<<

MIAMI UNIVERSITY The Graduate School

Certificate for Approving the Dissertation

We hereby approve the Dissertation

of

Daniel Lee Drew Jr

Candidate for the Degree

DOCTOR OF PHILOSOPHY

______Dr. Gary A. Lorigan, Director

______Dr. Rick Page, Reader

______Dr. Ellen Yezierski, Reader

______Dr. David L Tierney, Reader

______Dr. Paul Urayama, Graduate School Representative

ABSTRACT

INVESTIGATING THE AND DYNAMIC PROPERTIES OF BACTERIOPHAGE S21 PINHOLIN USING SOLID-STATE NUCLEAR MAGNETIC RESONANCE AND PARAMAGNETIC RESONANCE

by

Daniel L Drew Jr.

Holins are a family of lytic membrane responsible for the lysis of the cytosolic membrane in host cells of double stranded DNA bacteriophages. Recently a new family of holins have been discovered, the pinholin, which have been shown to depolarize the cytosolic membrane leading to the release of membrane-bound signal-anchor-released (SAR) endolysin from the bilayer. Despite the biological importance of pinholin the structure and dynamic properties have not been well characterized. The work in this dissertation will use a variety of powerful biophysical techniques to study these structure and dynamic properties of pinholin. First, we report the first in vitro synthesis of both the active S2168 and inactive S21IRS pinholin using solid phase peptide synthesis (SPPS) and reveals the first experimental data indicating the global α-helical structure of pinholin using circular (CD) spectroscopy. After sample optimization, electron spin echo envelope modulation (ESEEM) spectroscopy was used to determine the local α-helical secondary structure of the inhibitory and functional helices of pinholin. Following that, SS-NMR spectroscopy was utilized to probe the differences in the way the active and inactive forms of pinholin interact with the membrane and gives the first piece of direct quantitative evidence indicating TMD1 interacts with the lipid headgroups of the bilayer. Finally, DEER spectroscopy was used to probe the structural models of the active and inactive form of pinholin the membrane. This work led to a newly proposed model of active S2168 pinholin with TMD1 partially externalized from the membrane. This study expanded the application of SS-NMR and EPR spectroscopic techniques and provided a deeper understanding of the structure and dynamic properties of the complex pinholin membrane system.

INVESTIGATING THE STRUCTURE AND DYNAMIC PROPERTIES OF BACTERIOPHAGE S21 PINHOLIN USING SOLID-STATE NUCLEAR MAGNETIC RESONANCE AND 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 and Biochemistry

by

Daniel Lee Drew Jr

The Graduate School Miami University Oxford, Ohio

2021

Dissertation Director: Dr. Gary Lorigan

©

Daniel L Drew Jr

2021 TABLE OF CONTENTS

Chapter 1: Introduction Membrane Proteins, Solid-Phase Peptide Synthesis, Electron Paramagnetic Resonance, and Nuclear Magnetic Resonance………………………………….………..1 1.1 Membrane Proteins……………………………………………..…………………..2 1.2 The S21 Bacteriophage Lytic System………………………………….…………..4 1.3 Solid Phase Peptide Synthesis…………………………………………………….7 1.4 Electron Paramagnetic Resonance Spectroscopy………………………………9 1.5 Site Directed Spin Labeling……………………………………………………….10 1.6 Continuous Wave EPR……………………………………………………………11 1.7 Pulsed EPR Spectroscopic Techniques…………………………………………12 1.8 Solid State Nuclear Magnetic Resonance Spectroscopy………………………13

Chapter 2 Solid Phase Synthesis and Spectroscopic Characterization of the Active and Inactive Forms of Bacteriophage S21 Pinholin Protein…………………………………21 2.1 Abstract…………………………………………………………………….……....22 2.2 Introduction…………………………………………………………………..…….23 2.3 Materials and Methods…………………………………………….……………...25 2.3.1 Solid Phase Peptide Synthesis…………………………….…………..25 2.3.2 Protecting Group and Solid Phase Cleavage ……………….………..26 2.3.3 and Spin Labeling………………………….……..26 2.3.4 Peptide Incorporation into Lipid Mimetic Systems……………….…...27 2.3.5 Circular Dichroism…………………………………………….………...27 2.3.6 Continuous Wave EPR Spectroscopy………………………….……..28 2.3.7 31P Solid State NMR Spectroscopy……………………………..……..28 2.4 Results and Discussion…………………………………………………………...28 2.4.1 Optimization of Solid Phase Peptide Synthesis…………………..….29 2.4.2 Optimization of Peptide Cleavage and HPLC Purification………..….29 2.4.3 Optimization of MALDI-Tof Conditions and Matrix……………………30

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2.4.4 Circular Dichroism of S2168 and S21IRS Pinholin……………………..30 2.4.5 CW-EPR Measurements of Pinholin……………………………..……31 2.4.6 31P Solid State NMR Spectroscopy of Pinholin………………………33 2.5 Conclusions……………………...…………………………………………………34 2.6 Acknowledgments…………………...…………………………………………….34 2.7 Figures……………………………………..……………………………………….35

Chapter 3 Application of ESSEEM Detected Secondary Structure Technique to the Functional Pinholin System………………………………………………………………………………44 3.1 Abstract…………………………………………………………………………..…45 3.2 Introduction ………………………..…………………………………………..…..46 3.3 Experimental Methods…………………………………………………………….48 3.4 Results and Discussion……………………………………………………………51 3.5 Conclusions………………………………………...…………………………...... 54 3.6 Acknowledgments.……………………………………...…………………………55 3.7 Figures………………………………………………………...………………...….56

Chapter 4 Active S2168 and Inactive S21IRS Pinholin Interact Differently with the Lipid Bilayer: A 31P and 2H Solid State NMR Study………………………………………………………62 4.1 Abstract……………………………………………………………….…………….63 4.2 Introduction ………………………………………………….……………………..64 4.3 Materials and Methods…………………………………………..………………..65 4.3.1 Solid Phase Peptide Synthesis………………………………………...65 4.3.2 Protein Purification……………………………………………...... ……66 4.3.3 Proteoliposome Sample Preparation……..………………………...... 66 4.3.4 Solid State Nuclear Magnetic Resonance Spectroscopy……………67 4.4 Results and Discussion……………………………………………………….…..68 4.4.1 31P SS-NMR Chemical Shift Anisotropy Analysis…………………….68 4.4.2 2H SS-NMR Quadrupolar Splitting Analysis………………………..…70

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4.4.3 Depaking and Order Parameters (SCD)……………….………..……...71 4.5 Conclusions……………………………………………………...…………..…….73 4.6 Acknowledgments………………………………………………………………....75 4.7 Figures………………………………………………………………...……………76

Chapter 5 Pulsed EPR DEER Spectroscopic Study of Bacteriophage S21 Pinholin Reveals Two Different Structural Conformations Between the Active and Inactive Form………..94 5.1 Abstract…………………………………………………………..……..……..…. .95 5.2 Introduction……………………………………………………………..……….... 96 5.3 Experimental Methods…………………………………………………………….96 5.4 Results and Discussion……………………………………………………………98 5.5 Conclusions……………………………………………………………………....100 5.6 Acknowledgments………………………………………………………………..100 5.7 Figures…………………………………………………………………………….101

Chapter 6 Conclusions and Future Directions…………………………………………………...…108 6.1 Summary of Work……………………………………………………………...…109 6.2 Future Directions……………………………………...………………………….113 6.2.1 ESEEM Spectroscopic Studies……………………………………….113 6.2.2 SS-NMR Spectroscopic Studies……………….………………...…..113 6.2.3 DEER Spectroscopic Studies…………………….…………………..114

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LIST OF TABLES

Table 3.1 Primary sequence of active S2168 and inactive S21IRS pinholin outlining the experimental ESEEM d10-Leu and spin labeling positions…………...……….…………...49 Table 4.1 Experimental 31P SS-NMR chemical shift anisotropy (CSA) widths for active S2168 and inactive S21IRS pinholin at all concentrations and ……………...80 Table 4.2 Experimental 2H SS-NMR quadrupolar splitting values for active S2168 and inactive S21IRS pinholin at all concentrations and temperatures ….………………………83

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LIST OF FIGURES

Figure 1.1 Diagram of different membrane protein …………………….……..…2 Figure 1.2 A schematic showing the complex relationship between membrane and function …………………………………………………………………………..3 Figure 1.3 The primary sequence of S2168 pinholin and S2171 antipinholin ……...... 6 Figure 1.4 Flow diagram of Solid Phase Peptide Synthesis cycles ……………..……..…8 Figure 1.5 Energy level diagram of an unpaired electron with a spin of S=1/2…………10 Figure 1.6 MTSL structure and site directed spin labeling reaction………………………11 Figure 1.7 Energy level diagram of MTSL nitroxide spin label……………………………12

Figure 1.8 Structure of d54-DMPC lipid with deuterated acyl chains………………………14 Figure 2.1 Primary sequence for active S2168 pinholin, the cognate S2171 antipinholin, and the inactive S21IRS pinholin………………………………………………………………35 Figure 2.2 The schematic showing the hypothetical pinholin TMD1 and TMD2 conformations in the membrane………………………………………………………………36 Figure 2.3 MALDI-TOF mass spectrum of pure active S2168 and inactive S21IRS………37 Figure 2.4 Circular Dichroism spectra of both the active S2168 and inactive S21IRS pinholin……………………………………………………………………………………….…38 Figure 2.5 CW-EPR spectra of free MTSL, MTSL coupled to active S2168 pinholin in TFE, and MTSL labeled pinholin incorporated into lipid mimetic systems……………………..39 Figure 2.6 Static 31P solid-state NMR spectrum of empty DMPC MLVs, and the active S2168 and inactive S21IRS forms of pinholin…………………………………………………40 21 Figure 3.1 CD spectra of the active S 68 – d10-Leu50 (i-3) pinholin incorporated into DMPC MLVs……………………………………………………………………………………56 21 Figure 3.2 Three pulse ESEEM experimental data for active S 68 – d10-Leu25 pinholin incorporated into DMPC liposomes………………………………………………………….57 Figure 3.3 Three pulse ESEEM experimental data for active S2168 and inactive S21IRS pinholin at Leu25 and Leu50 positions incorporated into bicelle and liposome mimetic systems………………………………………………………………………..………………..58 Figure 4.1 The structural models of active S2168 and inactive S21IRS with the of d54-DMPC lipid……………………………………………………………………76

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Figure 4.2 dependent 31P SS-NMR spectra of active S2168 pinholin…...... 77 Figure 4.3 Temperature dependent 31P SS-NMR spectra of inactive S21IRS pinholin.....78 Figure 4.4 Comparison of 31P chemical shift anisotropy (CSA) width at 35°C for active S2168 and inactive S21IRS pinholin…………………………………………………………...79 Figure 4.5 Temperature dependent 2H SS-NMR spectra of active S2168 pinholin……...81 Figure 4.6 Comparison of the 2H quadrupolar splitting at 35°C between active S2168 and inactive S21IRS pinholin………………………………………………………………………..82 Figure 4.7 The original 2H SS-NMR spectra and the resulting dePaked spectra of empty and 1 mol% active S2168 proteoliposomes………………………………………………….84 21 Figure 4.8 Order Parameters (SCD) for empty and 1 mol% active S 68 proteoliposomes at increasing temperatures…………………………………………………………………....85 21 Figure 4.9 Order parameter (SCD) comparison of empty and 1 mol% active S 68 pinholin at 35°C…………………………………………………………………………………………..86 21 Figure 4.10 Order parameter (SCD) comparison of empty and 1 mol% active S 68 pinholin at 25°C……………………………………………………………………………...…87 Figure 4.S1 The dePaked 2H spectra of 0 mol%, 1 mol%, and 2 mol% S2168 at each temperature………………………………………………………………………………...... 88 Figure 4.S2 The overlaid dePaked spectra of 0 mol% and 1 mol% active S2168 pinholin at 35°C…………………………………………………………………………………..……...89 Figure 4.S3 The overlaid dePaked spectra of 0 mol% and 1 mol% active S2168 pinholin at 25°C…………………………………………………………………………………....…….90 Figure 5.1 The current functional model for the structural conformations of active S2168 and inactive S21IRS pinholin…………………………………………………………………101 Figure 5.2 Ribbon diagram of inactive S21IRS DEER substitution pairs and corresponding frequency modulation and distance distributions………………………………………….102 Figure 5.3 Ribbon diagram of active S2168 DEER substitution pairs and corresponding frequency modulation and distance distributions…………………………………...……..103 Figure 5.4 A preliminary molecular dynamic model comparison of the two proposed active S2168 pinholin conformations……………………………………………………...... 104 Figure 5.5 Newly proposed structural model of active S2168 pinholin………………….105

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DEDICATION

The work discussed in this dissertation is dedicated to my family and friends, without whom this would not have been possible.

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ACKNOWLEDGMENTS

Firstly, I would like to thank my advisor Dr. Gary A. Lorigan for the opportunity to work in his group and for all the help and support he gave me along the way.

I would also like to thank all the members of my committee for their guidance and advise throughout the years: Dr. Rick Page, Dr. David Tierney, Dr. Ellen Yezierski, and Dr. Paul Urayama.

I would like to thank both Dr. Robert McCarrick and Dr. Theresa Ramelot for training and lending me their expertise in regard to the EPR and NMR instrumentation.

I am thankful to all the members of the Lorigan group that I had the privilege of working with over years. Dr. Lishan Liu and Dr. Lauren Bottorf for training me when I was first starting out. Dr. Indra D. Sahu for all his help when it came to writing and editing. Dr. Rongfu Zhang for showing me hard work doesn’t have to come at the expense of a few laughs. Dr. Gunjan Dixit for all the different ways we’ve helped each other and for being my tether to sanity during the last years of our dissertation work. Tanbir Ahammad for taking up the pinholin mantel and always being there to discuss theories and hypotheses. Rachel Serafin for being the most dedicated undergraduate researcher I’ve had the pleasure of working with. Brandon Butcher, Rasal Khan, Sophia Rafferty, Andrew Craig, F. Dhilhani Mohammad and so many others.

Finally, I want to thank my mother, father, and brothers, as well as the rest of my family and Ashley Flye for their unconditional love and support.

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Chapter 1: Introduction

Membrane Proteins, S21 Bacteriophage Lytic System, Solid-Phase Peptide Synthesis, Electron Paramagnetic Resonance, and Solid-State Nuclear Magnetic Resonance

1

1.1 Membrane Proteins Membrane proteins play critical roles in biological systems. Nearly one third of all the proteins expressed by the human genome are membrane proteins and have been shown to be responsible for a variety of important biological functions, including cell signaling, ion and transport across membranes, and participants in energy transduction.1-5 Due to their diversity in function, membrane proteins are the target for more than half of the therapeutics and drugs available on the market.6-9 For these vital membrane protein systems to function properly they must fold into a specific secondary, and possibly tertiary and quaternary, structures unique to that membrane protein’s given function.10, 11 The different membrane protein structures and their corresponding functions is shown in Figure 1.1 below.

Figure 1.1 The different secondary and tertiary structures of membrane proteins with the corresponding function these types of structures can perform. (Copyright Dharmesh Patel)

Mutations in the membrane protein sequence and misfolding of the protein are responsible for some of the most severe human disorders and diseases including Alzheimer’s and Parkinson’s disease.12-14 Despite the importance of membrane protein roles in biological functions and the disastrous effects of mutations and misfolding the understanding between structures and function is still lacking.10 A thorough

2 understanding of protein structure and dynamics within the membrane helps to elucidate the pathway of function for these proteins. A schematic of the complex relationship between structure and function is shown in Figure 1.2.

Figure 1.2 A schematic showing the complex relationship between membrane protein structure and the overall function.

Equally important is the use of this information to cast on the mechanism through which diseases manifest at the cellular level. Although the importance of structural and dynamic information is well understood the quantity of unique membrane protein structures present in the Protein Data Bank (PDB) is drastically underrepresented at 0.5% of the total structures.15-17 There are currently 153,601 unique structures in the PDB with ~89% of those structures coming from X-ray diffraction, ~10% from protein NMR, and the remaining small percent a mix of electron microscopy, Cryo-EM and hybrid studies using multiple techniques.18 Of the 153,601 protein structures deposited in the PDB only 1644 represent structures of membrane proteins with only 919 being unique membrane protein structures.19 This underrepresentation is due to the notorious and inherently difficult nature of studying membrane proteins.16, 20 The initial problem to overcome is obtaining enough structurally and functionally stable protein. Typically, this is accomplished through overexpression of the target protein in E. coli, or some other bacterial system, but can often result in low expression yields.20, 21 Additionally, a number of mammalian membrane proteins require some degree of post-

3 translational modification accomplished through the cellular machinery of mammalian cells that is lacking in the bacterial cells.22 The absence of these modifications can affect the membrane insertion, folding/structure, and overall function of the membrane proteins.20, 22, 23 Another difficulty present in the study of membrane proteins is the amphipathic nature of the proteins. Most membrane proteins have highly hydrophobic portions which interact with the hydrophobic acyl chains comprising the core of the membrane. As seen in Figure 1.1, these sections can be comprised of singular or multiple helical domains, β- strands, or β-barrels, and are highly sensitive to the environment surrounding them. Membrane proteins can have hydrophilic domains of varying sizes and charges. These sections have been involved in a variety of roles ranging from folding as functional soluble domains to interacting with the charged lipid headgroups of the lipid bilayer.24 The duality of the hydrophobic and hydrophilic sections present on these membrane proteins makes it extremely important to reconstitute these proteins into synthetic membrane mimetic systems.25 Over the years there have been several synthetic membrane systems devised, such as detergent micelles, bicelles, vesicles, and nanodiscs, however, due to the varying degree of amphipathicity and charges present on each protein there is no one best mimetic and therefore each membrane protein system requires careful optimization. Ultimately, the lipid bilayer itself adds an additional layer on of complexity as the membrane is a heterogeneous and dynamic environment.26 This limits the application of many powerful biophysical techniques, such as x-ray crystallography, nuclear magnetic resonance (NMR), and circular dichroism (CD), all of which have no such fewer limitations when studying solution proteins. Fortunately, there are a diverse number of biophysical techniques, such as (MS), spectroscopy, solid-state NMR spectroscopy, and EPR spectroscopy, that can be utilized to investigate structures and dynamics of complex membrane proteins systems.

