

5.3.5 DEER Spectroscopic Measurements The four-pulse DEER experiments were conducted using a Bruker ELEXSYS E580 spectrometer with a SUPERQ-FT pulse Q-band system. For earlier data collection, the system used a 10 W amplifier, but recently it was upgraded to a more powerful 300 W amplifier with an EN5107D2 resonator. Approximately 70 µL of the sample was loaded into a 3 mm quartz EPR tube and flash-frozen with liquid nitrogen prior to insertion in the resonator cavity. Experimental data was collected with 16-step phase cycling at a temperature of 80 K. An optimized four-pulsed sequence [(π/2)ν1 – τ1 – (π)ν1 – t - (π)ν2 – (τ1 + τ2 – t) - (π)ν1 - τ2 – echo] was used for dead time free 41, 42 DEER experimental data collection. The probe pulse width was 8/16 ns, and pump (π)ν2 pulse width was of 24 ns. 120 MHz of frequency difference was used between the pump and probe pulses. In the upgraded instrumental set up, the pump (π)ν2 pulse was a 70 ns frequency-swept chirp pulse spanning 85 MHz. The shot repetition time was 1000 µs with 100 shots/point. Data acquisition time was 2~3 µs depending on the samples’ phase time memory (T2) and S/N ratio. Data acquisition was done overnight for signal averaging. The DEER data analysis was conducted using the MATLAB DEER Analysis 2015 Program.43 DEER distance distributions, P(r), were obtained using Tikhonov regularization in the distance domain with a minimum distance constraint P(r) > 0 under DEER Analysis 2015.44 The background correction was performed using a two- dimensional homogeneous model for proteoliposomes. The best fit of the time domain data was used for optimizing the regularization parameter in the L-curve.

5.4 Result and Discussion Recently, we reported the structural dynamics and topology of active pinholin (S2168) and 21 inactive antipinholin (S 68IRS) incorporated into DMPC proteoliposomes using CW-EPR line-

138 shape analysis, power saturation, and DEER spectroscopic experiments (unpublished Chapter 25, 45 21 21 4). Figure 5.1 shows the primary sequence of S 68 and S 68IRS with their structural topology model adapted from the literature.10, 25, 45

Figure 5.1: Primary sequence and structural topology model of active and inactive pinholin. TMD1 (green) and TMD2 (yellow) are depicted as cylinders. (A) and (B) show the primary amino 21 21 acid sequences of S 68 and S 68IRS where an ‘RYIRS’ tag has been incorporated between Met4 21 21 and Asp5 in the N-terminus of S 68IRS. (C) and (D) show the possible topology models of S 68 21 and S 68IRS, respectively.

For EPR spectroscopic experiments, a nitroxide spin label was incorporated onto the pinholin peptide using directed spin labeling (SDSL).46, 47 In our previous structural studies of S21, the spin label positions were judiciously selected to preserve the native conformations of active and inactive pinholin and consciously omitted those sites which might induce structural or functional perturbations according to the literaturea.24, 25, 45 The structural perturbation induced by the spin labels were more prominent for the inactive antipinholin since its TMD1 has a natural tendency to be externalized from the lipid bilayer.10, 45 It was hypothesized that those specific

139 residue positions have a significant mutational effect. Taking account of these observations and the previous mutational study reported by Pang et al (2010), certain residue positions (e.g. G14, 21 24 S16, A17) were selected to study the mutational effects on the structure and topology of S 68IRS.

In this study, CW-EPR power saturation experiments were conducted to investigate the 21 mutational effect on the structural conformation of S 68IRS TMD1. A nitroxide spin label was 21 placed at A20 of S 68IRS and denoted by IRS_A20R1 (R1 represents the MTSL spin label attached to Cys residue through a disulfide bond). This spin label position was selected as a control and anchoring point for the rest of the experiments, since A20 is positioned approximately at the center of TMD1 and a majority of the population of A20R1 was found buried inside of the lipid bilayer (positive  value) for the inactive conformation and outside of the lipid bilayer (negative  value) for the active conformation as reported previously (Figure 5.1 C and D).25, 45 The  values represent the relative accessibility of oxygen and NiEDDA for the nitroxide spin label and implies the relative population of the TMD1 inside vs outside of the lipid bilayer. Hence, the  values obtained for IRS_A20R1 in this study will report on the relative population of TMD1 inside vs outside of the lipid bilayer which could be correlated with the inactive and active conformation of pinholin S21.