1.2 The S21 Bacteriophage Lytic System The ultimate step of double stranded DNA bacteriophage infection is lysis of the host cell.27, 28 The mechanism of lysis is a highly regulated process divided into three steps with which each step involving the presence of a specific protein.29-31 The first step

4 is the permeabilization of the inner cytosolic membrane accomplished by a hole forming membrane protein known as holin.32 The holin protein accumulates in the cytosolic membrane harmlessly until the protein “triggers” at an allele-specific time.33 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.33, 34 Due to the variation in mechanisms and sizes of lesions formed between different classes of holins the lesions have been termed “holes” to show distinction from channels and other such membrane permeabilization pathways. Upon hole formation, the non-selective escape of the second required protein from the cytoplasm begins.32, 35 This fully folded and functional enzyme is known as the endolysin and is responsible for the degradation of the peptidoglycan. The final step in the lysis pathway requires the membrane-bound spanin protein, which fuses the inner and out cell membranes and allows for the escape of newly constructed bacteriophage to continue the bacteriophage infection cycle. Until recently it was believed that all holins, like the λ S105 canonical holin, trigger to form a non-selective micron-scale hole in the inner cytoplasmic membrane.35 Recent discoveries have uncovered a new class of holin and endolysin in the S21 gene of lambdoid bacteriophage φ21.36, 37 Instead of forming a large non-selective hole in the cytoplasmic membrane, these holins form nanometer-scale holes that are only responsible for the depolarization of the membrane.37, 38 Due to the small size of the holes, this new class of holin was named the pinholin.39 Unlike the large hole forming canonical holins the hole created by the pinholins is not large enough to allow for the nonspecific escape of functional endolysin from the cytoplasm. Instead these pinholins are paired with signal-anchor-release (SAR) endolysins.40-42 These proteins are named for a special N-terminal transmembrane helix that acts as an uncleaved signal sequence, resulting in a sec-mediated export.40 The SAR-endolysins accumulate in the periplasm as membrane-tethered, inactive enzymes. This prevents the premature degradation of the peptidoglycan layer. Upon pinholin triggering and the resulting membrane depolarization, the SAR domain freely exits the bilayer. Once untethered from the membrane the periplasmic catalytic domain refolds to the active form of the endolysin and begins degradation of the peptidoglycan.41, 42 This work will focus on the under characterized pinholin protein and pathway of membrane depolarization.

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Pinholin is a small amphipathic membrane protein comprised of two transmembrane domains (TMDs). Due to the presence of a dual start motif in the S21 gene there are two gene products for pinholin expression, S2171 ensuing from translational codon Met1, and S2168 resulting from Met4.36 The primary sequence for these two pinholin are shown in Figure 1.3.

Figure 1.3 The primary sequence of S2168 pinholin and S2171 antipinholin with predicted transmembrane domains underlined and the additional three amino acids of the antipinholin in red.

The S2168 is the active form of pinholin and is composed of 68 amino acids, with a short N-terminus, positively charged C-terminus, and two TMDs. S2171 is known as the antipinholin which is 71 amino acids in length, contains the same two TMDs, but has an additional three amino acids located on the N-terminus.36 Both the pinholin and antipinholin externalize the first TMD (TMD1) from the membrane, while the second (TMD2) remains incorporated into bilayer. This translocation of TMD1 is essential for the lytic function of pinholin and the ultimate depolarization of the membrane.43, 44 The antiholin is responsible for the delayed triggering of the pinholin due to the addition of a positively charged lysine on the N-terminus which drastically slows the externalization of TMD1 from the membrane. The overall lytic pathway initiates with the expression and accumulation of both the S2168 pinholin and S2171 antipinholin embedded in the bilayer. Starting as inactive monomers the proteins begin to interact and accumulate as dimers. Since the S2168/S2171 production ratio is ~2:1, the dominant dimer formations are that of the homodimers S2168:S2168 and heterodimers S2168:S2171.45 These dimers are in an inactive state where activation is dependent on the externalization of TMD1 from the bilayer for each monomer. Once both TMD1 segments have translocated, the pinholin dimer is known to be in an active form and continues through the lytic pathway by forming

6 oligomers.45 When concentration of the activated pinholin dimers reach a critical concentration, the pinholin population triggers and causes massive and sudden depolarization of the inner cell membrane, resulting in the release of SAR endolysin from the cytosolic membrane. In this study the structure, mechanism, and model pathway of the pinholin protein will be probed using powerful biophysical techniques such as circular dichroism (CD), solid-state nuclear magnetic resonance (SS-NMR), and ESEEM and DEER electron paramagnetic resonance (EPR) spectroscopy.

1.3 Solid Phase Peptide Synthesis Solid Phase Peptide Synthesis (SPPS) was pioneered by Robert Bruce Merrifield with his 1963 publication “Solid Phase Peptide Synthesis. I. The Synthesis of a Tetrapeptide.”46 Since then, SPPS has been improved through a variety of ways including microwave irradiation, addition of chemical compounds to reduce the required reaction energy, and reaction automation. All of these have led SPPS to become a widely used approach in many biochemistry and biophysical laboratories. The most common SPPS technique used in labs today, including the Lorigan Lab, is Fmoc protected SPPS due to its ability to produce high yields of peptide on a fast timescale and at low costs.47 The SPPS used for the following studies is a CEM Liberty Blue Synthesizer and is coupled with a Discovery-Bio microwave unit. Due to the sequential coupling of amino acids the yield and purity of SPPS is dependent on the length of peptides with a generally accepted limit of 70 to 80 amino acids.48 Each step in the synthesis is a series of repeated cycles which follows as N- terminus deprotection, washing away of the protecting group, coupling of the new amino acid, and finally washing away of any uncoupled amino acid. A flow diagram of these steps is outlined in Figure 1.4.

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Figure 1.4 A flow diagram of the cycles followed in Solid Phase Peptide Synthesis.49

The typical solid phase used in SPPS is a porous bead, known as a resin, and is usually composed of polystyrene (PS) or polyethylene glycol (PEG).50 These resins can either be purchased as blank resins or can come preloaded with the first amino acid in the sequence already attached. The desired peptide sequence can then be synthesized from C-terminal to the N-terminal following the series of steps outlined previously. All amino acids used in SPPS have an N-terminal protecting group, typically a Boc or Fmoc group, which must be removed before the addition of the next amino acids.51 Several amino acids with side chain functional groups require additional protecting groups which cannot be removed during the Fmoc or Boc deprotection. This is to prevent coupling of the next amino acid to the side chain of the previous residue creating a branch point and resulting in undesired product. At the end of the synthesis, the desired peptide will remain covalently bound to the resin and must be cleaved from the solid support using trifluoroacetic acid (TFA).52-54 This step will also remove the side chain protecting groups still present on many amino acids. Scavengers, such as water, anisole, 1,2-Ethanedithiol

8

(EDT), or triisopropylsilane (TIPS) can be added to the cleavage reaction to irreversibly bind to the protecting groups thus making them unreactive. After this, the peptide is able to be purified by high pressure liquid chromatography (HPLC) and confirmed using matrix assisted laser desorption ionization – time of flight (MALDI-Tof). The pure protein can then be coupled with the nitroxide spin label MTSL in order to conduct EPR experiments.55

1.4 Electron Paramagnetic Resonance (EPR) Spectroscopy Electron paramagnetic resonance (EPR) spectroscopy is a powerful biophysical technique used to study the structure, dynamics and electronic properties of systems containing unpaired .56-58 The presence of paramagnetic centers in organic radicals and complexes make them an ideal candidate for studies using EPR spectroscopy. With the introduction of site-directed spin labeling (SDSL), the technique is now routinely used to study the structure and dynamic properties of several biological systems including proteins and nucleic acids.55, 59 EPR spectroscopic studies involve electronic spin transitions unlike nuclear spin transitions in nuclear magnetic resonance (NMR) spectroscopy, making it a more sensitive technique for biophysical studies. The basic theory behind EPR transitions involve an unpaired electron spin residing in a . With an electronic spin of S= ½, this electron can be in a +1/2 or -

1/2 Ms quantum state. These states are usually degenerate in the absence of any magnetic field. However, when the magnetic field is applied, these states separate out with +1/2 state occupying a higher energy level and -1/2 with a lower energy state. Upon reaching resonance, these electrons can transition between the ground state and the exited state by absorbing a microwave with an energy, h= E.60 The energy difference can be given by,

∆퐸 = h = 𝑔훽푒퐵표 where, h is the Plank’s constant,  is the frequency of the photon, g is the electronic g- factor, 훽푒 is the Bohr Magnetron (Joules*Gauss-1), and 퐵표 is the applied magnetic field

9

(Gauss). The energy level diagram of an unpaired electron in the presence of magnetic field is shown in Figure 1.5. An EPR experiment measures an absorption spectrum, and the first derivative of this absorption spectrum is recorded.

Figure 1.5 The energy level diagram of an unpaired electron in a magnetic field showing the absorption peak and corresponding first derivative EPR signal.

EPR spectroscopy can provide higher sensitivity than NMR due to the inherent higher magnetic moment of an electron when compared to an NMR active nucleus. The technique is not limited by size, requires M range of sample concentration, L sample volume, and can be used to study complex biological systems like membrane proteins.61

1.5 Site-Directed Spin Labeling The EPR spectroscopic technique was initially developed to study systems with inherent paramagnetic centers meaning the scope of this technique was limited to the studies involving organic radicals and metal complexes.62 With the advent of site-directed spin labeling (SDSL) technique, the use of EPR is now extended to study otherwise EPR

10 silent systems.55 Synthetic peptides and biological systems can now be engineered with cysteine residues, which allow the introduction of a paramagnetic center. SDSL involves removal of all the native cysteines and replacing them with the amino acids that cause no structural or functional perturbation of the system under investigation. A unique cysteine is then introduced using SPPS in the case of small peptides or site-directed mutagenesis in the case of recombinant protein expression. This cysteine can be reacted with a sulfhydryl specific nitroxide reagent to form a di-sulfide linkage, resulting in a stable paramagnetic side chain within the peptide/protein. Figure 1.6 shows the most commonly used nitroxide spin label S-(2,2,5,5-tertamethyl-2,5-dihydro-1H-pyrrol-3-yl)methyl methanesulfonothioate (MTSL) and the site directed spin labeling reaction.12

Figure 1.6 MTSL structure and the bond formation between MTSL nitroxide spin label and the cysteine sidechain during site directed spin labeling.

1.6 Continuous Wave (CW) EPR Continuous Wave EPR (CW-EPR) is the most commonly used EPR technique. In a CW-EPR experiment, constant microwave power is applied to the sample, while the magnetic field (Bo) is swept. Once the resonance condition is reached, the first derivative of the absorption spectra is recorded. In the case of MTSL, the unpaired electron interacts with the nearby nucleus called the hyperfine interaction. This causes a small change in the allowed energy levels depending on the spin state of the nucleus. Following the multiplicity rule of 2nI+1, the resulting EPR spectrum can have multiple lines. Where, n is the number of magnetically-equivalent nuclei, and I represents the nuclear spin. In the case of MTSL, the unpaired electron interacts primarily with the 14N nucleus (n = 1, I= 1). The characteristic three-line spectrum for MTSL is shown in the Figure 1.7.

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Figure 1.7 Energy diagram of MTSL showing the hyperfine interaction of the unpaired electron with the 14N nucleus of the nitroxide label resulting in the three resonant EPR peaks.

When free in the solution, a nitroxide spin label gives rise to three sharp peaks. If the motion is restricted to its rigid limit, full, orientation dependent parameters can be observed. The changes in the mobility of the spin label can provide useful information about the interaction between the peptide and the membrane, the topology and dynamics of membrane proteins, insertion angles of transmembrane peptides, helical tilt angle etc.

1.7 Pulsed EPR Spectroscopic Techniques Measuring distances between the amino-acid residues in the case of peptides and proteins can provide valuable information about the structure-function and conformational dynamics.63-66 Pulse-EPR experiments involves a series of short, microwave pulses to perturb the system containing the spin label.67 Advances in pulsed-EPR spectroscopy have provided a major breakthrough in the study of membrane protein systems. One of the most useful pulsed-EPR techniques is electron spin echo envelope modulation (ESEEM) spectroscopy. Also referred to as EPR-detected NMR, ESEEM can

12 provide useful information about the nuclei near the paramagnetic electron center.68 This method can detect distances of up to ~8 Å between the unpaired electron spin and a nearby NMR active nuclei.57 The technique has been widely used to study accessibility in the membranes as well as protein incorporation into lipid bilayers.69-72 Recently, it has also found application in membrane protein secondary structure determination.72-74 Double electron-electron resonance (DEER) spectroscopy is another pulse EPR technique that can measure distances between two spin labels falling within 20 Å to 80 Å of each other.75-77 The results from DEER spectroscopy yield distance probability distributions that can elucidate structure and conformational dynamics of proteins and be utilized as restraints for molecular dynamic simulations.63, 77-80 Compiling multiple distance distributions can help to refine structural models, intermolecular interactions, as well as conformational changes of active or inactive proteins.

1.8 Solid-State Nuclear Magnetic Resonance Spectroscopy Solid-state NMR spectroscopy is a popular biophysical technique used to study the structural and dynamic properties of the membranes and the peptides and/ proteins incorporated into the lipid bilayers.81, 82 Due to a 100% natural abundance of 31P in the phospholipid headgroups of the bilayer, it can be used to study the perturbations in the polar head group region of the bilayer using 31P solid-state NMR spectroscopy.83 Chemical shift anisotropy (CSA) and proton-phosphorous heteronuclear dipolar couplings are predominant interactions in static 31P SS-NMR. In the case of 31P SS-NMR, high power proton-phosphorus decoupling is used to eliminate the effects of proton- phosphorous heteronuclear dipolar couplings. The CSA linewidth measurements can help to provide valuable information on the dynamics of the 31P head groups.84, 85 Reduction in the CSA linewidth is observed due to increased surface fluidity or increase in the dynamics of the lipid bilayer upon interaction of the peptides/ proteins with the head group region of the membrane.86, 87 The hydrophobic acyl chains face away from the aqueous environment and reside in the hydrophobic core of the lipid bilayer. The perturbations in the acyl chains brought

13 about by peptide incorporation can be probed by 2H NMR spectroscopy. To probe these changes lipids with deuterated side chains are used as shown in Figure 1.8.

Figure 1.8 Structure of d54-DMPC lipid with deuterated acyl chains.

The width of the spectral quadrupolar splitting in the case of 2H NMR gives rise to specific order parameters (SCD) of various CD2 and CD3 groups throughout the lipid acyl chain 88, 89 length. The measurement of SCD order parameters can reflect the region of local disorder, that can then be used to study the effect of peptide/protein incorporation on the overall membrane dynamics.

14

References:

1. Cournia, Z.; Allen, T. W.; Andricioaei, I.; Antonny, B.; Baum, D.; Brannigan, G.; Buchete, N. V.; Deckman, J. T.; Delemotte, L.; del Val, C.; Friedman, R.; Gkeka, P.; Hege, H. C.; Henin, J.; Kasimova, M. A.; Kolocouris, A.; Klein, M. L.; Khalid, S.; Lemieux, M. J.; Lindow, N.; Roy, M.; Selent, J.; Tarek, M.; Tofoleanu, F.; Vanni, S.; Urban, S.; Wales, D. J.; Smith, J. C.; Bondar, A. N., Membrane Protein Structure, Function, and Dynamics: a Perspective from Experiments and Theory. Journal of Membrane Biology 2015, 248 (4), 611-640. 2. Bordag, N.; Keller, S., alpha-Helical transmembrane peptides: A "Divide and Conquer" approach to membrane proteins. Chemistry and Physics of Lipids 2010, 163 (1), 1-26. 3. Moraes, I.; Evans, G.; Sanchez-Weatherby, J.; Newstead, S.; Stewart, P. D. S., Membrane protein structure determination The next generation. Biochimica Et Biophysica Acta-Biomembranes 2014, 1838 (1), 78-87. 4. Yu, X. T.; Lorigan, G. A., Secondary Structure, Backbone Dynamics, and Structural Topology of Phospholamban and Its Phosphorylated and Arg9Cys-Mutated Forms in Phospholipid Bilayers Utilizing C-13 and N-15 Solid-State NMR Spectroscopy. Journal of Physical Chemistry B 2014, 118 (8), 2124-2133. 5. Sachs, J. N.; Engelman, D. M., Introduction to the membrane protein reviews: The interplay of structure, dynamics, and environment in membrane protein function. Annual Review of Biochemistry 2006, 75, 707-712. 6. Bull, S. C.; Doig, A. J., Properties of Protein Drug Target Classes. Plos One 2015, 10 (3). 7. Bakheet, T. M.; Doig, A. J., Properties and identification of human protein drug targets. Bioinformatics 2009, 25 (4), 451-457. 8. Lieberman, R. L.; Culver, J. A.; Entzminger, K. C.; Pai, J. C.; Maynard, J. A., Crystallization chaperone strategies for membrane proteins. Methods 2011, 55 (4), 293- 302. 9. Overington, J. P.; Al-Lazikani, B.; Hopkins, A. L., How many drug targets are there? Nature Reviews Drug Discovery: 2006; Vol. 5, pp 993 – 996. 10. Elofsson, A.; von Heijne, G., Membrane protein structure: Prediction versus reality. Annual Review of Biochemistry 2007, 76, 125-140. 11. Garman, E. F., Developments in X-ray Crystallographic Structure Determination of Biological Macromolecules. Science 2014, 343 (6175), 1102-1108. 12. Sanders, C. R.; Nagy, J. K., Misfolding of membrane proteins in health and disease: the lady or the tiger? Current Opinion in 2000, 10 (4), 438- 442. 13. Tan, J. M. M.; Wong, E. S. P.; Lim, K. L., Protein Misfolding and Aggregation in Parkinson's Disease. Antioxidants & Redox Signaling 2009, 11 (9), 2119-2134. 14. Hartl, F. U., Protein Misfolding Diseases. Annual Review of Biochemistry, Vol 86 2017, 86, 21-26. 15. White, S. H., The progress of membrane protein structure determination. Protein Science 2004, 13 (7), 1948-1949. 16. Das, B. B.; Park, S. H.; Opella, S. J., Membrane protein structure from rotational diffusion. Biochimica Et Biophysica Acta-Biomembranes 2015, 1848 (1), 229-245.

15

17. Deisenhofer, J.; Epp, O.; Miki, K.; Huber, R.; Michel, H., Structure Of The Protein Subunits In The Photosynthetic Reaction Center Of Rhodopseudomonas-Viridis At 3a Resolution. Nature 1985, 318 (6047), 618-624. 18. Berman, H. M.; Westbrook, J.; Feng, Z.; Gilliland, G.; Bhat, T. N.; Weissig, H.; Shindyalov, I. N.; Bourne, P. E., The Protein Data Bank. Nucleic Acids Research 2000, 28 (1), 235-242. 19. White, S. Membrane Proteins of Known 3D Structure. https://blanco.biomol.uci.edu/mpstruc/. 20. Midgett, C. R.; Madden, D. R., Breaking the bottleneck: Eukaryotic membrane protein expression for high-resolution structural studies. Journal of Structural Biology 2007, 160 (3), 265-274. 21. Bahar, I.; Lezon, T. R.; Bakan, A.; Shrivastava, I. H., Normal Mode Analysis of Biomolecular Structures: Functional Mechanisms of Membrane Proteins. Chemical Reviews 2010, 110 (3), 1463-1497. 22. Baker, M., Making membrane proteins for structures: a trillion tiny tweaks. Nature Methods 2010, 7 (6), 429-434. 23. Walsh, C. T.; Garneau-Tsodikova, S.; Gatto, G. J., Protein posttranslational modifications: The chemistry of proteome diversifications. Angewandte Chemie- International Edition 2005, 44 (45), 7342-7372. 24. Leyva, J. A.; Bianchet, M. A.; Amzel, L. M., Understanding ATP synthesis: structure and mechanism of the F1-ATPase (Review). Molecular Membrane Biology 2003, 20 (1), 27-33. 25. Henry, G. D.; Sykes, B. D., Methods To Study Membrane-Protein Structure In Solution. Nuclear Magnetic Resonance, Pt C 1994, 239, 515-535. 26. Harayama, T.; Riezman, H., Understanding the diversity of membrane lipid composition. Nature Reviews Molecular Cell Biology 2018, 19 (5), 281-296. 27. Young, R.; Wang, I., Phage Lysis. The Bacteriophages. 2nd Ed ed.; Oxford Univ Press: Oxford, 2006. 28. Young, R.; Wang, I. N.; Roof, W. D., Phages will out: strategies of host cell lysis. Trends in Microbiology 2000, 8 (3), 120-128. 29. Young, R., Phage Lysis: Three Steps, Three Choices, One Outcome. Journal of Microbiology 2014, 52 (3), 243-258. 30. Young, R., Bacteriophage holins: Deadly diversity. Journal of Molecular Microbiology and Biotechnology 2002, 4 (1), 21-36. 31. Young, R., Phage Lysis: Do we have the hole story yet? Curr Opin Microbiol. 2013, 16 (6), 1-8. 32. Dewey, J. S.; Savva, C. G.; White, R. L.; Vitha, S.; Holzenburg, A.; Young, R., Micron-scale holes terminate the phage infection cycle. Proceedings of the National Academy of Sciences of the United States of America 2010, 107 (5), 2219-2223. 33. Garrett, J.; Young, R., Lethal action of bacteriophage lambda S gene. Journal of Virology: 1982; Vol. 44, pp 886-892. 34. Adhya, S.; Sen, A.; Mitra, S., The role of gene S In: The Bacteriophage Lambda. Cold Spring Harbor Laboratory: Cold Spring Harbor, NY, 1971; p 743-746. 35. To, K. H.; Young, R., Probing the Structure of the S105 Hole. Journal of Bacteriology 2014, 196 (21), 3683-3689.