Previous studies have suggested that the changes in hydrophobicity play a crucial role in the activation of pinholin by changing the probability of externalization of TMD1.24 To examine 21 21 this effect on the conformation of S 68IRS, two more S 68IRS constructs were designed keeping the spin label at the A20 position, with a single mutation (A17Q) , and a double mutation (G14QA17Q). These constructs were denoted as IRS_A20R1_A17Q, and IRS_A20R1_G14QA17Q, respectively (Figure 5.2).

140

21 Figure 5.2: Changes in hydrophobicity influenced the externalization of TMD1 of S 68IRS. (A) Tentative conformational change and equilibrium between to conformation of pinholin S21. The position of the spin label is in green and the points of mutation are the red letters in orange balls. CW-EPR power saturation data for IRS_A20R1 (B), IRS_A20R1_A17Q (C), IRS_A20R1_G14QA17Q (D), and IRS_A20R1_A17L (E).

141

The  value was calculated as 1.0 for IRS_A20R1 without an additional mutation which 21 implied that a majority of the S 68IRS TMD1 population was located inside of the lipid bilayer. However, the  values of -0.1 for IRS_A20R1_A17Q and -0.6 for IRS_A20R1_G14QA17Q were measured. A more negative  value indicated that a higher population of TMD1 was externalized as the hydrophilicity of the TMD1 region was increased. To examine the opposite effect, A17 was substituted with a more hydrophobic leucine (L) side chain, while keeping the spin label at the 21 same A20R1 position of S 68IRS. The construct was denoted as IRS_A20R1_A17L (Figure 5.2E).

21 The  value for IRS_A20R1_A17L was 1.0, which implied that a dominant population of S 68IRS 21 TMD1 were inside of the lipid bilayer like the inactive conformation of S 68IRS. This observation further confirmed that the increased hydrophobicity of TMD1 prevented the externalization of 21 S 68IRS TMD1, while increased hydrophilicity is more likely to induce the dissociation of TMD1 from the membrane. The cumulative change of  values due to changes in hydrophilicity, as observed in IRS_A20R1_A17Q and IRS_A20R1_G14QA17Q, indicated a change in relative population of TMD1, inside vs outside of lipid bilayer. This implies a dynamic equilibrium between active and inactive conformations of pinholin S21. The increased hydrophobicity of TMD1 21 shifted the equilibrium towards the inactive conformation and a greater population of S 68IRS TMD1 stayed inside of the lipid bilayer, while increased hydrophilicity shifted equilibrium 21 towards the active and more population of S 68IRS TMD1 was stayed outside of the lipid bilayer. To further explore the effect of varying types of sidechain and charge with its hydrophobic effect, S16 was replaced with the relatively hydrophilic amino acid, glutamate (E) or relatively hydrophobic amino acid, phenylalanine (F). For direct comparison, the spin label was placed at the same A20 position. To explore the effect of these mutations on the externalization of TMD1, two constructs (IRS_A20R1_S16E and IRS_A20R1_S16F) were probed using the CW-EPR power saturation experiments where the depth parameters were compared with IRS A20R1 (Figure 5.3).

142

21 Figure 5.3: Mutational effect on the externalization of S 68IRS TMD1. CW-EPR power saturation data collected for (A) IRS_A20R1 without additional mutation, (B) IRS_A20R1_S16E, and (C) IRS_A20R1_S16F.