16

36. 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. 37. Park, T.; Struck, D. K.; Dankenbring, C. A.; Young, R., The pinholin of lambdoid phage 21: Control of lysis by membrane depolarization. Journal of Bacteriology 2007, 189 (24), 9135-9139. 38. 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. 39. Bonovich, M. T.; Young, R., Dual Start Motif In 2 Lambdoid S-Genes Unrelated To Lambda-S. Journal of Bacteriology 1991, 173 (9), 2897-2905. 40. 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. 41. 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. 42. 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. 43. Pang, T.; Park, T.; Young, R., Mutational analysis of the S21 pinholin. Molecular Microbiology 2010, 76 (1), 68-77. 44. Park, T.; Struck, D. K.; Deaton, J. F.; Young, R., Topological dynamics of holins in programmed bacterial lysis. Proceedings of the National Academy of Sciences of the United States of America 2006, 103 (52), 19713-19718. 45. Pang, T.; Park, T.; Young, R., Mapping the pinhole formation pathway of S21. Molecular Microbiology 2010, 78 (3), 710-719. 46. Merrifield, R. B., Solid Phase Peptide Synthesis .1. Synthesis Of A Tetrapeptide. Journal of the American Chemical Society 1963, 85 (14), 2149-&. 47. Hansen, P. R.; Oddo, A., Fmoc Solid-Phase Peptide Synthesis. Peptide Antibodies: Methods and Protocols 2015, 1348, 33-50. 48. Bacsa, B.; Desai, B.; Dibo, G.; Kappe, C. O., Rapid solid-phase peptide synthesis using thermal and controlled microwave irradiation. Journal of Peptide Science 2006, 12 (10), 633-638. 49. Duro-Castano, A.; Conejos-Sanchez, I.; Vicent, M. J., Peptide-Based Polymer Therapeutics. Polymers 2014, 6 (2), 515-551. 50. Palomo, J. M., Solid-phase peptide synthesis: an overview focused on the preparation of biologically relevant peptides. Rsc Advances 2014, 4 (62), 32658-32672. 51. Stawikowski, M.; Fields, G. B., Introduction to Peptide Synthesis. Curr Protoc Protein Sci: 2002; Vol. 18, pp 1-17. 52. Lloyd-Williams, P.; Albericio, F.; Giralt, E., Chemical Approaches to the Synthesis of Peptides and Proteins. CRC Press: 1997; p 304. 53. King, D. S.; Fields, C. G.; Fields, G. B., A Cleavage Method Which Minimizes Side Reactions Following Fmoc Solid-Phase Peptide-Synthesis. International Journal of Peptide and Protein Research 1990, 36 (3), 255-266.

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54. Albericio, F.; Kneibcordonier, N.; Biancalana, S.; Gera, L.; Masada, R. I.; Hudson, D.; Barany, G., Preparation And Application Of The 5-(4-(9- Fluorenylmethyloxycarbonyl)Aminomethyl-3,5-Dimethoxyphenoxy)Vale Ric Acid (Pal) Handle For The Solid-Phase Synthesis Of C-Terminal Peptide Amides Under Mild Conditions. Journal of Organic Chemistry 1990, 55 (12), 3730-3743. 55. Cornish, V. W.; Benson, D. R.; Altenbach, C. A.; Hideg, K.; Hubbell, W. L.; Schultz, P. G., Site-Specific Incorporation Of Biophysical Probes Into Proteins. Proceedings of the National Academy of Sciences of the United States of America 1994, 91 (8), 2910-2914. 56. Sahu, I. D.; Lorigan, G. A., Site-Directed Spin Labeling EPR for Studying Membrane Proteins. Biomed Research International 2018. 57. Sahu, I. D.; McCarrick, R. M.; Lorigan, G. A., Use of Electron Paramagnetic Resonance To Solve Biochemical Problems. Biochemistry 2013, 52 (35), 5967-5984. 58. 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. 59. Stone, T. J.; Buckman, T.; Nordio, P. L.; McConnell, H. M., SPIN-LABELED BIOMOLECULES. Proceedings of the National Academy of Sciences of the United States of America 1965, 54 (4), 1010-+. 60. Weil, J. A.; Bolton, J. R., Electron paramagnetic resonance: Elementary Theory and Practical Applications. Wiley-Interscience: Hoboken, NJ, USA, 2007. 61. Klare, J. P.; Steinhoff, H. J., Spin labeling EPR. Photosynthesis Research 2009, 102 (2-3), 377-390. 62. Leman, J. K.; Ulmschneider, M. B.; Gray, J. J., Computational modeling of membrane proteins. Proteins-Structure Function and Bioinformatics 2015, 83 (1), 1-24. 63. Sahu, I. D.; Kroncke, B. M.; Zhang, R. F.; Dunagan, M. M.; Smith, H. J.; Craig, A.; McCarrick, R. M.; Sanders, C. R.; Lorigan, G. A., Structural Investigation of the Transmembrane Domain of KCNE1 in Proteoliposomes. Biochemistry 2014, 53 (40), 6392-6401. 64. Aitha, M.; Moritz, L.; Sahu, I. D.; Sanyurah, O.; Roche, Z.; McCarrick, R.; Lorigan, G. A.; Bennett, B.; Crowder, M. W., Conformational dynamics of metallo-beta- lactamase CcrA during catalysis investigated by using DEER spectroscopy. Journal of Biological 2015, 20 (3), 585-594. 65. Glaenzer, J.; Peter, M. F.; Hagelueken, G., Studying structure and function of membrane proteins with PELDOR/DEER spectroscopy - The crystallographers' perspective. Methods 2018, 147, 163-175. 66. Vicente, E. F.; Sahu, I. D.; Costa, A. J.; Cilli, E. M.; Lorigan, G. A., Conformational changes of the HsDHODH N-terminal Microdomain via DEER Spectroscopy. Journal of Physical Chemistry B 2015, 119 (28), 8693-8697. 67. Prisner, T.; Rohrer, M.; MacMillan, F., Pulsed EPR spectroscopy: Biological applications. Annual Review of Physical Chemistry 2001, 52, 279-313. 68. Stoll, S.; Calle, C.; Mitrikas, G.; Schweiger, A., Peak suppression in ESEEM spectra of multinuclear spin systems. Journal of Magnetic Resonance 2005, 177 (1), 93- 101.

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69. Bartucci, R.; Guzzi, R.; Esmann, M.; Marsh, D., Water Penetration Profile at the Protein-Lipid Interface in Na,K-ATPase Membranes. Biophysical Journal 2014, 107 (6), 1375-1382. 70. Carmieli, R.; Papo, N.; Zimmermann, H.; Potapov, A.; Shai, Y.; Goldfarb, D., Utilizing ESEEM spectroscopy to locate the position of specific regions of membrane- active peptides within model membranes. Biophysical Journal 2006, 90 (2), 492-505. 71. Milov, A. D.; Samoilova, R. I.; Tsvetkov, Y. D.; De Zotti, M.; Formaggio, F.; Toniolo, C.; Handgraaf, J. W.; Raap, J., Structure of Self-Aggregated Alamethicin in ePC Membranes Detected by Pulsed Electron-Electron Double Resonance and Electron Spin Echo Envelope Modulation . Biophysical Journal 2009, 96 (8), 3197-3209. 72. Zhou, A. D.; Abu-Baker, S.; Sahu, I. D.; Liu, L. S.; McCarrick, R. M.; Dabney- Smith, C.; Lorigan, G. A., Determining alpha-Helical and beta-Sheet Secondary Structures via Pulsed Electron Spin Resonance Spectroscopy. Biochemistry 2012, 51 (38), 7417-7419. 73. Liu, L. S.; Lorigan, G., Probing the Secondary Structure of Membrane Proteins with the Pulsed EPR ESEEM Technique. Biophysical Journal 2014, 106 (2), 192A-192A. 74. Liu, L. S.; Sahu, I. D.; Bottorf, L.; McCarrick, R. M.; Lorigan, G. A., Investigating the Secondary Structure of Membrane Peptides Utilizing Multiple H-2-Labeled Hydrophobic Amino Acids via Electron Spin Echo Envelope Modulation (ESEEM) Spectroscopy. Journal of Physical Chemistry B 2018, 122 (16), 4388-4396. 75. Baber, J. L.; Louis, J. M.; Clore, G. M., Dependence of Distance Distributions Derived from Double Electron-Electron Resonance Pulsed EPR Spectroscopy on Pulse- Sequence Time. Angewandte Chemie-International Edition 2015, 54 (18), 5336-5339. 76. Jeschke, G.; Polyhach, Y., Distance measurements on spin-labelled biomacromolecules by pulsed electron paramagnetic resonance. Physical Chemistry Chemical Physics 2007, 9 (16), 1895-1910. 77. 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. 78. Li, Q. F.; Wanderling, S.; Sompornpisut, P.; Perozo, E., Structural basis of lipid- driven conformational transitions in the KvAP voltage-sensing domain. Nature Structural & Molecular Biology 2014, 21 (2), 160-+. 79. Jao, C. C.; Hegde, B. G.; Chen, J.; Haworth, I. S.; Langen, R., Structure of membrane-bound alpha-synuclein from site-directed spin labeling and computational refinement. Proceedings of the National Academy of Sciences of the United States of America 2008, 105 (50), 19666-19671. 80. Jeschke, G.; Abbott, R. J. M.; Lea, S. M.; Timmel, C. R.; Banham, J. E., The characterization of weak protein-protein interactions: Evidence from DEER for the trimerization of a von Willebrand factor A domain in solution. Angewandte Chemie- International Edition 2006, 45 (7), 1058-1061. 81. Kinnun, J. J.; Leftin, A.; Brown, M. F., Solid-State NMR Spectroscopy for the Physical Chemistry Laboratory. Journal of Chemical Education 2013, 90 (1), 123-128. 82. Molugu, T. R.; Lee, S.; Brown, M. F., Concepts and Methods of Solid-State NMR Spectroscopy Applied to Biomembranes. Chemical Reviews 2017, 117 (19), 12087- 12132.

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83. Santos, J. S.; Lee, D. K.; Ramamoorthy, A., Effects of antidepressants on the conformation of phospholipid headgroups studied by solid-state NMR. Magnetic Resonance in Chemistry 2004, 42 (2), 105-114. 84. Dave, P. C.; Tiburu, E. K.; Damodaran, K.; Lorigan, G. A., Investigating structural changes in the lipid bilayer upon insertion of the transmembrane domain of the membrane-bound protein phospholamban utilizing P-31 and H-2 solid-state NMR spectroscopy. Biophysical Journal 2004, 86 (3), 1564-1573. 85. Abu-Baker, S.; Lorigan, G. A., Phospholamban and its phosphorylated form interact differently with lipid bilayers: A (31)P, (2)H, and (13)C solid-state NMR spectroscopic study. Biochemistry 2006, 45 (44), 13312-13322. 86. Seelig, J., P-31 Nuclear Magnetic-Resonance And Head Group Structure Of Phospholipids In Membranes. Biochimica Et Biophysica Acta 1978, 515 (2), 105-140. 87. McLaughlin, A. C.; Cullis, P. R.; Berden, J. A.; Richards, R. E., P-31 Nmr Of Phospholipid Membranes - Effects Of Chemical-Shift Anisotropy At High Magnetic-Field Strengths. Journal of Magnetic Resonance 1975, 20 (1), 146-165. 88. Koenig, B. W.; Ferretti, J. A.; Gawrisch, K., Site-specific deuterium order parameters and membrane-bound behavior of a peptide fragment from the intracellular domain of HIV-1 gp41. Biochemistry 1999, 38 (19), 6327-6334. 89. Yamaguchi, S.; Huster, D.; Waring, A.; Lehrer, R. I.; Kearney, W.; Tack, B. F.; Hong, M., Orientation and dynamics of an antimicrobial peptide in the lipid bilayer by solid- state NMR spectroscopy. Biophysical Journal 2001, 81 (4), 2203-2214.

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Chapter 2

Solid Phase Synthesis and Spectroscopic Characterization of the Active and Inactive Forms of Bacteriophage S21 Pinholin Protein.

Daniel L. Drew Jr., Tanbir Ahammad, Rachel A. Serafin, Brandon J. Butcher, Katherine R. Clowes, Zachary Drake, Indra D. Sahu, Robert M. McCarrick, and Gary A. Lorigan*

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

This work has been published in the Journal of Analytical Biochemistry:

Drew, D. L. et al. Solid phase synthesis and spectroscopic characterization of the active and inactive forms of bacteriophage S21 pinholin protein. Analytical Biochemistry 2019, 567, 14-20

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2.1 Abstract: The mechanism for the lysis pathway of double-stranded DNA bacteriophages involves a small hole-forming class of membrane proteins, the holins. This study focuses on a poorly characterized class of holins, the pinholin, of which the S21 protein of phage ϕ21 is the prototype. Here we report the first in vitro synthesis of the wildtype form of the S21 pinholin, S2168, and negative-dominant mutant form, S21IRS, both prepared using solid phase peptide synthesis and studied using biophysical techniques. Both forms of the pinholin were labeled with a nitroxide spin label and successfully incorporated into both bicelles and multilamellar vesicles which are membrane mimetic systems. Circular dichroism revealed the two forms were both >80% alpha helical, in agreement with the predictions based on the literature. The molar ellipticity ratio [θ]222/

[θ]208 for both forms of the pinholin was 1.4, suggesting a coiled-coil tertiary structure in the bilayer consistent with the proposed oligomerization step in models for the mechanism of hole formation. 31P solid-state NMR spectroscopic data on pinholin indicate a strong interaction of both forms of the pinholin with the membrane headgroups. The 31P NMR data has an axially symmetric line shape which is consistent with lamellar phase proteoliposomes lipid mimetics.

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2.2 Introduction: The final step of the double-stranded DNA bacteriophage infection cycle is host lysis.[1-3] The mechanism for this lysis pathway involves three proteins, a small hole- forming inner membrane protein known as the holin, a muralytic enzyme known as the endolysin, and the spanin complex responsible for outer membrane disruption.[3, 4] The function of the holin protein is to permeabilize the inner phospholipid bilayer allowing the release of the endolysin to begin the degradation of the peptidoglycan.[5] This is accomplished by a harmless accumulation of the holin in the host cell membrane until the protein “triggers” at an allele-specific time. 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.[3] Due to the variation in mechanisms and sizes of lesions formed between different classes of holins the lesions have been termed “holes” to show distinction from channels and other such membrane permeabilization pathways.[2, 3] Initially it was believed that all holins, like the λ S105 canonical holin, trigger to form micron-scale holes in the inner cell membrane.[3] These holes allowed for non- selective escape of fully folded and functional endolysin enzymes. However, more recently a second type of holin has been discovered.[2, 6] Instead of forming micron-scale non-selective holes in the cytoplasmic membrane, these holins form nanometer-scale holes that are only responsible for the depolarization of the membrane.[2, 6] Due to the small size of the holes, this new class of holin was named the pinholin. Unlike the large hole forming canonical holins the hole created by the pinholins is not large enough to allow for the nonspecific escape of functional endolysin from the cytoplasm. Instead these pinholins are paired with signal-anchor-release (SAR) endolysins.[2, 3] These proteins are named for a special N-terminal transmembrane helix that acts as an uncleaved signal sequence, resulting in a sec-mediated export. The SAR-endolysins accumulate in the periplasm as membrane-tethered, inactive enzymes. When the pinholins trigger and cause depolarization of the membrane, the SAR domain exits the bilayer, allowing the periplasmic catalytic domain to refold to its active form and begin degradation of the peptidoglycan.[6-8] This study focuses on the optimization of the solid phase peptide synthesis and spectroscopic characterization of the pinholin membrane protein system. More

23 specifically, the system under study is encoded by the S21 holin gene of the lambdoid bacteriophage ϕ21. The 71-codon S21 gene has a dual translational start motif.[9, 10] This results in the synthesis of two gene products, S2171 and S2168, resulting from translational initiations from codon Met1 and Met4, respectively. These two gene products are outlined in Figure 2.1. The S2168 is the functional or active form of the pinholin. This form of pinholin has two transmembrane domains (TMDs), the first of which (TMD1) externalizes from the membrane, while the second (TMD2) remains embedded in the bilayer and is essential for lytic functionality.[11-13] The second form, S2171, is known as the antiholin. The antiholin is responsible for the delayed triggering of the pinholin due to the addition of a positively charged lysine on the N-terminus (Figure 2.1) which drastically slows the externalization of TMD1. A basic schematic for the orientation of the pinholin in the membrane as well as hypothetical conformations of externalized TMD1 can be seen in Figure 2.2. The pinholin pathway begins with the accumulation of both forms of pinholin embedded in the bilayer, accumulating as dimers. Since the S2168/S2171 production ratio is ~2:1, the dimers are mostly S2168:S2168 homodimers and S2168:S2171 heterodimers.[14] Activation is thought to require externalization of TMD1 from the bilayer. Once both TMD1 segments are externalized, the pinholin dimer is licensed to continue in a pathway of oligomerization. When concentration of the activated pinholin dimers reach a critical concentration, the pinholin population triggers and causes massive and sudden depolarization of the inner cell membrane.[6, 15] The structure, mechanism, and model pathway of the pinholin protein has been difficult to study primarily because pinholin is not only a hydrophobic membrane protein but also expresses lethal in function, thus prohibiting high-level biosynthesis. The pinholin system poses an interesting challenge as the length of the pinholin and antiholin proteins are near the limit of solid phase peptide synthesis. The resulting function of the pinholin pathway is well known, but the individual steps of the dimerization and oligomerization in the pathway are not well studied. The Young group has shown that the addition of the five amino acid sequence ‘RYIRS’ to the N-terminus of the S2168 pinholin, shown in Figure 2.1, prevents the externalization of TMD1 and ultimately the function of the pinholin pathway.[12] This is called the inactive IRS form of the pinholin. This presents an opportunity for recapitulating the holin pathway in vitro, using appropriate proportions of

24 the wildtype pinholin (S2168) and the S21IRS, as a surrogate antipinholin to control the pathway. Utilizing this control, the structure and dynamics of the pinholin protein as it progresses through the lytic pathway can be spectroscopically studied with a variety of biophysical techniques. This study represents the first time that solid phase peptide synthesis has been used to study any holin system in vitro. Circular dichroism has been used to determine alpha helical protein secondary structure as well as probe the initial oligomerization steps required in the proposed pinholin lytic pathway. The extant biophysical information on the oligomerization of pinholin was conducted on a truncated version of the protein where only TMD2 was present.[11, 14] However, this study uses the full length pinholin system. Electron paramagnetic resonance (EPR) spectroscopy was used to confirm pinholin incorporation into an in vitro multilamellar vesicle mimetic system. In conjunction with EPR spectroscopy, 31P solid-state NMR spectroscopy was used to discern the effect the different pinholin forms have on the membrane from the perspective of the phospholipids.