The  values of -0.4 for IRS_A20R1_S16E clearly demonstrated that a greater population of A20R1 spin-labeled TMD1 stayed outside of the lipid bilayer when compared to that for IRS_A20R1 (negative vs positive  value). These data implied that this mutation (S16E) enhanced 21 the externalization of S 68IRS TMD1 and hence most of the population adapted an active conformation of pinholin S21. However, for IRS_A20R1_S16F,  value (1.1) was similar to IRS_A20R1 indicating that the most of the population of A20R1 spin-labeled TMD1 was buried inside of the lipid bilayer similar to that of IRS_A20R1 and implied that this mutation does not 21 enhance the externalization of S 68IRS TMD1 and the majority of the population adapted an 21 inactive conformation of pinholin S 68IRS. These observations are consistent with earlier CW- EPR power saturation data (Figure 5.2) suggesting that the increased hydrophobicity of the IRS_A20R1_S16F variant prevented the externalization of TMD1 while increased hydrophilicity in IRS_A20R1_S16E enhanced the externalization of TMD1. However, sidechain type (aromatic vs aliphatic) and charge (positively vs negatively charged) did not play a significant role on the externalization of TMD1 as observed for relative hydrophobicity of TMD1. In addition to the mutational effect induced by amino acids, the spin label can lead to structural perturbation in certain positions and is comparable to the mutations that caused the

143 externalization of TMD1 of inactive pinholin. For example, for positions IRS_S16R1 and IRS_A17R1, it was expected that a majority of the population will be located inside of the lipid bilayer as suggested by the previous study that showed G14, S19, and A20 were located inside of the lipid bilayer for antipinholin.45 However, the calculated depth parameters () were -0.8 and - 1.0 for IRS_S16R1 and IRS_A17R1, respectively, which clearly demonstrated that a majority of the populations were externalized from the lipid bilayer and stayed as solvent exposed (Figure 5.4).

21 Figure 5.4: Effect of MTSL on the externalization of S 68IRS TMD1. CW-EPR power saturation data collected for variants (A) IRS S16R1 and (B) IRS A17R1.

These observations are consistent with the mutational results observed in the previous section (Figure 5.2 and 5.3) where changes in hydrophobicity of these positions significantly affected the TMD externalization process. However, externalization of TMD1, induced by nitroxide spin labels are more likely linked to the change in the intramolecular or intermolecular interactions with TMD2 rather than changes in TMD1 hydrophobicity. To visualize the relative effect of each individual mutation on the relative population of 21 S 68IRS TMD1, calculated depth parameters are summarized in Figure 5.5.

144

21 Figure 5.5: Comparison of the depth parameter for different S 68IRS mutants with respect to the IRS_A20R1. The red columns indicate a negative depth parameter where majority of the spin label was outside of the lipid bilayer. The green columns indicate a positive depth parameter where majority of the spin label was inside of the lipid bilayer.

Figure 5.5 clearly indicate that the increased hydrophilicity at A17, G14, and A16 21 positions increased the propensity of S 68IRS TMD1 to externalize. Hence, more and more populations adapt the active conformation. However, the increased hydrophobicity at A17 and 21 A16 positions prevented S 68IRS TMD1 externalization (Figure 5.5). Additionally, the placement 21 of the R1 spin label at A17 and S16 also caused the externalization of S 68IRS TMD1. Taking account these results suggest that some pinholin S21 residue positions such as G14, S16 and A17 have significant effect on the protein's structural topology as the changes to those positions induce significant structural perturbation. It is important to mention here that TMD1 has a putative glycine zipper (G10xxxG14xxxG18) which may have significant role on the structural conformation of 21 S 68IRS and certain changes in this region might have caused structural perturbations. Hence, these types of residues should be avoided during the SDSL to study the native structural topology of this system.