2.3 Material and Methods:

2.3.1 Solid Phase Peptide Synthesis The solid phase peptide synthesis of pinholin peptides was conducted using a CEM Liberty Blue Peptide Synthesizer with Discovery Bio Microwave System. The synthesis used a NovaSyn TG amino resin, a composite of low cross-linked polystyrene with the PEG chains terminally functionalized with an amino group. All syntheses were run at a 0.1 mM scale with Dimethylformamide (DMF) as the base solvent. All Fmoc protected amino acid solutions were prepared at a 0.2 M concentration and coupled using a standard activator and activator base pair of DIC and oxyma, respectively. The coupling reactions were run at 90°C for 4 min while the Fmoc deprotection was run with 20% v/v piperidine in DMF at 93°C for 1 min.[16] As seen in Figure 2.1, the pinholin protein is naturally cys-less, requiring the introduction of only one site-specific cysteine into the primary sequence for site directed spin labeling EPR experiments.

25

2.3.2 Protecting Group and Solid Phase Cleavage The solid phase bound pinholin peptide was washed three times with and allowed to dry on a vacuum filter. The amino acid side chain protecting groups as well as the solid phase resin were cleaved from the peptide in a three-hour Trifluoroacetic acid (TFA) cleavage reaction.[17-19] The cleavage solution was then gravity filtered to remove the cleaved solid phase resin. TFA was evaporated from the reaction solution using inert nitrogen gas flow. Tert-butyl ether was added in excess to precipitate the pinholin peptide and separate from the still solubilized protecting groups.[16] The precipitated peptide was centrifuged down into a pellet by spinning for 15 min at 9000 rpm and the excess ether was decanted off. This procedure was repeated three times to ensure that all the protecting groups and scavengers were removed. Following the three ether washes the pinholin peptide was placed in a vacuum desiccator to dry for at least 8 hrs.

2.3.3 Protein Purification and Spin Labeling The crude pinholin peptide was purified by reverse phase high pressure liquid chromatography (RP-HPLC) using a C4 column running a two-solvent gradient. The first solvent was deionized water, the second was 90% HPLC grade . Both were degassed and then acidified with 0.1% TFA by volume. The pinholin peptide was collected in fractions and the molecular weight of the peptide was confirmed using Matrix Assisted Laser Desorption Ionization – Time of Flight Spectrometry (MALDI-TOF). Collected fractions were dried using lyophilization to recover the pure pinholin. The dry, pure peptide was then spin-labeled with S-(1-oxyl-2,2,5,5-tetramethyl-2,5- dihydro-1H-pyrrol-3-yl)methyl methanesulfonothioate (MTSL), a nitroxide spin label at position Leu25. This position was chosen due to its location in TMD1, and therefore will test our ability to spin label the different conformations adopted by the active S2168 and inactive S21IRS pinholin. This was performed by dissolving the pinholin peptide and MTSL, at a 5x molar excess, in DMSO and letting it react while being stirred continuously for 24 hrs. The reaction was stopped by freezing the solution in liquid nitrogen and then dried using lyophilization.

26

The resulting crude spin-labeled pinholin was again purified using RP-HPLC and the same two-solvent system on a C4 semi-prep column to remove the excess MTSL. MTSL addition to the pinholin peptide was confirmed through MALDI-TOF mass, as shown in Figure 2.3. Collected pure peptide fractions were dried using lyophilization.

2.3.4 Peptide Incorporation into Lipid Mimetic Systems The pure peptide was incorporated into one of two different lipid mimetic environments, 1,2-Dimyristoyl - sn - Glycero - 3 - Phosphocholine (DMPC) / 1,2 - Diheptanoyl - sn - Glycero – 3 - Phosphocholine (DHPC) bicelles, or DMPC multilamellar vesicles (MLV). MLVs are a commonly used mimetic system and have been shown to be successful in mimicking a bilayer for membrane protein studies.[20, 21] MLVs were created by mixing the peptide dissolved in 2,2,2-Trifluoroethanol (TFE) with DMPC in at the desired protein concentration or protein to lipid ratio. Solvents were evaporated off using inert N2 gas and the remaining lipid/protein film was rehydrated using 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer at a concentration of 20mM adjusted to a neutral pH of ~7.0. The protein/lipid solution was flash frozen in liquid nitrogen and then sonicated. This process was repeated 3 times to ensure MLV formation. Bicelles were made by taking the dissolved peptide in TFE and adding it to a solution of DMPC and DHPC in chloroform at an optimized q value of 3.6.[22] Then the solvents were evaporated off using inert N2(g) and the same process as MLV formation was followed until sample became clear.

2.3.5 Circular Dichroism Circular Dichroism (CD) measurements on pinholin were performed on an Aviv Circular Dichroism Model 435 in a quartz with a 1.0 mm path length. Data was collected from 260 to 190 nm with 1 nm bandwidth at 25°C. CD data was collected on pinholin MLV samples prepared using the conditions outlined in the previous section. CD spectral simulations and secondary structural content calculations were performed using DICHROWEB software found on http://dichroweb.cryst.bbk.ac.uk.[23] The CDSSTR algorithm was used for all simulations and compared back to reference

27 data set SMP180 with a spectral width of 190-240nm.[24-28] Molar ellipticity ratios [θ]222/

[θ]208 were calculated to determine the presence of tertiary coiled-coil helices.[29, 30]

2.3.6 Continuous Wave Electron Paramagnetic Resonance Spectroscopy CW-EPR experiments were performed at the Ohio Advanced EPR Laboratory at Miami University. CW-EPR spectra were collected at X-band on a Bruker EMX EPR spectrometer using an ER041xG microwave bridge and ER4119-HS cavity coupled with a BVT 3000 nitrogen gas temperature controller. Each CW-EPR spectrum was acquired with 42 s field sweep with a central field of 3315 G and sweep width of 100 G, modulation frequency of 100 kHz, modulation amplitude of 1 G, and microwave power of 10 mW at room temperature.

2.3.7 31P Solid State Nuclear Magnetic Resonance Spectroscopy The 31P solid-state nuclear magnetic resonance measurements were conducted at 25°C using a Bruker 500 MHz WB UltraShield NMR spectrometer with a 4mm triple resonance CP-MAS probe. 31P NMR spectra were recorded with 1H decoupling using a 4 µs π/2 pulse and a 4 s recycle delay, a spectral width of 300 ppm, and by averaging 4K scans. The free induction decay was processed using 200 Hz of line broadening. All figures were generated using the Igor software package.

2.4 Results and Discussion: This study reports the successful in vitro synthesis of both the active S2168 and inactive S21IRS forms of the pinholin system using solid phase peptide synthesis (SPPS). The full length active 68 and inactive IRS forms of the pinholin protein are composed of 68 and 73 amino acids respectively. The optimization of the sample preparation was confirmed through MALDI-TOF spectra that match the predicted molecular weights for each of the wild type pinholins, 7548 Da for the active S2168 and 8223 Da for the inactive S21IRS (Figure 2.1 and 2.3). A cysteine was substituted into the primary sequence for the leucine at position 25 for future nitroxide spin labeling. After synthesis, the solid phase resin and all amino acid protecting groups were removed in a three-hour reaction using a 30 mL cleavage solution of trifluoroacetic acid (TFA). The resulting peptide was

28 precipitated from solution and washed 3 times using tert-butyl ether. Following the ether washes, the protein was purified using RP-HPLC and fractions were analyzed using MALDI-TOF to confirm the accuracy and overall purity of the synthesis.

2.4.1 Optimization of the Solid Phase Peptide Synthesis Long hydrophobic peptides are difficult to synthesize, therefore the initial pinholin synthesis was of the first 20 amino acids (see Figure 2.1) with each of the following syntheses adding 10 more amino acids to the chain length. The peptide from each synthesis was analyzed using MALDI-TOF to determine which amino acids were not coupling completely. Coupling times and temperatures where adjusted to increase successful coupling as the chain length was extended to the full 73 amino acids of the inactive IRS pinholin. Since the pinholin system is naturally cys-less the incorporation of a cysteine at any position along the primary sequence allows for disulfide bond formation to the spin label at specific positions. The MALDI-TOF results for the synthesis of both the active S2168 and inactive S21IRS pinholin wild type using the fourth cleavage condition in the following section can be seen in Figure 2.3. The target MW for the active S2168 and inactive S21IRS pinholin were 7546 and 8222 Da, respectively. The observed MW from Figure 2.3 for the active S2168 was found to be 7548 Da while the inactive S21IRS shows a peak at 8223 Da confirming a successful synthesis for both forms of the WT pinholin. Figure 2.3 also shows the MALDI-TOF data for the active S2168-L25C-MTSL and inactive S21IRS-L25C-MTSL forms of pinholin. The +185 Da shift in the m/z, 7722 for the active and 8391 for the inactive, confirms the successful spin labeling of both forms of the pinholin. In all cases there is a small peak appearing at one half the target MW corresponding to detection of a doubly charged pinholin ion.

2.4.2 Optimization of Peptide Cleavage Conditions and HPLC Purification The cleavage protocol was optimized by monitoring the cleavage reaction of various cleavage solutions over time. Four different cleavage conditions were tested based on type and number of certain amino acid side chain protecting groups present.[16] The cleavage conditions for four different solutions are as follows:

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Cleavage condition 1[17] – TFA/EDT/thioanisole/water – 88/5/2/5 Cleavage condition 2[18] – TFA/phenol/water/thioanisole/EDT – 82.5/5/5/5/2.5 Cleavage condition 3[19] – TFA/thioanisole/EDT/anisole – 90/5/3/2 Cleavage condition 4[17] – TFA/triisoproylsilane/EDT/water – 94/1/2.5/2.5 Each of the different components of the cleavage reaction was measured out as a %v/v with a final cleavage solution volume of 30 mL. As each cleavage condition was tested a 5 mL aliquot of the reaction solution was removed at t = 30, 60, 90, 120, 150, and 180 minutes. Each aliquot was worked up following the procedure outlined in section 2.2. Finally, each time point was analyzed using MALDI-TOF to confirm the target protein molecular weight. The MALDI-TOF results clearly identified cleavage condition 4 as the best cleavage condition for this system as it gave the sharpest target peak with the fewest impurity peaks. The MALDI peak at the target molecular weight does not begin to appear until an hour after the reaction begins. The MALDI results show no change in peak shape between the 2.5 and 3 hour aliquots indicating the completion of the cleavage reaction.

2.4.3 Optimization of MALDI-TOF Sample Conditions and Matrix To optimize the MALDI-TOF analysis the pinholin peptide was dissolved in solutions of increasing amounts of acetonitrile in water. Each solution was spotted using one of three different matrices at a 1:1 ratio, α-Cyano-4-hydroxycinnamic acid (CHCA) matrix, 2,5-Dihydroxybenzoic acid (DHB) matrix, and Sinapic Acid (SA) matrix. The sample condition of 85% acetonitrile spotted with sinapic acid matrix gave the clearest resolution of MS peaks. Closer analysis of MALDI-TOF results revealed peaks at a higher m/z ratio indicating the presence of unremoved protecting groups during cleavage step. This process lead to the optimization of the cleavage reaction to remove these protecting groups.

2.4.4 Circular Dichroism of Active S2168 and Inactive S21IRS Pinholin CD spectroscopy is primarily used in the biochemical field to determine the global secondary structure of large macromolecules, such as peptides and proteins. This is accomplished through measuring the difference in absorption between left and right-

30 handed circularly polarized light over a range of .[31] The folding of the protein into a helical secondary structure was confirmed for both the active S2168 and inactive S21IRS pinholin using CD spectroscopy. CD samples for both the active S2168 and inactive S21IRS forms of wild type pinholin were separately prepared in DMPC MLVs at a protein to lipid ratio of 1:500 with a final concentration of protein equal to 50 µM. Background at lower wavelengths was minimized by subtracting the data collected from empty DMPC MLVs run at the same lipid concentration from the protein sample in DMPC MLVs. The resulting CD spectra of the active S2168 pinholin (blue) and the inactive S21IRS (red) are shown in Figure 2.4. Both forms of pinholin show a predominately helical structure with double minima at 208 nm and 222 nm and a large positive peak at 195 nm. The active S2168 pinholin showed a calculated helical content of 83%, while the inactive S21IRS pinholin secondary structure calculation determined 82% helical content. The percent normalized RMSD value calculated for both forms of the pinholin were less than 1.0%.[28] The helical content agrees well to the expected secondary structure of the protein based on previous studies in the literature.[11] To give a more in-depth analysis of the CD spectra the molar ellipticity ratio of

[θ]222/ [θ]208 was calculated for both forms of the pinholin. A molar ellipticity ratio exceeding a value of 1 is indicative of coiled-coil helical structures.[29, 30, 32] The molar ellipticity ratio [θ]222/ [θ]208 for both forms of the pinholin was found to be 1.4 and therefore, this coiled-coil structure was determined to be present for both the active S2168 and inactive S21IRS forms. This matches with the current predicted pinholin lysis pathway in which the pinholin begin to oligomerize, forming more coiled-coils, as more pinholin proteins accumulate in the membrane ultimately resulting in the lysis of the membrane.[11, 15] Further evidence of this will be shown in future experiments in which the oligomerization state of pinholin will be investigated as a function of concentration.

2.4.5 Continuous Wave EPR Measurements of Pinholin CW-EPR spectroscopy can be used to probe the dynamic and structural properties of both solution and membrane proteins. CW-EPR spectroscopy of these spin-labeled

31 molecules can reveal information about the motion of the nitroxide side chain, solvent accessibility, and the polarity of the surrounding environment.[21, 33] The successful spin labeling of the pinholin protein and incorporation into both bicelle and MLV lipid mimetic systems is shown in the CW-EPR spectra in Figure 2.5. This EPR data was also used to calculate spin labeling efficiency for each peptide which ranged from ~85-90% labeling efficiency. The EPR spectra for unbound nitroxide spin labels typically consist of three sharp peaks of relatively similar intensities, as seen in Figure 2.5A. Upon binding to a protein, such as pinholin, the MTSL will have a more restricted mobility due to the presence of the protein backbone. This decrease in the motion of the MTSL will broaden the lines in the EPR spectra and cause a decrease in their amplitude which is present in Figure 2.5B.[33, 34] The broadening of the EPR spectral linewidth is quantitatively determined by measuring the central line width. In Figure 2.5A the free MTSL spectra shows a central line width of 1.4 G, while the MTSL bound to the pinholin, 5B, has a central line width of 2.9 G. The line broadening of the EPR spectra from Figure 2.5A to 2.5B confirms the successful disulfide bond formation between the free MTSL and the Cys side chain of the pinholin due to a more restricted environment for the bound SL. Incorporation of the pinholin into a lipid mimetic system should restrict the motion of the spin label even further through interactions with the lipid’s hydrocarbon acyl chains. Since the broadening of the EPR spectra is proportional to the mobility of the spin label, the EPR spectra from pinholin incorporated into bicelles and MLVs show a greater degree of broadening than the labeled protein in solution (TFE) as seen when comparing Figure 2.5B to 2.5C, D. The central line widths of pinholin in bicelles versus pinholin in MLVs are 3.2 G and 3.3 G, respectively. The similarity of the central line width between the lipid incorporated samples is due to the similarity of the local environment around the spin label. Both bicelles and MLVs are good mimetics for lipid bilayers as opposed to a mimetic like micelles, which only mimic the hydrophobic environment of the membrane but cannot recreate the conditions of a bilayer. Therefore, the local environment and acyl chain packing around the spin label for both bicelles and MLVs will be similar in either mimetic.

32

2.4.6 31P Solid State – NMR Spectroscopy of Pinholin Solid State NMR spectroscopy is a powerful biophysical technique which utilizes the presence of directionally dependent anisotropic interactions to probe the dynamics or kinetics of a system, specifically the membrane system for this study.[35] 31P SS NMR experiments were used to measure the chemical shift anisotropy (CSA) of the phosphorus head groups of DMPC lipids.[20] The degree in which the pinholin interacts with the 31P lipid head group will help to probe the differences between the roles of the active S2168 and inactive S21IRS forms of the pinholin in the lytic pathway from the perspective of the membrane. 31P SS-NMR samples were prepared using the wild type active S2168 and inactive S21IRS forms of the pinholin incorporated into DMPC MLVs at 1 mol%. More protein is required here to account for the lower sensitivity of NMR when compared to EPR spectroscopy. Empty DMPC MLVs, active S2168 pinholin in DMPC MLVs, and inactive S21IRS pinholin in DMPC MLVs at 25°C are shown in Figures 2.6A, B, and C, respectively. The shape of the static 31P SS-NMR spectra for the empty MLVs, active S2168, and inactive S21IRS are characteristic of lamellar phase lipid mimetics and show axial symmetry. The CSA width for each of the NMR spectra were found to be 48.0 ppm for empty DMPC MLVs, 44.7 ppm for active S2168 pinholin incorporated in DMPC MLVs, and 42.2 ppm for the inactive S21IRS pinholin incorporated in DMPC MLVs. The smaller CSA width for both the active S2168 and inactive S21IRS pinholin when compared to the empty MLVs confirms the interaction of the protein with the lipid mimetic system. The absence of an isotropic peak suggests the integrity of the MLV mimetic system is not compromised with the addition of the pinholin protein. The smaller CSA for the inactive S21IRS pinholin (42.2 ppm) indicates the inactive form is influencing the 31P DMPC lipid head groups more than the active S2168 form. These differences suggest that the active S2168 and inactive S21IRS pinholins behave differently once incorporated into the lipid bilayer. This difference 31 2 will be further explored in future studies using both P T1 measurements and H lipid acyl chain experiments.