145

5.4.1 Pinholin Conformational changes observed with DEER Distance Measurements The CW-EPR power saturation data represent whether a majority of the population is located inside or outside of the lipid bilayer and are more applicable to studying the systems with a single conformation. DEER spectroscopy is a powerful biophysical technique that can provide direct evidence of the conformational changes of biomolecules and relative population of multiple conformations by monitoring the spin label distance distribution.41, 48-53 To further validate EPR power saturation results and directly provide the evidence of the effect of mutations on pinholin conformational change, the DEER spectroscopic technique was employed. For DEER 21 spectroscopic measurements, two spin labels were attached at S8 and L53 in S 68IRS and this construct was denoted as IRS_S8R1/L53R1. These positions were selected as an anchoring point since they are located at the N-terminus side of TMD1 and the C-terminus side of TMD2, respectively. These two residues were also found in close proximity in the inactive conformation 21 of antipinholin S 68IRS with a reported average distance of 26 Å (Unpublished data, Chapter 4). Additionally, no significant functional and structural perturbations have been reported for mutations at these positions.24 This construct will demonstrate a distinction of distance distribution 21 between the inactive conformation of pinholin S 68IRS against any structural perturbation induced by the mutation. Three more constructs were designed with single, double, and triple mutations based on our EPR power saturation results (see previous section above) and those reported by Pang et al. (2010).24 These constructs were denoted as IRS_S8R1/L53R1_A17Q, IRS_S8R1/L53R1_A17QG21Q, and IRS_S8R1/L53R1_G14A17QG21Q with the spin labels at the S8 and L53 positions. The DEER distance distributions obtained from DEER measurements were then compared with that of the IRS_S8R1/L53R1 (Figure 5.6).

146

Figure 5.6: Effect of mutations on DEER distance distributions of IRS_S8R1/L53R1 with or 21 without additional mutations. (A) inactive and active conformation of S 68IRS. Spin label positions are shown in green and mutation points are shown as a red letter in the orange circles. (B) Distance probability distributions are color-coded as the same color of the construct name. The highest distances distributions are normalized to one.

Inspection of the DEER distance distribution data (Figure 6B), it is clear that all mutant constructs showed a shorter distance peak centered around 26 Å similar to that of IRS_S8R1/L53R1, and another distance population at around 49 Å (Figure 5.6B). This longer distance population was not prominent for the single mutant IRS_S8R1/L53R1_A17Q (Figure 5.6B). However, the longer distance population was increased in the double mutation variant (IRS_S8R1L53R1_A17QG21Q) and was further pronounced for the triple mutation variant (IRS S8R1_L53R1_G14A17QG21Q) (Figure 5.6B). These data clearly demonstrated that increased

147 hydrophilicity at those residue positions enhanced the externalization of TMD1 and that a subpopulation was moving apart from TMD2, whereas others remained in close proximity as in the inactive conformation. The propensity of externalization increased with increased hydrophilicity and a cumulative effect was seen with the incorporation of additional mutations. This mutational effect is consistent with the power saturation data that indicated a higher number of populations of TMD1 were externalized when the hydrophilicity of TMD1 was increased. These results are also consistent with the mutational results reported by Pang et al. (2010) where G14Q, 21 A17Q, and G21Q mutations caused activation of the dominate antipinholin S 68IRS, although those mutants took additional time to trigger lysis.24 DEER data clearly indicate that the activation 21 of the antipinholin S 68IRS associated with the enhancement of the externalization of TMD1 due to the increased hydrophobicity of TMD1.

In addition to the mutational effects caused by natural amino acids, structural perturbations were also observed due to the presence of spin labels themselves, as was observed for CW-EPR power saturation experiments. A previous study showed that parallel distances between TMD1 and TMD2 of inactive pinholin were ~23-26 Å (Unpublished data; Chapter 4). However, several dual-spin labeled antipinholin constructs, IRS_S16R1/G48R1, IRS_S16R1/V46R1, and IRS_A20R1/L45R1 independently showed a longer distance distribution around 49 Å, 48 Å, and 45 Å, respectively (Figure 5.7). These distances match more closely with the active conformation of pinholin S21 instead of the inactive conformation which implies that the spin label itself caused a structural perturbation and externalized the TMD1 of antipinholin from the lipid bilayer.

148

21 Figure 5.7: Spin label induced activation of inactive antipinholin S 68IRS. Pair wise spin label 21 positions and probable distances are shown with arrows for active conformation of S (A). DEER time domain spectrum and corresponding distance distribution for IRS_S16R1/G48R1 (B), IRS_S16R1/V46R1 (C), IRS_A20R1/L45R1 (D). (The First spectrum was collected with 10 W amplifier).

In these DEER samples, one or both spin labels in a spin pair could induce structural perturbations due to the changes in intra and inter molecular interactions. In the earlier section (Figure 5.4), CW-EPR power saturation data demonstrated that IRS_S16R1 had an influence on the externalization of TMD1, which could be one of the reasons for the active conformation of IRS_S16R1/G48R1, and IRS_S16R1/V46R1 (Figure 5.7). In addition, it is worthy to mention here, G40xxxS44xxxG48 is the glycine zipper present in TMD2 and has significant effect on the structural conformation and oligomerization of pinholin S21. 24 Any residue in this region (e.g. L45, V46, G48) may contribute to the structural perturbation in addition to the mutational effects from TMD1. The mutational effects observed in the current study was based on the incorporation of the pinholin protein into the DMPC lipid bilayer. These mutational effects may vary in different lipid bilayers. Further studies are needed with different lipid bilayers to generalize the mutational affect in the membrane environment. Similarly, a more detailed study is needed to explore further mutational points and the underlying reasons for the structural perturbations caused by those residue mutations. 149

5.5 Conclusion This is the first EPR spectroscopic study which showed direct evidence of structural perturbations of pinholin S21 by positional mutations. This work also correlated these mutations with conformational changes and functionality of pinholin S21. The CW-EPR power saturation and DEER data clearly demonstrated that the relative hydrophobicity and interaction of different residues impact the observed conformational changes that have been directly linked to the pinholin S21 activation and triggering time. This study judiciously utilized CW-EPR power saturation and DEER spectroscopy to probe the effects of several amino acid residue mutations on antipinholin 21 S 68IRS TMD1 externalization. This approach will pave the way for the application of additional biophysical techniques for mutational studies in several other biologically significant systems especially those in which point mutations are likely related to functional/structural changes.

Acknowledgments We are grateful to the members of the Ry Young group at Texas A&M University for their experimental suggestions. This work was generously supported by the NSF CHE-1807131 grant, the NSF (MRI-1725502) grant, the Ohio Board of Regents, and Miami University. Gary A. Lorigan would also like to acknowledge support from the John W. Steube Professorship.

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47. Sahu, I. D.; Lorigan, G. A., Site-Directed Spin Labeling EPR for Studying Membrane Proteins. Biomed Research International 2018. 48. Sahu, I. D.; Lorigan, G. A., Biophysical EPR Studies Applied to Membrane Proteins. J Phys Chem Biophys: 2015; Vol. 5. 49. Jeschke, G.; Polyhach, Y., Distance measurements on spin-labelled biomacromolecules by pulsed electron paramagnetic resonance. Physical Chemistry Chemical Physics 2007, 9 (16), 1895- 1910. 50. Borbat, P. P.; McHaourab, H. S.; Freed, J. H., Protein structure determination using long- distance constraints from double-quantum coherence ESR: Study of T4 lysozyme. Journal of the American Chemical Society 2002, 124 (19), 5304-5314. 51. Vincent, E. F.; Sahu, I. D.; Costa-Filho, A. J.; Chilli, E. M.; Lorigan, G. A., Conformational changes of the HsDHODH N-terminal Microdomain via DEER Spectroscopy. J. Phys. Chem. B 2015, 119, 8693-8697. 52. Sahu, I. D.; McCarrick, R. M.; Troxel, K. R.; Zhang, R. F.; Smith, H. J.; Dunagan, M. M.; Swartz, M. S.; Rajan, P. V.; Kroncke, B. M.; Sanders, C. R.; Lorigan, G. A., DEER EPR Measurements for Membrane Protein Structures via Bifunctional Spin Labels and Lipodisq Nanoparticles. Biochemistry 2013, 52 (38), 6627-6632. 53. Sahu, I. D.; McCarrick, R. M.; Lorigan, G. A., Use of Electron Paramagnetic Resonance To Solve Biochemical Problems. Biochemistry 2013, 52 (35), 5967-5984.