33

2.5 Conclusion: In this study we report the synthesis of both the active S2168 and inactive S21IRS forms of the pinholin protein using solid phase peptide synthesis. The measured CD data of both forms of the pinholin, matched the predicted alpha helical content. The CD molar ellipticity ratio also provided preliminary data of the helical packing required of the pinholin to attain functionality and will be explored to a greater extent in future works. CW-EPR spectroscopy was used to successfully show spin labeling of the pinholin as well as incorporation into both bicelles and MLV lipid mimetic systems. The success of this measurement opens the door for more in depth EPR structural and dynamic studies to be conducted in the future, such as pulsed EPR measurements.[36-40] 31P SS-NMR spectroscopy allowed for the study of the pinholin system from the lipid perspective and showed interactions of both forms of the pinholin with the lipid membrane, to varying degrees, through decreases in the CSA width when compared to the empty DMPC MLVs. These differences in the way the active S2168 and inactive S21IRS forms of the pinholin interact with the membrane suggest differences in the externalization of TMD1, and 31 ultimately the role each form plays in the bacteriophage lytic pathway. Additionally, P T1 relaxation time measurements and 2H NMR experiments can be conducted to further probe the interaction of the pinholin with the acyl chain and lipid head groups to better understand how S2168 and S21IRS differ in their membrane interaction.

2.6 Acknowledgments: We would like to thank members of the Young group at Texas A&M University for their enthusiasm in the continued pinholin work as well as their experimental suggestions. This work was generously supported by a NSF CHE-1807131 grant and a NIGMS/NIH Maximizing Investigator’s Research Award (MIRA) R35 GM126935 award. Gary A. Lorigan would also like to acknowledge support from the John W. Steube Professorship.

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2.7 Figures:

Figure 2.1: This table shows the primary sequence for the active S2168 form of pinholin, its cognate antiholin S2171, and the inactive S21IRS form of the pinholin. Transmembrane domains (TMD) 1 and 2 are underlined with the spin labeled position highlighted in yellow. The three additional amino acids for the antiholin are shown in red with the positive charge responsible for timing denoted under the Lys. The IRS tag responsible for preventing TMD1 externalization of S2168 is highlighted in red with the additional two positive charges denoted on the Arg residues.

35

Figure 2.2: The schematic shows the hypothetical pinholin TMD1 and TMD2 conformations in the membrane as well as possible TMD1 externalization orientations.

36

Figure 2.3: The MALDI-TOF mass spectrum of active (blue) S2168-WT and S2168-L25C- MTSL with a WT m/z of 7546 and MTSL m/z of 7722 and the inactive (red) S21IRS-WT and S21IRS-L25C-MTSL with a WT m/z of 8222 and MTSL m/z of 8397. The shift of +185 comes from the successful coupling of the MTSL to the Cys substituted Leu25 position. The doubly charged ion peaks for all spectra can be seen at one half the target m/z value.

37

Figure 2.4: Circular Dichroism spectra of both the active S2168 (blue) and inactive S21IRS (red) forms of the pinholin protein in DMPC MLVs showing local minima at 208 and 222 nm with a large positive peak at 195 nm, indicative of alpha helical secondary structure.

38

Figure 2.5: CW-EPR spectra of A) free MTSL at 300 µM in water B) MTSL coupled to the active S2168 pinholin protein at Leu25 dissolved in TFE C) MTSL labeled pinholin incorporated into 1:500 DMPC/DHPC bicelles and D) MTSL labeled pinholin incorporated into 1:500 DMPC MLVs.

39

Figure 2.6: Static 31P solid-state NMR spectrum of: A) empty DMPC MLVs (black). B) 1mol% active S2168-WT pinholin in DMPC MLVs (blue). C) 1mol% inactive IRS-WT pinholin in DMPC MLVs (red).

40

References:

[1] R. Young, I. Wang, Phage Lysis. The Bacteriophages, 2nd Ed ed., Oxford Univ Press, Oxford, 2006. [2] R. Young, Phage Lysis: Do we have the hole story yet?, Curr Opin Microbiol., 16 (2013) 1-8. [3] R. Young, Phage Lysis: Three Steps, Three Choices, One Outcome, Journal of Microbiology, 52 (2014) 243-258. [4] R. Young, I.N. Wang, W.D. Roof, Phages will out: strategies of host cell lysis, Trends in Microbiology, 8 (2000) 120-128. [5] R. Young, Bacteriophage holins: Deadly diversity, Journal of Molecular Microbiology and Biotechnology, 4 (2002) 21-36. [6] T. Park, D.K. Struck, C.A. Dankenbring, R. Young, The pinholin of lambdoid phage 21: Control of lysis by membrane depolarization, Journal of Bacteriology, 189 (2007) 9135- 9139. [7] M. Xu, D.K. Struck, J. Deaton, I.N. Wang, R. Young, 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, 101 (2004) 6415-6420. [8] J.H. Grose, S.R. Casjens, Understanding The Enormous Diversity Of Bacteriophages: The Tailed Phages That Infect The Bacterial Family Enterobacteriaceae, Virology, 468 (2014) 421-443. [9] M. Barenboim, C.Y. Chang, F.D. Hajj, R. Young, Characterization of the dual start motif of a class II holin gene, Molecular Microbiology, 32 (1999) 715-727. [10] M.T. Bonovich, R. Young, dual start motif in 2 lambdoid s-genes unrelated to lambda- s, Journal of Bacteriology, 173 (1991) 2897-2905. [11] T. Pang, C.G. Savva, K.G. Fleming, D.K. Struck, R. Young, Structure of the lethal phage pinholin, PNAS, 106 (2009) 18966-18971. [12] T. Pang, T. Park, R. Young, Mutational analysis of the S21 pinholin, Molecular Microbiology, 76 (2010) 68-77. [13] T. Park, D.K. Struck, J.F. Deaton, R. Young, Topological dynamics of holins in programmed bacterial lysis, Proceedings of the National Academy of Sciences of the United States of America, 103 (2006) 19713-19718. [14] T. Pang, T. Park, R. Young, Mapping the pinhole formation pathway of S21, Molecular Microbiology, 78 (2010) 710-719. [15] T. Pang, T.C. Fleming, K. Pogliano, R. Young, Visualization of pinholin lesions in vivo, Proceedings of the National Academy of Sciences of the United States of America, 110 (2013) E2054-E2063. [16] P.R. Hansen, A. Oddo, Fmoc Solid-Phase Peptide Synthesis, Peptide Antibodies: Methods and Protocols, 1348 (2015) 33-50. [17] P. Lloyd-Williams, F. Albericio, E. Giralt, Chemical Approaches to the Synthesis of Peptides and Proteins, CRC Press1997. [18] D.S. King, C.G. Fields, G.B. Fields, a cleavage method which minimizes side reactions following fmoc solid-phase peptide-synthesis, International Journal of Peptide and Protein Research, 36 (1990) 255-266.

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[19] F. Albericio, N. Kneibcordonier, S. Biancalana, L. Gera, R.I. Masada, D. Hudson, G. Barany, Preparation And Application Of The 5-(4-(9- Fluorenylmethyloxycarbonyl)Aminomethyl-3,5-Dimethoxyphenoxy)Vale Ric Acid (Pal) Handle For The Solid-Phase Synthesis Of C-Terminal Peptide Amides Under Mild Conditions, Journal of Organic Chemistry, 55 (1990) 3730-3743. [20] S. Abu-Baker, G.A. Lorigan, Phospholamban and its phosphorylated form interact differently with lipid bilayers: A (31)P, (2)H, and (13)C solid-state NMR spectroscopic study, Biochemistry, 45 (2006) 13312-13322. [21] I.D. Sahu, R.M. McCarrick, G.A. Lorigan, Use of Electron Paramagnetic Resonance To Solve Biochemical Problems, Biochemistry, 52 (2013) 5967-5984. [22] T.B. Cardon, E.K. Tiburu, G.A. Lorigan, Magnetically Aligned Phospholipid Bilayers In Weak Magnetic Fields: Optimization, Mechanism, And Advantages For X-band EPR Studies, Journal of Magnetic Resonance, 161 (2003) 77-90. [23] L. Whitmore, B.A. Wallace, Protein Secondary Structure Analyses From Circular Dichroism Spectroscopy: Methods And Reference Databases, Biopolymers, 89 (2008) 392-400. [24] A. Abdul-Gader, A.J. Miles, B.A. Wallace, A Reference Dataset For The Analyses Of Membrane Protein Secondary Structures And Transmembrane Residues Using Circular Dichroism Spectroscopy, Bioinformatics, 27 (2011) 1630-1636. [25] L.A. Compton, W.C. Johnson, Analysis Of Protein Circular-Dichroism Spectra For Secondary Structure Using A Simple Matrix Multiplication, Analytical Biochemistry, 155 (1986) 155-167. [26] P. Manavalan, W.C. Johnson, Variable Selection Method Improves The Prediction Of Protein Secondary Structure From Circular-Dichroism Spectra, Analytical Biochemistry, 167 (1987) 76-85. [27] N. Sreerama, R.W. Woody, Estimation Of Protein Secondary Structure From Circular Dichroism Spectra: Comparison of CONTIN, SELCON, and CDSSTR Methods With An Expanded Reference Set, Analytical Biochemistry, 287 (2000) 252-260. [28] D. Mao, E. Wachter, B.A. Wallace, Folding Of The Mitochondrial Proton Adenosine- Triphosphatase Proteolipid Channel In Phospholipid-Vesicles, Biochemistry, 21 (1982) 4960-4968. [29] J. Liu, Y. Zheng, C.-S. Cheng, N.R. Kallenbach, M. Lu, A seven-helix coiled coil., PNAS, 103 (2006) 15457-15462. [30] N.E. Zhou, C.M. Kay, R.S. Hodges, Synthetic Model Proteins - Positional Effects Of Interchain Hydrophobic Interactions On Stability Of 2-Stranded Alpha-Helical Coiled- Coils, Journal of Biological Chemistry, 267 (1992) 2664-2670. [31] S.M. Kelly, T.J. Jess, N.C. Price, How to study proteins by circular dichroism, Biochimica Et Biophysica Acta-Proteins and Proteomics, 1751 (2005) 119-139. [32] T.T. Zheng, A. Boyle, H.R. Marsden, D. Valdink, G. Martelli, J. Raap, A. Kros, Probing coiled-coil assembly by paramagnetic NMR spectroscopy, Organic & Biomolecular Chemistry, 13 (2015) 1159-1168. [33] I.D. Sahu, G.A. Lorigan, Site-Directed Spin Labeling EPR for Studying Membrane Proteins, Biomed Research International, (2018). [34] I.D. Sahu, A.F. Craig, M.M. Dunagan, K.R. Troxel, R.F. Zhang, A.G. Meiberg, C.N. Harmon, R.M. McCarrick, B.M. Kroncke, C.R. Sanders, G.A. Lorigan, Probing Structural Dynamics and Topology of the KCNE1 Membrane Protein in Lipid Bilayers via Site-

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Directed Spin Labeling and Electron Paramagnetic Resonance Spectroscopy, Biochemistry, 54 (2015) 6402-6412. [35] T.R. Molugu, S. Lee, M.F. Brown, Concepts and Methods of Solid-State NMR Spectroscopy Applied to Biomembranes, Chemical Reviews, 117 (2017) 12087-12132. [36] I.D. Sahu, B.M. Kroncke, R.F. Zhang, M.M. Dunagan, H.J. Smith, A. Craig, R.M. McCarrick, C.R. Sanders, G.A. Lorigan, Structural Investigation of the Transmembrane Domain of KCNE1 in Proteoliposomes, Biochemistry, 53 (2014) 6392-6401. [37] E.F. Vincent, I.D. Sahu, A.J. Costa-Filho, E.M. Chilli, G.A. Lorigan, Conformational changes of the HsDHODH N-terminal Microdomain via DEER Spectroscopy, J. Phys. Chem. B, 119 (2015) 8693-8697. [38] L.S. Liu, G. Lorigan, Probing the Secondary Structure of Membrane Proteins with the Pulsed EPR ESEEM Technique, Biophysical Journal, 106 (2014) 192A-192A. [39] L.S. Liu, I.D. Sahu, L. Bottorf, R.M. McCarrick, G.A. Lorigan, Investigating the Secondary Structure of Membrane Peptides Utilizing Multiple H-2-Labeled Hydrophobic Amino Acids via Electron Spin Echo Envelope Modulation (ESEEM) Spectroscopy, Journal of Physical Chemistry B, 122 (2018) 4388-4396. [40] G.A. Lorigan, Probing the Structure of Membrane Proteins with ESEEM and DEER Pulsed EPR Techniques, Biophysical Journal, 102 (2012) 423A-423A.

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Chapter 3

Probing the Local Secondary Structure of Bacteriophage S21 Pinholin Membrane Protein using Electron Spin Echo Envelope Modulation Spectroscopy

Daniel L. Drew Jr., Rachel A. Serafin, Tanbir Ahammad, Indra D. Sahu, Robert M. McCarrick, Gary A. Lorigan*

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

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3.1 Abstract: There have recently been advances in biophysical techniques for detecting local secondary structures of membrane protein using electron paramagnetic resonance (EPR). One of these methods is the three pulse electron spin echo envelope modulation (ESEEM) approach which can be utilized to determine the local secondary structure of membrane proteins. The pulsed EPR ESEEM technique was used in this work to probe the local secondary structure of the small hole forming membrane protein, pinholin. This poorly characterized protein is responsible for the depolarization of the inner cytosolic membrane of double stranded DNA bacteriophage host cells. In this study circular dichroism (CD) spectra of pinholin in a membrane, with double minima present at 208 nm and 222 nm, indicated that the global secondary structure was α-helical. Secondary structural calculations using CD data fitting DICHROWEB software showed ~60% α- helical content. Comparable CD data with and without the spin label indicate no significant difference in the secondary structure of pinholin. The local α-helical secondary structure was confirmed using three pulse ESEEM spectroscopy for both transmembrane domains for the active S2168 and inactive S21IRS forms of pinholin. Comparison of the ESEEM normalized frequency domain intensity for each i-n transmembrane position determined the α-helical folding nature of these domains and excluded the possibility of π or 310- helices.

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3.2 Introduction: Membrane proteins are responsible for a wide variety of cellular functions including transport, signaling, lysis, and are targets for over 50% of small molecule drug binding.1- 3 However, determining membrane protein secondary structure has proved to be a challenge for the scientific community due to their hydrophobic nature, poor overexpression yields, and lack of high quality crystals.4, 5 Previously the Lorigan group has demonstrated that electron spin echo envelope modulation (ESEEM) spectroscopy coupled with site direct spin labeling (SDSL) and 2H isotopic amino acid labeling can be utilized to determine the local secondary structure of model membrane proteins.6-9 This technique has been used to differentiate between α-helices and β-sheets as well as detect 6 the presence of α-helices versus 310-helices. The use of this ESEEM technique gives advantage over other biophysical structural techniques, like circular dichroism (CD), as the experimental design allows for selective probing of local secondary structure as opposed to global structures. Knowing the local secondary structure is critical as subtle changes in structure have been shown to affect the packing of membrane proteins or their incorporation and interactions with the surrounding lipid environment.10 There are also families of membrane proteins which must undergo local conformational changes and refolding of their secondary structure in order to attain functionality.11, 12 This ESEEM approach therein overcomes concentration and size limitations seen in CD or NMR spectroscopy and supplies a method for determining these small, yet crucial changes in the local secondary structure of proteins.

This ESEEM technique requires a deuterated amino acid side chain, such as d10 Leu, with a site-specific cysteine substitution within four amino acids of the 2H-labeled amino acid. A nitroxide spin label, in this case S-(1-oxyl-2,2,5,5-tetramethyl-2,5-dihydro- 1H-pyrrol-3-yl)methyl methanesulfonothioate (MTSL), is attached via disulfide bond formation to the cysteine sidechain. ESEEM spectroscopy detects the weak dipolar interaction between the unpaired electron of the spin label and the deuterium of the labeled Leu sidechain within a detection limit of 8 Å.13, 14 The characteristic periodicity of helices and the linear nature of β-sheets reveals unique patterns in the ESEEM spectra as the spin label is moved step wise away from the d10 Leu. In a helix, when the spin label is at a position 2 amino acids away from the d10 Leu, the spin label and the d10 Leu appear

46 on opposite sides on the helix and therefore outside of the 8 Å ESEEM detection limit. As the spin label is moved to positions 3 or 4 amino acids away from the d10 Leu the helical structure puts both labels on the same side of the helix allowing for deuterium echo modulation to be observed.8 Alternatively, the exact opposite pattern would be detected when probing a β-sheet. The linear nature of β-sheets would place the spin label within the detection limit at position 1 or 2 amino acids away from the d10 Leu allowing for deuterium modulation. Unlike the α-helix there would be no deuterium modulation detected at positions 3 and 4 amino acids away from the d10 Leu as the linear β-stand would place the spin label more than 8 Å away from the d10 Leu. To this point, the application of the ESEEM technique has been mostly limited to model peptides or small protein segments of known structure.6, 8 This study will apply the outlined ESEEM approach to a more complicated full-length membrane protein of unknown structure known as pinholin. Until this point, the predicted local helical secondary structure for pinholin has only been hypothesized using computational molecular modeling simulations and through comparison to other classes of bacteriophage holin proteins.15 Thus, more structural studies are needed to investigate the complicated membrane protein system. The function of the pinholin protein is to depolarize the cytosolic membrane during double stranded DNA bacteriophage host cell lysis. This depolarization step allows the release of the signal-anchored-release (SAR) endolysin from the membrane to begin the degradation of the peptidoglycan.16 The mechanism for membrane depolarization is accomplished through externalization of the first of two pinholin transmembrane domains (TMD) from the membrane. This results in pinholin shifting from an inactive form, with both TMD1 and TMD2 are incorporated into the membrane, to an active form of pinholin with TMD1 externalized. Functional studies of pinholin have shown that truncated forms of pinholin with only TMD2 present still show lytic function. Therefore, TMD2 is considered the functional domain of pinholin while, TMD1 is the inhibitory domain. The lysis pathway for the remaining membrane incorporated TMD2 involves the oligomerization of pinholin until the protein “triggers.” Triggering is the term used to denote when the pinholin reaches a critical concentration in the membrane and undergoes a in the membrane forming heptameric “pinholes.”15 Due to the variation in mechanisms and sizes of lesions formed between different classes of holins the lesions have been termed

47

“holes” to show distinction from channels and other such membrane permeabilization pathways.3, 15, 17 Studying the secondary structure of the pinholin’s TMDs in an inactive and active form is vital for understanding the mechanism of not only pinholin activation but ultimately the oligomerization and resulting depolarization of the membrane. There is evidence of channel and hole forming membrane proteins undergoing a refolding of helical secondary structure into α-, 310-, or π-helices in changing environmental conditions. The work presented here will be the first set of biophysical experiments aimed at probing the local secondary structure of the TMDs of pinholin in an active and inactive form.

3.3 Experimental Methods: Pinholin is encoded by the S21 holin gene of the lambdoid bacteriophage ϕ21. The S2168 is the active form of the pinholin, while the S21IRS will represent the inactive or nonfunctional form.18 All pinholin proteins were synthesized using solid phase peptide synthesis conducted on a CEM Liberty Blue Peptide Synthesizer with Discovery Bio Microwave System.19 Fourteen different variations of the active S2168 and inactive S21IRS forms were created to complete the study. Table 1 outlines all fourteen different deuterated and spin labeled pinholin positions.

48

Table 3.1: Active and Inactive Pinholin Primary Sequence

Table 3.1: This table shows the primary sequence of active S2168 and inactive S21IRS pinholin. The deuterated d10 Leu position is denoted by i while the position of the cysteine substitution is shown as X. The underlined sections of the primary sequence correspond to the two predicted transmembrane domains.

These pinholin constructs were generated to position a d10 Leu amino acid at a specific position designated as i. A cysteine residue, denoted in Table 1 as X, is then substituted in at specific position -2, -3, or -4 residues away from the deuterated leucine, i. This cysteine residue allows for disulfide bond formation to the nitroxide spin label MTSL to make the pinholin peptide EPR active. A control sample was synthesized by creating an i-4 pinholin with a natural (undeuterated) leucine side chain at position Leu25. After successful solid phase synthesis, the pinholin was cleaved from the solid phase resin using a trifluoroacetic acid (TFA) cleavage solution which also removed the remaining amino acid side chain protecting groups. The resin bound protein was added to the cleavage solution of TFA/Triisoproylsilane/1,2-Ethanedithiol/Water (94%/1%/2.5%/2.5%) and left to stir for 3 hrs at room temperature.20 Once complete, the solid phase resin was removed from the cleavage reaction through gravity filtration and the remaining TFA was evaporated off by inert N2 gas flow to an approximate volume of

49

5 mL. The peptide was separated from the still soluble scavengers through addition of methyl tert-butyl ether to precipitate the pinholin. Any remaining TFA or scavengers were removed by performing three additional centrifugations with ether washes. The final ether wash was decanted off and the crude pinholin was dried under vacuum overnight. The crude pinholin was purified using reverse-phase high pressure liquid chromatography (RP-HPLC) with a C4 prep column run at a gradient of 5 to 95% Solvent B (90% acetonitrile/10% water/0.1% TFA).21 The collected pure pinholin fractions were lyophilized, and the resulting pure pinholin was spin labeled using S-(1-oxyl-2,2,5,5- tetramethyl-2,5-dihydro-1H-pyrrol-3-yl)methyl methanesulfonothioate (MTSL). Labeling of the pinholin with the nitroxide spin label MTSL was run at a 5-fold molar excess in dimethyl sulfoxide (DMSO) for 24 hrs. The resulting solution was then lyophilized, and the remaining crude spin labeled pinholin powder was purified using reverse-phase HPLC to remove the excess spin label. This purification was run using a C4 semiprep column with the same gradient and solvent system as previously mentioned. Matrix assisted laser desorption ionization – time of flight (MALDI-TOF) was used to confirm the target molecular weight of the pinholin plus successful MTSL coupling. All the remaining pure, spin labeled pinholin fractions were lyophilized into a powder to use for lipid incorporation and the resulting experiments.18 The pinholin peptides were determined to be approximately 90% pure after purification. The pure spin labeled pinholins were incorporated into two different lipid mimetic systems at a 500:1 lipid to protein ratio. 1,2-Dimyristoyl - sn - Glycero - 3 - Phosphocholine (DMPC) / 1,2 - Diheptanoyl - sn - Glycero – 3 - Phosphocholine (DHPC) bicelles were created by mixing the 2 lipids, dissolved in chloroform, with a q-value of 3.6 and then adding the corresponding amount of pinholin dissolved in 2,2,2-Trifluoroethanol (TFE).

The solvents were evaporated off using inert N2 gas and the remaining lipid/protein film was rehydrated using 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer at a concentration of 20 mM adjusted to a neutral pH of ~7.0. The lipid/protein solution was flash frozen in liquid nitrogen and then vortexed and sonicated until the buffer solution went clear. The second lipid mimetic used were multilamellar vesicles (MLV) created by mixing the peptide dissolved in 2,2,2-Trifluoroethanol (TFE) with DMPC in chloroform at a 500:1 lipid to protein ratio. Then the solvents were evaporated off using inert N2(g) and

50 the same process as bicelle formation was followed with a minimum of three freeze thaw cycles. The global secondary structure was determined using circular dichroism (CD) spectroscopy where the thin-film sample was rehydrated with a mixture of 10% 10mM HEPES buffer/90% 15 mM buffer. The dilution of HEPES buffer with phosphate buffer was needed to counteract the of HEPES at wavelengths lower than 200 nm. The measurements were performed on an Aviv Circular Dichroism Spectrometer Model 435 in a quartz cuvette with a 1.0 mm path length. Data was collected from 260 nm to 190 nm with 1 nm bandwidth at 25°C. X-band CW-EPR (~9 GHz) was used to calculate spin labeling efficiency with all samples showing greater than 85% labeling efficiency. Three-pulse (π/2 – τ – π/2 – T – π/2 – τ – echo) ESEEM measurements were conducted on a Bruker ELEXSYS E580 with an ER4118X MS3 resonator using a 200 ns tau value for 1H modulation suppression. The ESEEM data for each sample was collected at a microwave frequency of ~9.269 GHz and a magnetic field of ~3300 G at a temperature of 80 K with 4-step phase cycling. The spectra were collected with a starting T value of 386 ns with an increase in 12 ns increments for a total of 512 points.6, 7 All ESEEM data was collected on a 45 µL sample incorporated in to either 500:1 DMPC MLVs or 500:1 DMPC/DHPC bicelles. A two- component exponential decay was used to fit the time domain data. The maximum value for the exponential fit as well as the collected time domain data were both scaled to one, according to the literature.22 The normalized decay curve was subtracted from the experimental data to give a scaled ESEEM spectrum.23-25 A Fourier Transformation (FT) was then applied to the scaled ESEEM spectra to yield the corresponding frequency domain.26 The detected deuterium peak appears at ~2.3 MHz corresponding to the Larmor frequency of 2H.

3.4 Results and Discussion: In the pathway of bacteriophage cell lysis, the pinholin protein is responsible for the permeabilization, and resulting depolarization, of the inner cell membrane.27, 28 In this study, CD spectroscopy as well as an established three pulse ESEEM approach were used to determine both global and local secondary structure of the active and inactive

51 forms of pinholin in the membrane, respectively. Figure 3.1 shows the CD spectra of the 21 deuterated and spin labeled S 68 – d10 Leu50 (i-3) incorporated into DMPC MLVs at a 500:1 lipid to protein ratio. This data shows a large positive peak near 195 nm and double minima at 208 and 222 nm corresponding to a global α-helical structure. Secondary structural content calculations were performed using DICHROWEB software and resulted in ~60% α-helical structure. This indicates the deuterated side chain and site-directed spin labeling have no significant impact on the global helical secondary structure when compared to the previously publish wildtype pinholin data.18 This CD data is also comparable for both the active and inactive forms of pinholin. Following the confirmation that the labels are not impacting the global folding of pinholin the ESEEM approach outlined above was used to probe the local secondary structure. Positions were chosen within both TMDs predicted by the literature to be helical in nature.15, 29 These experiments were conducted for both the active S2168 and inactive S21IRS forms of pinholin.29 Position L25 is located in the first predicted helical domain, while L50 is located in the second transmembrane domain. Study of the local secondary structure of L25 in TMD1 for the active S2168 and inactive S21IRS forms of pinholin probed any conformational changes in TMD1 when spanning the membrane or after externalization. The L50 position in TMD2 of the inactive S21IRS pinholin determines the secondary structure of the functional domain of pinholin while in an inactive form. Any changes in the secondary structure of the functional domain of pinholin during oligomerization or hole formation will be seen by comparing the active S2168 to the inactive S21IRS data of the L50 site. Figure 3.2 shows three pulse ESEEM spectra of active S2168 form at pinholin in DMPC lipid bilayers. Deuterium modulation is observed for the i-3 and i-4 positions at the Larmor frequency. The modulation depth seen in the frequency domain is proportional to 1/r6, in which r is the distance between the spin label and the 2H labeled found on the Leu sidechain. This modulation in the time domain results in a peak in the frequency domain that can be observed in the i-3 and i-4 positions of the spin label in Figure 3.2B. There is some variability present in the intensity between each position when comparing the different TMDs in the active and inactive forms of pinholin within the same mimetic system. Intensities of the i-4 positions ranged from 0.13 – 0.21 in liposomes, while the i-

52

3 ranged from 0.14 – 0.17. This variability is due to different local environments between the 2 different TMDs. There are also multiple r values between each of the individual 2H atoms found on the d10 Leu side chain and the spin label. The distance can be affected by the spin label itself as it can adopt multiple conformations through the 5 torsion angles found throughout the sidechain.15, 19, 27, 29-31 The presence of the deuterium peak at positions i-3 and i-4 with the absence of the peak at i-2 is indicative of an α-helical secondary structure.7, 32, 33 This pattern comes from the 3.6 amino acid periodicity of the α-helical secondary structure. This places the spin label, at i-3 and i-4, and the deuterated leucine side chain on the same face of the helix allowing dipolar coupling while the i-2 position falls on the opposite face of the helix outside of the 8 Å detection limit.34 Figure 3.3 shows the normalized ESEEM frequency data for all pinholin data collected in DMPC/DHPC 1:500 bicelles (A) and DMPC 1:500 liposomes (B). The α- helical pattern can be observed when looking at deuterated position L25 located in TMD1 (top row) for both the active S2168 (blue) and the inactive S21IRS (red). This data indicates that regardless of the lipid mimetic system (bicelles or liposomes), the local α-helical structure of TMD1 is not changing between being membrane incorporated or after translocation. This α-helical secondary structure is also present in the functional domain, TMD2, of both active and inactive forms of pinholin. This is seen in the bottom row of Figure 3.3 with deuterated position L50 showing deuterium dipolar coupling peaks for the i-4 position, but not for i-2. There is a small observable peak in the i-2 at the d10 L50 position in TMD2 for only the S2168 active form of pinholin. This may be due to the helical packing of multiple TMD2s in the oligomeric state of the pinholin. The inability of the inactive S21IRS form of pinholin to reach this oligomeric state would indicate why this is only observed in TMD2 of the active S2168 form. The presence of this peak, along with the previous results of TMD1 folded as an α-helix in both forms of pinholin, would suggest that there is no need for a refolding of either TMD during the oligomerization step of the pinholin depolarization pathway. A comparison of the normalized frequency domain intensity was conducted. Due to unique turn periodicities for different helices, such as 3.1 for a 310 helix or 4.1-4.4 for a π helix, comparing the normalized frequency domain intensity will allow for differentiation

53 between these helical structures.6, 35, 36 The intensity comparison between the i-3 and i-4 peaks for bicelle incorporated pinholin yields frequency intensities within 4% of each other with the i-4 positions all having slightly higher intensities. This pattern would be expected and consistent with the accept turn periodicity of 3.6 amino acids per turn. The liposome samples followed this same trend showing intensities within 13% of the i-3 and i-4 positions. These results support the conclusion that, for the case of the inhibitory domain, TMD1, of pinholin the helix is folding as an α-helix and there is no change in this secondary structure as the helix externalizes from the bilayer. The data for the TMD2 L50 position in the functional domain in the inactive S21IRS sample indicates an α-helical secondary structure. This α-helical secondary structure of TMD2 is also retained through the externalization of TMD1 and possibly, due to the presence of the small peak observed in the i-2 positions discussed previously, conserved during the oligomerization in the penultimate step of the depolarization pathway.15

3.5 Conclusions: Knowing membrane protein secondary structure is pivotal in understanding protein functions and dynamics. Techniques such as CD spectroscopy can give global secondary structure information but lack the ability to specify local secondary structures. As opposed to techniques like NMR which require higher sample concentrations and are restricted by protein size, this ESEEM approach requires only ~50 µM and ~50 µL of sample and has no limit on protein size.6, 8, 33 The application of ESEEM outlined in this paper demonstrated the ability to probe local protein secondary structure, not only for model peptides as previously published, but of full-length functional systems. The α-helical local secondary structure of both predicted pinholin transmembrane domains was confirmed with the presence of deuterium modulation observed at the i-3 and i-4 positions.12 The confirmation of the α-helical structure in TMD1 and TMD2 for both the active S2168 and inactive S21IRS forms of the pinholin demonstrates the applicability of this technique to both peripheral and integral membrane proteins. In addition, the comparison of ESEEM frequency domain intensities between the active S2168 and inactive S21IRS conformations indicates there is little to no change in helical structure after TMD1 externalizes from the membrane. This work, along with previously published

54 work from the Lorigan group, demonstrated that the lipid composition or memetic system have no effect on in vitro protein studies.6, 8, 21, 33 Future application of this work will include probing helical boundaries to observe the loss of helical folding pattern to identify the end point of the helix. This ESEEM approach, along with EPR DEER spectroscopy, will be utilized to probe tertiary interactions of functional systems known to form dimers or oligomeric states and the use of deuterated buffers to probe membrane protein topology.

3.6 Acknowledgment: This work was generously supported by a NSF CHE-1807131 grant and a NIGMS/NIH Maximizing Investigator’s Research Award (MIRA) R35 GM126935 award. The pulsed EPR spectrometer used to conduct he experiments was purchased through the NSF and the Ohio Board of Reagents grants (MRI-0722403). Gary A. Lorigan would also like to acknowledge support from the John W. Steube Professorship.

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3.7 Figures:

21 Figure 3.1: CD spectra of the active S 68 – d10 L50 (i-3) pinholin incorporated into DMPC MLVs at a 500:1 lipid to protein ratio. The double minimums at 208 nm and 222 nm are indicative of α-helical global secondary structure of pinholin and validate that the deuterated and spin labeled side chains do not affect folding.

56

Figure 3.2: Three pulse ESEEM experimental data for the active S2168 pinholin form with 2 H labeled d10 Leu side chain at position L25 in TMD1 incorporated into DMPC liposomes at 500:1 lipid to protein ratio. 2H modulation can be observed in the time domain (A) at positions i-4 and i-3 which translates to the peaks seen in the normalized frequency domain intensity (B).

57

A) B)

DMPC/DHPC Bicelles DMPC Liposomes

Figure 3.3: Normalized ESEEM frequency domain spectra for active S2168 pinholin (blue) and inactive S21IRS pinholin (red) for transmembrane domain 1 (top) and transmembrane domain 2 (bottom). A) The ESEEM data set of active and inactive pinholin incorporated into DMPC/DHPC bicelles at a 500:1 lipid to protein ratio. B) The ESEEM data set of active and inactive pinholin incorporated into DMPC liposomes at a 500:1 lipid to protein ratio

58

References:

1. Lappano, R.; Maggiolini, M., G protein-coupled receptors: novel targets for drug discovery in cancer. Nature Reviews Drug Discovery 2011, 10 (1), 47-60. 2. Tautermann, C. S., GPCR structures in drug design, emerging opportunities with new structures. Bioorganic & Medicinal Chemistry Letters 2014, 24 (17), 4073-4079. 3. Young, R., Phage Lysis: Three Steps, Three Choices, One Outcome. Journal of Microbiology 2014, 52 (3), 243-258. 4. Bordag, N.; Keller, S., alpha-Helical transmembrane peptides: A "Divide and Conquer" approach to membrane proteins. Chemistry and Physics of Lipids 2010, 163 (1), 1-26. 5. Huang, C. D.; Mohanty, S., Challenging the Limit: NMR Assignment of a 31 kDa Helical Membrane Protein. Journal of the American Chemical Society 2010, 132 (11), 3662-+. 6. Bottorf, L.; Rafferty, S.; Sahu, I. D.; McCarrick, R. M.; Lorigan, G. A., Utilizing Electron Spin Echo Envelope Modulation To Distinguish between the Local Secondary Structures of an alpha-Helix and an Amphipathic 3(10)-Helical Peptide. Journal of Physical Chemistry B 2017, 121 (14), 2961-2967. 7. Liu, L. S.; Lorigan, G., Probing the Secondary Structure of Membrane Proteins with the Pulsed EPR ESEEM Technique. Biophysical Journal 2014, 106 (2), 192A-192A. 8. Liu, L. S.; Sahu, I. D.; Bottorf, L.; McCarrick, R. M.; Lorigan, G. A., Investigating the Secondary Structure of Membrane Peptides Utilizing Multiple H-2-Labeled Hydrophobic Amino Acids via Electron Spin Echo Envelope Modulation (ESEEM) Spectroscopy. Journal of Physical Chemistry B 2018, 122 (16), 4388-4396. 9. Lorigan, G. A., Probing the Structure of Membrane Proteins with ESEEM and DEER Pulsed EPR Techniques. Biophysical Journal 2012, 102 (3), 423A-423A. 10. Kurochkina, N., Helix-helix interactions and their impact on protein motifs and assemblies. Journal of Theoretical Biology 2010, 264 (2), 585-592. 11. Gross, M., Proteins that Convert from to : Implications for Folding and Disease. Current Protein & Peptide Science 2000, 1 (4), 339-347. 12. Kubota, T.; Lacroix, J. J.; Bezanilla, F.; Correa, A. M., Probing alpha-3(10) Transitions in a Voltage-Sensing S4 Helix. Biophysical Journal 2014, 107 (5), 1117-1128. 13. Carmieli, R.; Papo, N.; Zimmermann, H.; Potapov, A.; Shai, Y.; Goldfarb, D., Utilizing ESEEM spectroscopy to locate the position of specific regions of membrane- active peptides within model membranes. Biophysical Journal 2006, 90 (2), 492-505. 14. Sun, L.; Hernandez-Guzman, J.; Warncke, K., OPTESIM, a versatile toolbox for numerical simulation of electron spin echo envelope modulation (ESEEM) that features hybrid optimization and statistical assessment of parameters. Journal of Magnetic Resonance 2009, 200 (1), 21-28. 15. Pang, T.; Savva, C. G.; Fleming, K. G.; Struck, D. K.; Young, R., Structure of the lethal phage pinholin. PNAS 2009, 106 (45), 18966-18971. 16. Park, T.; Struck, D. K.; Dankenbring, C. A.; Young, R., The pinholin of lambdoid phage 21: Control of lysis by membrane depolarization. Journal of Bacteriology 2007, 189 (24), 9135-9139. 17. Young, R., Phage Lysis: Do we have the hole story yet? Curr Opin Microbiol. 2013, 16 (6), 1-8.