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Chapter 6

Conclusions and Future Directions

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6.1 Summary of Dissertation Research Membrane proteins (MPs) are the key mediators of numerous cellular activities in biological systems.1-3 Approximately 20-30% of the total proteome of most cells are MPs which are associated with numerous physiological and pathological roles.3-8 As such, nearly 60% of currently available drug therapies target MPs.2, 9-11 For efficient drug design, it is crucial to know the structural details of the MP of interest due to the fact that stability and function are dictated by the MPs structural topology, conformational dynamics, and the surrounding environment.12-16 However, it is notoriously difficult to study the structure and dynamics of MPs due to several limitations including poor expression, complicated purification process, lipid environments, as well as limited biophysical techniques.

Chapter 1 (Part A) of this dissertation provides a comprehensive overview of MPs, the challenges associated with the study of MPs, and the application of EPR spectroscopy to overcome those challenges. This dissertation focuses on a structural study of a bacteriophage MP, pinholin S21, using state-of-the-art EPR spectroscopic techniques. Chapter 1 (Part B) of this thesis provides a comprehensive overview of the bacteriophage lytic system while emphasizing holin and pinholin. Holins are the group of MPs which form holes to permeabilize the cytoplasmic membranes of bacteria to initiate host cell lysis and control the phage infection cycle.17 Holin represents a diverse group of MPs which consists of ~900 proteins.18 A subclass of holin known as pinholin makes smaller holes than canonical holin and represents approximately one-fourth of holin proteins.19 Phage 21 encodes for one such holin system known as pinholin S21.19, 20 Pinholin S21 has two holin proteins: active pinholin S2168 and inactive antipinholin S2171.20 Both pinholin proteins have a short N-terminus region followed by two transmembrane domains (TMDs) that are connected through a short loop and terminating in a long, positively charged C-terminus.21 To overcome the challenge associated with expression and purification of MPs, solid phase peptide synthesis and reverse phase-HPLC was used to synthesize and purify pinholin S21 for the structural studies reported in this dissertation.

The work in Chapter 2 was focused on determining the structural topology and dynamics of active pinholin S2168 using Continuous Wave (CW)-EPR line shape analysis and power saturation techniques. Systematic site-directed spin labeling (SDSL) was used to introduce the R1 side chain for 39 residue positions of S2168. Pinholin S2168 was incorporated into DMPC

157 proteoliposomes to mimic the plasma membrane environment. Circular dichroism (CD) spectroscopic data confirmed that pinholin S2168 maintains a predicted native α-helical secondary structure in DMPC proteoliposomes. The calculated dynamic parameters clearly illustrated the restricted mobility of the pinholin S2168 TMDs and the higher mobility of the N- and C-termini regions. The depth parameter calculated from CW-EPR power saturation data indicated that TMD1 was partially externalized from the lipid bilayer and interacted with the membrane surface, whereas TMD2 remained buried in the lipid bilayer in the active conformation of pinholin S2168. At the end of this chapter, a tentative structural topology model of pinholin S2168 was presented based on collected EPR spectroscopic data.22

The work presented in Chapter 3 was carried out to determine the structural topology and 21 dynamics of antipinholin S 68IRS in DMPC proteoliposomes using CW-EPR line shape analysis and power saturation techniques. The native phage 21 antipinholin is S2171 has 71 amino acids and is a weak antipinholin as it achieves the same conformation as active pinholin after an initial 21 delay. However, S 68IRS is a dominant antiholin that was generated by the addition of five amino acids (RYIRS) to the N-terminus of S2168 as these additional residues preserve the inactive 20, 23 21 conformation of pinholin. CD data confirmed the α-helical secondary structures of S 68IRS in solution with or without a bound nitroxide spin-label. CW-EPR line shape analysis experiments 21 of 35 different R1 side chains of S 68IRS indicated restricted mobility of the TMDs which were predicted to be inside the lipid bilayer as compared with the N- and C-termini R1 side chains. In addition, R1 accessibility experiments performed on 24 residues using CW-EPR power saturation 21 experiments indicated that TMD1 and TMD2 of S 68IRS were incorporated into the lipid bilayer while the N- and C-termini were located outside of the lipid bilayer. Based on this study, a tentative 21 structural model of S 68IRS was proposed in which both TMDs remain in the lipid bilayer while the N- and C-termini were located outside of it.24