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18. Drew, D. L.; 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 S-21 pinholin protein. Analytical Biochemistry 2019, 567, 14-20. 19. Chandrudu, S.; Simerska, P.; Toth, I., Chemical Methods for Peptide and Protein Production. Molecules 2013, 18 (4), 4373-4388. 20. King, D. S.; Fields, C. G.; Fields, G. B., A Cleavage Method Which Minimizes Side Reactions Following Fmoc Solid-Phase Peptide-Synthesis. International Journal of Peptide and Protein Research 1990, 36 (3), 255-266. 21. Mayo, D. J.; Inbaraj, J. J.; Subbaraman, N.; Grosser, S. M.; Chan, C. A.; Lorigan, G. A., Comparing the structural topology of integral and peripheral membrane proteins utilizing electron paramagnetic resonance spectroscopy. Journal of the American Chemical Society 2008, 130 (30), 9656-+. 22. Heming, M.; Narayana, M.; Kevan, L., Analysis Of Nuclear-Quadrupole Interaction Effects In Electron Spin-Echo Modulation Spectra By 2nd-Order Perturbation-Methods. Journal of Chemical Physics 1985, 83 (4), 1478-1484. 23. Urban, L.; Steinhoff, H. J., Hydrogen bonding to the nitroxide of protein bound spin labels. Molecular Physics 2013, 111 (18-19), 2873-2881. 24. Milov, A. D.; Samoilova, R. I.; Shubin, A. A.; Gorbunova, E. Y.; Mustaeva, L. G.; Ovchinnikova, T. V.; Raap, J.; Tsvetkov, Y. D., Self-Aggregation and Orientation of the Ion Channel-Forming Zervamicin IIA in the Membranes of ePC Vesicles Studied by cw EPR and ESEEM Spectroscopy. Applied Magnetic Resonance 2010, 38 (1), 75-84. 25. Bartucci, R.; Guzzi, R.; Sportelli, L.; Marsh, D., Intramembrane Water Associated with TOAC Spin-Labeled Alamethicin: Electron Spin-Echo Envelope Modulation by D2O. Biophysical Journal 2009, 96 (3), 997-1007. 26. Stoll, S.; Britt, R. D., General and efficient simulation of pulse EPR spectra. Physical Chemistry Chemical Physics 2009, 11 (31), 6614-6625. 27. Pang, T.; Park, T.; Young, R., Mapping the pinhole formation pathway of S21. Molecular Microbiology 2010, 78 (3), 710-719. 28. 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. 29. Pang, T.; Park, T.; Young, R., Mutational analysis of the S21 pinholin. Molecular Microbiology 2010, 76 (1), 68-77. 30. Columbus, L.; Kalai, T.; Jeko, J.; Hideg, K.; Hubbell, W. L., Molecular motion of spin labeled side chains in alpha-helices: Analysis by variation of side chain structure. Biochemistry 2001, 40 (13), 3828-3846. 31. Columbus, L.; Hubbell, W. L., A new spin on protein dynamics. Trends in Biochemical Sciences 2002, 27 (6), 288-295. 32. Zhou, A. D.; Abu-Baker, S.; Sahu, I. D.; Liu, L. S.; McCarrick, R. M.; Dabney- Smith, C.; Lorigan, G. A., Determining alpha-Helical and beta-Sheet Secondary Structures via Pulsed Electron Spin Resonance Spectroscopy. Biochemistry 2012, 51 (38), 7417-7419. 33. Zhang, R. F.; Sahu, I. D.; Gibson, K. R.; Muhammad, N. B.; Bali, A. P.; Comer, R. G.; Liu, L. S.; Craig, A. F.; McCarrick, R. M.; Dabney-Smith, C.; Sanders, C. R.; Lorigan, G. A., Development of electron spin echo envelope modulation spectroscopy to

60 probe the secondary structure of recombinant membrane proteins in a lipid bilayer. Protein Science 2015, 24 (11), 1707-1713. 34. Cieslak, J. A.; Focia, P. J.; Gross, A., Electron Spin-Echo Envelope Modulation (ESEEM) Reveals Water and Phosphate Interactions with the KcsA Potassium Channel. Biochemistry 2010, 49 (7), 1486-1494. 35. Kumar, P.; Bansal, M., Dissecting pi-helices: sequence, structure and function. Febs Journal 2015, 282 (22), 4415-4432. 36. Low, B. W.; Baybutt, R. B., The Pi-Helix - A Hydrogen Bonded Configuration Of The Polypeptide Chain. Journal of the American Chemical Society 1952, 74 (22), 5806- 5807.

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Chapter 4

Active S2168 and Inactive S21IRS Pinholin Interact Differently with the Lipid Bilayer: A 31P and 2H Solid State NMR Study

Daniel L. Drew Jr., Gunjan Dixit, Brandon Butcher, Indra D. Sahu, Gary A. Lorigan*

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

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4.1 Abstract: Pinholins are a family of lytic membrane proteins responsible for the lysis of the cytosolic membrane in host cells of double stranded DNA bacteriophages. Protein-lipid interactions have been shown to influence membrane protein topology as well as their function. This work investigated the interactions of pinholin with the phospholipid bilayer while in an active and inactive confirmation to elucidate the different interactions the two forms have with the bilayer. Pinholin incorporated into deuterated DMPC-d54 lipid bilayers, along with 31P and 2H solid state NMR (SS-NMR) spectroscopy were used to probe the protein-lipid interactions with the phosphorus head group at the surface of the bilayer while interactions with the 2H nuclei were used to study the hydrophobic core. A comparison of the 31P chemical shift anisotropy (CSA) values of the active S2168 pinholin and inactive S21IRS pinholin indicated stronger head group interactions for the pinholin in its active form when compared to that of the inactive form supporting the model of a partially externalized peripheral transmembrane domain (TMD). The 2H quadrupolar splitting analysis showed a decrease in spectral width for both forms of the pinholin when compared to the empty bilayers at all temperatures. In this case the decrease in the spectral width of the inactive S21IRS form of the pinholin showed stronger interactions with the acyl chains of the bilayer. The presence of the inactive form’s additional TMD within the membrane was supported by the loss of peak resolution observed in the 2H 2 NMR spectra. The order parameters (SCD) were calculated from dePaked H SS-NMR spectra and further supported the interaction of active pinholin S2168 partially externalized TMD with the surface of the bilayer. This study also highlights the versatility and application of SS-NMR to differentiate between two different conformations of the same protein.

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4.2 Introduction: The initial step of double-stranded DNA bacteriophage host cell lysis is the permeabilization of the cytosolic membrane allowing for the release of the muralytic endolysin enzyme to begin peptidoglycan degradation. This is accomplished by a family of membrane proteins known as holins.1, 2 There are many subfamilies of holins, the most well studied being the λ S105 canonical holin which has three transmembrane helices and forms a single micron-scale hole in the membrane.3-5 This allows for the release fully folded and functional endolysin from the cytoplasm.6 This study will focus on the less studied lambdoid bacteriophage ϕ21 pinholin, named such due to the numerous nanometer-scale holes it forms throughout the membrane. Pinholin is composed of two amphipathic helical transmembrane domains and, due to a dual translational start motif in the S21 gene, is expressed as an active S2168 pinholin and a S2171 antipinholin.7, 8 This S2171 antipinholin is the negative-dominant form of the pinholin and is responsible for slowing the timing of membrane lysis. The pinholin pathway begins with the harmless accumulation of both forms of the pinholin in the cytosolic membrane where nonfunctional S2168 homodimers and S2168:S2171 heterodimers begin to form at a 2:1 ratio.9 These dimers require the externalization of the first transmembrane helical domain (TMD1) from the membrane to become functional.10 Due to the additional positively charged lysine found on the N-termini of the S2171 antipinholin the externalization of TMD1 in this case is slowed which delays the timing of lysis. Understanding the differences between these two forms of the pinholin and their interactions with the lipid bilayer will help to gain a better insight into the beginning steps of the bacteriophage lytic pathway. This biophysical work will probe the different protein-lipid interactions pinholin has with the membrane bilayer while in the inactive conformation, before TMD1 externalization, and in the active conformation with TMD1 externalized. This system will be studied utilizing 31P and 2H solid state nuclear magnetic resonance (SS-NMR) spectroscopy. SS-NMR spectroscopy is a powerful biophysical technique that has been widely used to study the structure, topology, and dynamics of membrane proteins as well as their interactions with the lipid bilayer.11, 12 These protein-lipid interactions have been shown to be critical in several biological processes and have been known to impact the aggregation or segregation of protein within the membrane, the overall function of the protein, and the

64 association of lytic proteins to the membrane.13-16 Incorporation of proteins into synthetic phospholipid bilayers yields a more relevant biological simulation of protein-lipid interactions than monolayers or detergent micelles.17-19 The effect of pinholin on the hydrophilic phospholipid headgroups and hydrophobic acyl chains of these bilayers can be probed through SS-NMR spectroscopy and incorporation of pinholin into deuterated 31 DMPC (d54-DMPC) liposomes. P SS-NMR spectral line shapes are sensitive to the local environment around the 31P head group and can give insight into the overall lipid dynamics within the system, and lipid phases. The 31P nuclei of the phosphocholine headgroup of DMPC can reveal different dynamic effects between active and inactive pinholin at the surface of the membrane.20-22 2H SS-NMR spectroscopy of deuterated lipid acyl chains can be used to obtain insight into the dynamics and chain order or packing of the acyl chains within the hydrophobic core of the membrane.20, 23, 24 The use of both NMR active nuclei can be combined to determine the interactions of the pinholin with both the lipid head groups and the hydrophobic core of the membrane and how those interactions differ when the pinholin is in its active or inactive conformation. This study focuses on the different protein-lipid interactions between active and inactive pinholin with respect to TMD1’s partial externalization from the membrane, the influence of the remaining TMDs on the packing of the acyl chains within the membrane, and the effects of both pinholin concentration and temperature changes on the properties of the DMPC lipid bilayer. This study also highlights the versatility and application of the biophysical technique SS-NMR to differentiate between two different conformations of the same protein.

4.3 Materials and Methods:

4.3.1 Solid Phase Peptide Synthesis All pinholin proteins were synthesized on a CEM Liberty Blue Solid Phase Peptide Synthesizer with a Discover Bio Microwave System. The solid phase was a NovaSyn TG amino resin, a composite of cross-linked polystyrene with the PEG chains terminally functionalized with the first amino acid group of the pinholin sequence. All Fmoc protected amino acids, as well as the activator diisopropylcarbidimide (DIC), and activator base

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Oxyma, were purchased from Millipore Sigma. Amino acid solutions were prepared at a 0.2 M concentration and coupled using DIC and oxyma at 90°C for 4 min. Fmoc deprotection was run with 20% piperidine in DMF at 93°C for 1 min.25 The resin and side chain protecting groups were cleaved from the protein using a 30 mL, three-hour Trifluoroacetic acid (TFA) cleavage reaction, [94% TFA, 2.5 TIPs, 2.5% EDT, 1% 26-28 water]. The TFA was evaporated off with a N2 gas flow and the crude pinholin was precipitated from the remaining solution using tert butyl ether.

4.3.2 Protein Purification The crude pinholin peptide was purified using reverse phase high pressure liquid chromatography (RP-HPLC) on a C4 column and was eluted using a two-solvent gradient. The first solvent was deionized water, the second was 90% HPLC grade acetonitrile. Both solvents were degassed and then acidified with 0.1% TFA by volume. The pinholin protein was collected in fractions and the molecular weight of the protein was confirmed using Matrix Assisted Laser Desorption Ionization – Time of Flight Spectrometry (MALDI-TOF).28 Collected fractions were dried using lyophilization to recover the purified pinholin.

4.3.3 Proteoliposome Sample Preparation The active S2168 and inactive S21IRS wild type pinholin were incorporated into 1,2-

Dimyristoyl – d54 – sn – Glycero – 3 – Phosphocholine (DMPC) multilamellar vesicles (MLV) which have been shown to be successful at mimicking a lipid bilayer for membrane 29, 30 protein studies. DMPC-d54 was purchased from Avanti Polar Lipids. These MLVs were created by dissolving a known amount of pinholin in 2,2,2-Trifluoroethanol (TFE) and adding the protein to DMPC dissolved in chloroform at a protein concentration at either 1 mol% or 2 mol%. The solvents were evaporated off using inert N2 gas and the remaining lipid/protein film was rehydrated using 4-(2-hydroxyethyl)-1- piperazineethanesulfonic acid (HEPES) buffer created with deuterium depleted water at a concentration of 10 mM and adjusted to a neutral pH of ~7.0. All samples were rehydrated with the HEPES buffer to a final lipid concentration of 50 mM. To improve

66 incorporation and MLV formation the protein/lipid solution was flash frozen in liquid nitrogen and then sonicated. This freeze-thaw cycle was repeated 3 times.

4.3.4 Solid State Nuclear Magnetic Resonance Spectroscopy The solid-state nuclear magnetic resonance spectroscopy measurements were conducted using a Bruker 500 MHz WB UltraShield NMR spectrometer with a 4 mm triple resonance CP-MAS probe. 31P NMR spectra were recorded with 1H decoupling using a 4 µs π/2 pulse and a 4 s recycle delay, a spectral width of 300 ppm, and by averaging 6K scans. The collected free induction decay was processed using 300 Hz of line broadening. 2H NMR spectra were collected at 76.77 MHz using a standard quadrupolar echo pulse sequence (3 µs 90° pulse length, 40 µs inter-pulse delay with a 0.5 sec recycle delay).31 The spectral width was set to 100 kHz and 80 K transients were averaged for every 2H NMR spectra. Exponential line broadening of 100 Hz was applied to the free induction decay before the Fourier transformation was taken. Both 31P and 2H SS-NMR experiments were collected from 25°C to 55°C in 10°C increments and the sample was left to equilibrate to each temperature for 5 min before data acquisition. Depaking of 2H SS-NMR spectra was conducted using MATLAB with the dePaking script published and provided by the Brown group at the University of Arizona.11, 12 The dePaked NMR spectra were calculated such that the bilayer normal was perpendicular ( = 0) with respect to the static magnetic field.32 The dePaked peak picking and quadrupolar splitting values were determined by plotting the dePaked spectra using Igor Pro. The quadrupolar splitting of each doublet pair corresponds to the deuterium atoms bonded to a different carbon on the lipid acyl chain. The peak of the three 2H nuclei bound to the terminal methyl carbon appear as the doublet closest to 0 kHz and is assigned as carbon number 14. The remaining carbon number assignments were made in decreasing order as the quadrupolar splitting values for the respective 2H atoms increased. The quadrupolar splitting for the 2H atoms on the carbons closest to the glycerol backbone appear as a plateau and were estimated by integrating the last broad peak. Order parameters for 2H atoms on each carbon were calculated using the quadrupolar splitting values and the following equation.33

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3 푒2푞푄 ∆푣 = ( ) ∙ 푆 [1] 푄 4 ℎ 퐶퐷

2 푒 푞푄⁄ Where ∆푣푄 is the quadrupolar splitting value, ( ℎ) is the deuterium quadrupolar coupling constant which is equal to 167 kHz for all C-D covalent bonds, and 푆퐶퐷 is the order parameter for the deuterium on the given carbon.

4.4 Results and Discussion: The initial pinholin work showed the successful synthesis, purification, and incorporation of pinholin into membrane mimetic systems.28 This work utilizes 31P and 2H SS-NMR to investigate the effects of the active S2168 and inactive S21IRS pinholin on the dynamic properties of the phospholipid bilayer. The lipid headgroups and acyl chains of 31 DMPC-d54 multi lamellar vesicles (MLVs) were studied using P chemical shift anisotropy

2 (CSA) and H quadrupolar splitting (∆푣푄), respectively. Figures 4.1A and 4.1B shows a schematic representation of the active and inactive forms of pinholin within the membrane while the chemical structure of d54-DMPC can be seen in Figure 4.1C.

4.4.1 31P SS-NMR CSA Analysis for the Active and Inactive Forms of Pinholin To investigate the interactions of the active S2168 and inactive S21IRS forms of pinholin with the headgroups of the DMPC membrane 31P SS-NMR static spectra were taken at increasing protein concentrations (0 – 2 mol%). The 31P NMR static spectra at varying mol% for active S2168 pinholin are shown in Figure 4.2 with increasing temperatures from 25°C to 55°C at 10°C intervals. All motionally averaged 31P powder- patter spectra are characteristic of axially symmetric (휎11 ≅ 휎22 ≠ 휎33) phospholipid bilayers, in this case multi lamellar vesicles, in the liquid crystalline phase (Lα) for all mol% and temperature variants. The chemical shift anisotropy (CSA) for each spectrum was calculated by measuring the difference between 11 (휎11 ≅ 휎22) and 휎33 of the spectra with all 31P CSA values in the range of 49.3 – 35.3 ppm (Table 1).34, 35 For both the 1 and 2 mol% cases of active S2168 and inactive S21IRS, the 31P CSA width decreases as the temperature increases. CSA for the active S2168 decreased from 45.2 ppm at 25°C to 42.6 at 55°C for 1 mol% and from 40.7 ppm to 35.3 ppm for 2 mol%

68 over the same temperature range. The inactive S21IRS pinholin showed a similar indirect trend between temperature and 31P CSA width with the 1 mol% inactive decreasing from 46.6 ppm at 25°C to 43.3 ppm at 55°C and 2 mol% decreasing from 44.2 ppm to 39.1 ppm over the same temperature range (Figure 4.3). A comparison of the 31P CSA width between 0, 1, and 2 mol% S2168 and S21IRS pinholin is shown in Figure 4.4. All data presented in Figure 4.4 was collected at 35°C to ensure the DMPC proteoliposomes are in the liquid crystalline phase. The 31P CSA width is observed to decrease as the mol% of the protein increases for both the active and inactive pinholin. At 35°C the 0 mol% MLVs have a CSA of 47.3 ppm, the active S2168 CSA decreased from 44.5 ppm to 38.4 ppm, while the inactive S21IRS decreased from 46.0 ppm to 43.7 ppm. The overall 31P CSA width comparison between active S2168 and inactive S21IRS at 1 and 2 mol% at each temperature reveals in each case that the active S2168 pinholin has a lower CSA width value than the inactive S21IRS indicating a higher degree of interaction with the lipid headgroup, when compared to the inactive S21IRS. This trend would be consistent with a partially externalized TMD1 of the active pinholin form laying on the surface of the membrane while TMD1 from the inactive form is still incorporated within the membrane. Additionally, each spectrum in Figures 4.2 and 4.3 show the presence of an isotropic peak near 0 ppm. This peak in the 31P NMR static spectra is indicative of the fast-relative motion or reorientation of the phospholipid headgroup of DMPC with respect to the 31P NMR timescale. This peak can appear from the presence of lipids tumbling quickly in solution, from lateral diffusion of the lipid, or displacement of lipids over the surface of the membrane.34, 36 The isotropic linewidth for active and inactive pinholin varies from 1.3 ppm for the 1 mol% active S2168 at 25°C to 8.0 ppm for 2 mol% active S2168 at 55°C. A similar trend to that of the CSA can be seen for the isotropic component of active S2168 and inactive S21IRS pinholin. For each temperature, as the concentration of the protein increases from 1 to 2 mol% the linewidth of the isotropic peak also increases indicating a greater perturbation of the membrane lipid head groups. Just like in the 31P CSA width analysis the active S2168 pinholin shows a greater increase of the isotropic peak when compared to that of than the inactive S21IRS pinholin. This would be consistent with the overall function of the active pinholin as it is a membrane lysing protein. The

69 observed trends of the isotropic peak are indicative of the pinholin’s disruption of the lipid bilayer or displacement of the lipids in the bilayer from the partial externalization of TMD1.

4.4.2 2H SS-NMR Quadrupolar Splitting for the Active and Inactive Forms of Pinholin The effect of active and inactive pinholin on the acyl chain dynamics and overall order within the DMPC bilayer were studied using 2H SS-NMR and measuring the 2 corresponding quadrupolar splittings (∆푣푄) and order parameters (SCD). The H NMR spectra of 0, 1, and 2 mol% active pinholin incorporated into DMPC-d54 MLVs at varying temperatures are shown in Figure 4.5. The 2H NMR spectra shown in Figure 4.5 are characteristic of axially symmetric motions for the phospholipids about the membrane normal. The overall 2H NMR spectra are composed of a series of overlaying doublet resonances originating from the twelve different CD2 positions of DMPC with the highest intensity central doublet corresponding to the terminal CD3 methyl group of the acyl chain.37, 38 The spectral width for the empty MLVs, active, and inactive pinholin at 1 and 2 mol% can be seen in Table 2 and range from 29.8 – 19.2 kHz. The decrease in spectral width when compared to the 0 mol% sample suggests the presence of both active S2168 and inactive S21IRS pinholin transmembrane domain interactions with the acyl chains of the DMPC MLVs. The spectral width for the active S2168 pinholin ranged from 29.3 kHz at 25°C to 21.6 kHz at 55°C for 1 mol%, while the spectra for the 2 mol% decreased from 28.5 kHz to 19.8 kHz over the same temperature range. The inactive S21IRS sample showed a similar temperature trend between the spectral widths as the 1 mol% inactive S21IRS decreased from 28.9 kHz at 25°C to 21.1 kHz at 55°C while the 2 mol% ranged from 27.9 kHz to 19.2 kHz over the same temperatures. A comparison of the spectral width as active S2168 and inactive S21IRS pinholin concentrations are increased from 1 to 2 mol% at 35°C is shown in Figure 4.6. Active S2168 and inactive S21IRS pinholin both show a decrease in the spectral width as the concentration of the protein increases from 1 to 2 mol%. Unlike in the 31P CSA analysis, the inactive S21IRS shows a greater decrease in the spectral width when compared to that of the active S2168 at the same concentration and temperature.