In Chapter 4, double electron-electron resonance (DEER) spectroscopy was used for detailed structural study and direct comparison of the active and inactive forms of pinholin S21. DEER spectroscopy coupled with SDSL is a powerful biophysical tool for long-range distance (20-80 Å) measurements used to elucidate the structural and conformational properties of biomolecules.25-31 In this study, the structural topology and conformation of active pinholin S2168 21 and antipinholin S 68IRS in DMPC proteoliposomes was investigated using a four-pulse DEER

158 sequence to measure the distances between TMD1 and TMD2. Five inter-label distances were measured for both the active and inactive forms of pinholin. A model for the ten lowest-energy structures of the active pinholin and inactive antipinholin in DMPC proteoliposomes were obtained using the experimental DEER distances coupled with the simulated annealing software package Xplor-NIH. In the proposed model, TMD2 of active pinholin S2168 remains in the lipid bilayer and TMD1 is partially externalized from the bilayer with some residues located on the surface. 21 However, both TMDs of inactive S 68IRS remain incorporated in the lipid bilayer. This study clearly demonstrated the differences in structural topology and conformations between the two forms of pinholin S21. The results from this study validated the structural model for active and inactive conformations of pinholin reported in our earlier reports.22, 24, 32

Chapter 5 is focused on a novel approach to study the effects residue mutation on the structural topology and conformational changes of pinholin S21 using EPR techniques. Phages use a predefined combination of pinholin and antiholin concentrations to define the timing of host lysis.3, 22 In addition, phages tune their lytic function by the residue mutation of pinholin and antiholin to adjust to different host environments.3, 22 Typically, the effects of pinholin residue mutation are probed by monitoring the timing of cell lysis which is not enough to define pinholin activity, as SAR endolysin alone can cause cell lysis after induction, regardless of expression of the pinholin allele.22, 24 This study reports the effects of various mutations on the activation or 21 inactivation of inactive pinholin S 68IRS using CW-EPR power saturation and DEER spectroscopic techniques. This work directly evaluated the structural and conformational changes of pinholin S21 induced by residue mutations and correlated the conformational changes with activation or inactivation of the pinholin S21. CW-EPR power saturation data indicated that 21 increased hydrophilicity of S 68IRS TMD1 accelerated its externalization from the lipid bilayer as relative to wildtype. DEER experiments were used to probe how different amino acid mutations 21 changed the relative populations of active vs inactive conformations of S 68IRS. This mutational study clearly demonstrated that the relative hydrophobicity of TMD1 impacts its externalization which further controls the activation of inactive dimers and ultimately the formation of pinholes. This study also illustrated how the interactions between TMD1 and TMD2 were changed by mutations that impacted the externalization of TMD1 and changed host lysis triggering time. These results provide a more comprehensive picture of how conformational changes influence the

159 functionality of pinholin systems by defining of the effects of residue mutation on S21 pinholin structural topology and conformational change.

Finally, the works presented in this dissertation reflect the detailed structural dynamics, conformational changes, and topology of the active and inactive conformations of pinholin S21 as determined by several different EPR spectroscopic techniques. The work presented in this dissertation have helped refine the structural model of active pinholin reported previously in the literature.33 We have also clearly demonstrated how to apply EPR spectroscopic techniques to mutational studies of membrane proteins. Additionally, we are the first research group to use a synthetic pinholin S21 peptide in biophysical studies and our work will pave the way for the structural study of many other challenging membrane proteins.

6.2 Future Directions

6.2.1 Oligomerization of Pinholin S21 Studied Using DEER Spectroscopy Pinholin S21 is incorporated into the cytoplasmic membrane as inactive monomers that further associate into inactive dimers.20, 23 When the critical concentration of active dimers is reached, they interact with other adjacent dimers and form oligomers that make the nanoscale membrane holes.20, 23, 34 Hence, pinhole formation is a concentration-dependent dynamic equilibrium process among the monomer, inactive dimers, active dimers, 2D aggregates, and heptamers.20 Currently, there have been no detailed biophysical studies to directly probe the concentration-dependent oligomerization of full-length pinholin using a suitable biophysical technique. DEER spectroscopy is a powerful biophysical technique that can be used to study the oligomeric states of proteins.35-38 In DEER experiments, for a given sample, the modulation depth depends on the fraction of specifically interacting spin pairs and the distance between them that are excited by the microwave pulse. For a single spin-labeled protein, if the protein is a monomer with a random distribution of inter-spin distances, there will be no specific spin interaction and the modulation depth will be zero. If pinholin forms an oligomer, an increase in spin-labeled protein concentration, will cause the spin labels to come in close enough contact to form spin pairs that will cause the modulation amplitude to increase indicating the formation of oligomers.36-38 Moreover, DEER data will provide the distances between monomers which can be used to evaluate

160 how the oligomeric states are formed as well as the orientation of monomers in the oligomer complexes.35-38

6.2.2 CW-EPR Alignment Techniques to Probe the Relative Orientation of TMD1 and TMD2 CW-EPR alignment techniques are useful tools for predicting the relative orientation and calculating helical tilt of TMDs with respect to the membrane normal. The hyperfine tensorial components of a rigid spin-label (e.g. TOAC) depends on its relative orientation within a magnetic field while significantly changing the hyperfine splitting values at various orientations. Using this parameter, the relative orientation and helical tilt angle of the transmembrane peptides can be calculated for magnetically or mechanically aligned samples.39, 40 From the structural topology of active and inactive pinholin, it was found that TMD1 aligns parallel or perpendicular to the lipid bilayer if the pinholin S21 is in the active and inactive conformation, respectively. This can be further validated by the CW-EPR alignment technique with a TOAC labeled pinholin S21. It was also predicted that TMD2 has a certain degree of helical tilt which can be calculated using a CW- EPR alignment study.20, 39, 40

6.2.3 Application of ESEEM to probe the helical length Electron spin echo envelope modulation (ESEEM) spectroscopy is used to gain information about NMR-active nuclei near an unpaired electron spin system and is also known as EPR-detected NMR spectroscopy.41, 42 This method can be used to identify the local secondary structure of a membrane protein as well as being able to differentiate between beta-sheet, alpha- 41-43 helix, and 310-helix motifs. ESEEM techniques can also be used to determine the actual length of α-helical regions. There is no experimental evidence of the residue or distance lengths of the pinholin S21 α-helical TMDs. The previously reported helix lengths were predictions based on mathematical algorithms and these values differ from the helical lengths we report in Chapter 4. For these experiments, a deuterated amino acid would be placed near the predicted end of the α helix and a spin-label will be placed at various distances from the deuterated amino acid.

6.3 Final Remarks Holins and holin-like proteins serve a variety of functions and are ubiquitous and present in all three domains of life including bacteria, eukaryotes, and archaea.44 Despite its ubiquitous presence and potential application in biotechnology and the pharmaceutical industry, very little structural information is known for this group of proteins.44-52 Like many other MPs, structural 161 study of holins has been severely limited by challenges associated with expression, purification, and incorporation into stable membrane-like native environment as well as biophysical techniques that are compatible with lipid bilayers. This dissertation highlights the application of solid-phase peptide synthesis and reverse phase-HPLC to overcome the problems associated with expression and purification of membrane proteins. Furthermore, exploration of the application of EPR spectroscopic techniques to a wide variety of holin systems will provide for a greater in-depth knowledge of these otherwise challenging to study systems. A continued exploration of EPR spectroscopy will allow this approach to continue to emerge as a widely accepted tool for examining a wide variety of membrane proteins.

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