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Additionally, a loss of spectral resolution is apparent in Figures 4.5 and 4.6 as the concentration of pinholin increases, as seen by the absence of the individual sharp doublet peaks throughout each 2H NMR spectrum. The loss in the resolution of each 2H spectrum along with the changes in overall spectral width, is a result of the interactions of the active S2168 and inactive S21IRS pinholin packing against the acyl chains of the lipid bilayer.21, 39, 40 The comparison of 1 mol% active S2168 and 1 mol% inactive S21IRS seen in Figure 4.6 shows a greater loss in the peak resolution for the inactive form of pinholin, when compared to the active pinholin at the same concentration. This suggests a stronger interaction of the inactive S21IRS pinholin with the acyl chains of the lipid bilayer when compared to that of the pinholin in the active conformation. The greater loss of resolution seen in the 1 mol% inactive S21IRS would indicate a greater number of TMDs present within the bilayer. This is consistent with trends obtained in the quadrupolar splitting analysis and provides further evidence towards the externalization of active S2168 pinholin TMD1 from the membrane yielding fewer acyl chain interactions when compared to inactive S21IRS pinholin. Figure 4.5 also shows an appearance of an isotropic peak centered around 0 kHz which increases as the concentration of active S2168 pinholin increases. This isotropic component is indicative of the fragmentation of the larger MLVs into small sized vesicles which have a fast-relative motion with respect to the NMR timescale. The trends observed in Figure 4.5 show a direct correlation between the intensity of the isotropic component and the concentration of active S2168 pinholin in the DMPC bilayers. These results are consistent with the 31P SS-NMR data which both show an increase in the isotropic component as a function of the concentration of pinholin but show a higher degree of membrane disruption for active S2168 when compared to that of the inactive S21IRS pinholin.

4.4.3 Depaking and Order Parameters (SCD) In order to further explore the dynamic interactions of the active and inactive forms of pinholin with the phospholipid bilayer, order parameters (SCD) were calculated from the 2H SS-NMR data. The packing of DMPC acyl chains or order parameters defines the dynamic perturbations or local packing of each individual C–D bond of the standard

71

DMPC acyl chain conformations as the concentration of active and inactive pinholin increases within the bilayer. These order parameters were calculated using equation [1] where the quadrupolar splitting values were determined through dePaking of the original 2H SS-NMR spectra shown in Figure 4.5. Due to the nature of calculating order parameters large changes in the quadrupolar splitting of peaks in the dePaked spectra results in small shifts in the order parameter plot. An illustrative example of dePaking for empty 0 mol% MLVs and 1 mol% active S2168 is seen in Figure 4.7 and shows the original and dePaked spectra for both cases. While the decrease in spectral width and increase in the isotropic component are consistent with expected trends, the order parameters for both 2 mol% active S2168 pinholin and 1 mol% inactive S21IRS pinholin could not be calculated due to the loss in resolution of the original spectra that was discussed in the previous section. The dePaked spectra for all active S2168 samples can be seen in Supplemental Figures 4.S1. Order parameter values ranged from the most disordered,

0.015 for the CD3 terminal methyl, to the most ordered, 0.23 for the CD2 closest to the glycerol backbone, all of which are characteristic values of DMPC bilayers in the liquid- crystalline phase. The trends of the order parameters at varying temperatures for empty 0 mol% MLVs and 1 mol% active S2168 pinholin show a decrease in the overall order of the system as the temperature increases. The overall decrease in order parameters for each CD2 moving further away from the glycerol backbone can be seen in Figure 4.8 for both 0 mol% and 1 mol% active S2168 at all temperatures. This profile is characteristic of an increase in mobility, or decrease in order, of the acyl chains both of which would be true of a system at higher temperatures. To better see the differences between order parameters an overlaid comparison of the order parameters between 0 mol% and 1 mol% active S2168 pinholin at 35°C can be seen in Figure 4.9. A greater separation between the order parameters of the carbons near the glycerol backbone indicate more disorder in the presence of 1 mol% active S2168. The overlaid dePaked spectra used to find quadrupolar splitting values of 0 mol% and 1 mol% active S2168 at 35°C can be seen in the Supplementary Information (Figure 4.S2) which highlights the shift observed for each peak throughout the acyl chain. The trends observed in Figure 4.9 are consistent with all the previous data shown and support the partial externalization of active S2168 TMD1

72 from the membrane as the helix interacts with the carbons closest to the membrane surface. Alternatively, an interesting trend was observed for the order parameters of active S2168 pinholin near the phase transition (25°C) of the DMPC bilayer. A comparison of the order parameters between 0 mol% and 1 mol% active S2168 pinholin at 25°C can be seen in Figure 4.10. This shows more disorder for the carbons closer to the glycerol backbone in the presence of 1 mol% active S2168. The overlaid dePaked spectra used to find quadrupolar splitting values of 0 mol% and 1 mol% active S2168 at 25°C can be seen in the Supplementary Information (Figure 4.S3) which highlights the shifts observed for each peak throughout the acyl chain. Interestingly, at temperatures near the phase transition of DMPC the order parameters after carbon-7 indicate a more ordered environment in the presence of active S2168 pinholin when compared to the empty DMPC liposomes. This is due to the lipid compensation for a positive hydrophobic mismatch occurring in the DMPC bilayer. For a typical alpha helix each helical turn comprises of 3.6 amino acids, correlating to each amino acid contributing 1.5 Å to the helix length. This indicates the TMDs of pinholin are both >33 Å in length. The average hydrophobic thickness for a DMPC bilayer is ~25Å. This results in residues of the helix intended to be located within the lipid bilayer instead residing outside of the membrane bilayer. This is known as a positive hydrophobic mismatch.41-43 This mismatch can be resolved through an ordering of the acyl chains packing around the transmembrane helix resulting in an increase in the local hydrophobic thickness of the bilayer.41, 42 In this case, the higher order parameters seen after carbon- 7 in Figure 4.10 would be indicative of this higher degree of ordering of the acyl chains. Future studies could be conducted using longer chained lipids to avoid the presence of this mismatch at lower temperatures.

4.5 Conclusion: Utilizing 31P and 2H SS-NMR spectroscopy, this study compares the interactions of phospholipid bilayers with the active S2168 and inactive S21IRS forms of pinholin. 31P SS-NMR nuclei were used to probe the interaction of pinholin with the lipid headgroups. The 31P CSA width values of 1 and 2 mol% active S2168 are smaller than the inactive

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S21IRS CSA values at the same mol% and temperature. The decrease in CSA indicates a higher degree of perturbations at the surface of the membrane for the active S2168 pinholin than the inactive S21IRS form of pinholin. This is consistent with the previously proposed model of TMD1 partially externalizing from the lipid bilayer. The 31P SS-NMR spectra also show an isotropic peak with an increasing line width as the concentration of pinholin increases. A greater linewidth for the active over the inactive pinholin could be a result of displacement of the lipids in the bilayer coming from the partial externalization of TMD1 or possibly due to the lysing function of the active S2168 pinholin which would induce a greater amount of disruption of the lipid bilayer The 2H SS-NMR quadrupolar splitting data showed an overall decrease in the spectral width as the mol% of pinholin increased. Unlike 31P CSA trends, the inactive S21IRS showed a greater decrease in the 2H quadrupolar splitting. This trend indicates a greater degree of interaction with the acyl chains of the lipid coming from the inactive form of the pinholin. These interactions are originating from the presence of TMD1 remaining within the membrane for the inactive form of the pinholin. This is further supported by the additional loss in resolution of the doublet peaks observed when comparing 1 mol% inactive S21IRS to the spectra for 1 mol% active S2168 pinholin. The order parameters determined from the quadrupolar splitting of the dePaked 2H spectra give further support for the conclusions drawn from the 31P and 2H data. The decrease in the order parameters of carbon 2-6 in the presence of 1 mol% active S2168 pinholin support the idea of a partial TMD1 externalization from the membrane. This study also highlights the application of SS-NMR spectroscopy to distinguish between the two different conformations of the pinholin protein and the resulting interactions with the lipid bilayer

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4.6 Acknowledgements: We would like to thank Dr. Theresa Ramelot for her support for the SS-NMR instrument and data processing. The MATLAB software code used to dePake our 2H spectrum was kindly provided by the Brown group at University of Arizona. We are also grateful to the members of the Young group at Texas A&M University for their experimental suggestions. This work was generously supported by a NSF CHE-1807131 grant and a NIGMS/NIH Maximizing Investigator’s Research Award (MIRA) R35 GM126935 award. Gary A. Lorigan would also like to acknowledge support from the John W. Steube Professorship.

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4.7 Figures:

Figure 4.1: A) The currently proposed model of the active S2168 pinholin with TMD1 partially externalized from the membrane. B) The inactive S21IRS with the additional 5 amino acids seen on the N-terminus of the protein. C) Chemical structure of deuterated

DMPC-d54 lipid.

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31 Figure 4.2: Temperature dependent P SS-NMR spectra of empty d54-DMPC MLVs (A), 1 mol% active S2168 pinholin (B), and 2 mol% active S2168 pinholin (C).

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Figure 4.3: Temperature dependent 31P NMR spectra of 1 mol% inactive S21IRS pinholin (A), and 2 mol% inactive S21IRS pinholin (B).

78

Figure 4.4: Comparison of chemical shift anisotropy (CSA) width at 35°C between empty 21 21 DMPC-d54 MLVs, active S 68, and inactive S IRS pinholin for 1 and 2 mol% pinholin.

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31P CSA Width (ppm)

25°C 35°C 45°C 55°C

0 mol% 48.9 47.3 46.5 43.8

1 mol% S2168 45.2 44.5 43.8 42.6

2 mol% S2168 40.7 38.4 36.8 35.3

1 mol% S21IRS 46.6 46.0 45.2 43.3

2 mol% S21IRS 44.2 43.7 42.9 39.1

Table 4.1: The 31P NMR chemical shift anisotropy width (± 0.5 ppm) of empty 0 mol% MLVs, Active S2168 pinholin and Inactive S21IRS pinholin at 1 and 2 mol%.

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2 Figure 4.5: Temperature dependent H SS-NMR spectra of empty DMPC-d54 MLVs (A), 1 mol% active S2168 pinholin (B), and 2 mol% active S2168 pinholin (C).

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Figure 4.6: Comparison of the 2H quadrupolar splitting at 35°C between empty DMPC- 21 21 d54 liposomes, active S 68, and inactive S IRS pinholin at 1 and 2 mol% pinholin.

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2H Quad. Splitting (kHz)

25°C 35°C 45°C 55°C

0 mol% 29.8 26.5 24.2 22.6

1 mol% S2168 29.3 25.6 23.4 21.6

2 mol% S2168 28.5 23.6 20.5 19.8

1 mol% S21IRS 28.9 24.9 22.9 21.1

2 mol% S21IRS 27.9 22.9 20.0 19.2

Table 4.2: The 2H quadrupolar splitting spectral width analysis of empty d54-DMPC MLVs, active S2168, and inactive S21IRS pinholin at one and two mol% for increasing temperatures.

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Figure 4.7: (A) The original 2H SS-NMR spectra of empty 0 mol% MLVs at 25°C. (B) The resulting dePaked spectra for empty 0 mol% MLVs. (C and D) The original 2H SS-NMR spectra of 1 mol% active S2168 pinholin at 25°C and the resulting dePaked spectra, respectively.

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Figure 4.8: A) Order Parameters (SCD) for 0 mol% MLVs at increasing temperatures. B) 21 Order Parameters (SCD) for 1 mol% active S 68 pinholin at increasing temperatures.

85

Figure 4.9: Order parameter (SCD) comparison at 35°C between empty 0 mol% MLVs (black) and 1 mol% active S2168 pinholin (blue).

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Figure 4.10: Order parameter (SCD) comparison at 25°C between empty 0 mol% MLVs (black) and 1 mol% active S2168 pinholin (blue). The order parameters after carbon-7 for 1 mol% active S2168 show more order than the 0 mol% sample to compensate for the positive hydrophobic mismatch of the bilayer.

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Supplementary Figures:

Figure 4.S1: The dePaked 2H spectra of 0 mol% (black), 1 mol% S2168 (blue), and 2 mol% S2168 (green) at each temperature overlaid on top of the original 2H spectra. The loss in peak resolution at 2 mol% S2168 prevents the calculation of order parameters with any degree of certainty.

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Figure 4.S2: The overlaid dePaked spectra of 0 mol% and 1 mol% active S2168 pinholin at 35°C highlighting the shifts in CD2 quadrupolar splitting peaks.

89

Figure 4.S3: The overlaid dePaked spectra of 0 mol% and 1 mol% active S2168 pinholin at 25°C. The positive hydrophobic mismatch is resolved through the ordering of the CD2 near the tail of the acyl chain observed as leftward shifts in the peaks from 0 to 1 mol% and result in higher order parameter values.

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References:

1. Young, R., Phage Lysis: Do we have the hole story yet? Curr Opin Microbiol. 2013, 16 (6), 1-8. 2. Young, R., Phage Lysis: Three Steps, Three Choices, One Outcome. Journal of Microbiology 2014, 52 (3), 243-258. 3. To, K. H.; Young, R., Probing the Structure of the S105 Hole. Journal of Bacteriology 2014, 196 (21), 3683-3689. 4. Dewey, J. S.; Savva, C. G.; White, R. L.; Vitha, S.; Holzenburg, A.; Young, R., Micron-scale holes terminate the phage infection cycle. Proceedings of the National Academy of Sciences of the United States of America 2010, 107 (5), 2219-2223. 5. Wang, I. N.; Deaton, J.; Young, R., Sizing the holin lesion with an endolysin-beta- galactosidase fusion. Journal of Bacteriology 2003, 185 (3), 779-787. 6. Park, T.; Struck, D. K.; Dankenbring, C. A.; Young, R., The pinholin of lambdoid phage 21: Control of lysis by membrane depolarization. Journal of Bacteriology 2007, 189 (24), 9135-9139. 7. 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. 8. Bonovich, M. T.; Young, R., DUAL START MOTIF IN 2 LAMBDOID S-GENES UNRELATED TO LAMBDA-S. Journal of Bacteriology 1991, 173 (9), 2897-2905. 9. Pang, T.; Park, T.; Young, R., Mapping the pinhole formation pathway of S21. Molecular Microbiology 2010, 78 (3), 710-719. 10. Park, T.; Struck, D. K.; Deaton, J. F.; Young, R., Topological dynamics of holins in programmed bacterial lysis. Proceedings of the National Academy of Sciences of the United States of America 2006, 103 (52), 19713-19718. 11. Kinnun, J. J.; Leftin, A.; Brown, M. F., Solid-State NMR Spectroscopy for the Physical Chemistry Laboratory. Journal of Chemical Education 2013, 90 (1), 123-128. 12. Molugu, T. R.; Lee, S.; Brown, M. F., Concepts and Methods of Solid-State NMR Spectroscopy Applied to Biomembranes. Chemical Reviews 2017, 117 (19), 12087- 12132. 13. Arora, A.; Tamm, L. K., Biophysical approaches to membrane protein structure determination. Current Opinion in Structural Biology 2001, 11 (5), 540-547. 14. Dempsey, C. E.; Ryba, N. J. P.; Watts, A., Evidence From Deuterium Nuclear- Magnetic-Resonance For The Temperature-Dependent Reversible Self-Association Of Erythrocyte Band-3 In Dimyristoylphosphatidylcholine Bilayers. Biochemistry 1986, 25 (8), 2180-2187. 15. Lemmon, M. A.; Engelman, D. M., Specificity And Promiscuity In Membrane Helix Interactions. Febs Letters 1994, 346 (1), 17-20. 16. Watts, A., Protein-Lipid Interactions - Do The Spectroscopists Now Agree. Nature 1981, 294 (5841), 512-513. 17. Morrow, M. R.; Grant, C. W. M., The EGF transmembrane domain: Peptide-peptide interactions in fluid bilayer membranes. Biophysical Journal 2000, 79 (4), 2024-2032. 18. Rigby, A. C.; Barber, K. R.; Shaw, G. S.; Grant, C. W. M., Transmembrane region of the epidermal growth factor receptor: Behavior and interactions via H-2 NMR. Biochemistry 1996, 35 (38), 12591-12601.

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19. Sharpe, S.; Barber, K. R.; Grant, C. W. M.; Goodyear, D.; Morrow, M. R., Organization of model helical peptides in lipid bilayers: Insight into the behavior of single- span protein transmembrane domains. Biophysical Journal 2002, 83 (1), 345-358. 20. Dave, P. C.; Tiburu, E. K.; Damodaran, K.; Lorigan, G. A., Investigating structural changes in the lipid bilayer upon insertion of the transmembrane domain of the membrane-bound protein phospholamban utilizing P-31 and H-2 solid-state NMR spectroscopy. Biophysical Journal 2004, 86 (3), 1564-1573. 21. Abu-Baker, S.; Lorigan, G. A., Phospholamban and its phosphorylated form interact differently with lipid bilayers: A (31)P, (2)H, and (13)C solid-state NMR spectroscopic study. Biochemistry 2006, 45 (44), 13312-13322. 22. Santos, J. S.; Lee, D. K.; Ramamoorthy, A., Effects of antidepressants on the conformation of phospholipid headgroups studied by solid-state NMR. Magnetic Resonance in Chemistry 2004, 42 (2), 105-114. 23. Koenig, B. W.; Ferretti, J. A.; Gawrisch, K., Site-specific deuterium order parameters and membrane-bound behavior of a peptide fragment from the intracellular domain of HIV-1 gp41. Biochemistry 1999, 38 (19), 6327-6334. 24. Yamaguchi, S.; Huster, D.; Waring, A.; Lehrer, R. I.; Kearney, W.; Tack, B. F.; Hong, M., Orientation and dynamics of an antimicrobial peptide in the lipid bilayer by solid- state NMR spectroscopy. Biophysical Journal 2001, 81 (4), 2203-2214. 25. Hansen, P. R.; Oddo, A., Fmoc Solid-Phase Peptide Synthesis. Peptide Antibodies: Methods and Protocols 2015, 1348, 33-50. 26. Lloyd-Williams, P.; Albericio, F.; Giralt, E., Chemical Approaches to the Synthesis of Peptides and Proteins. CRC Press: 1997; p 304. 27. King, D. S.; Fields, C. G.; Fields, G. B., A Cleavage Method Which Minimizes Side Reactions Following Fmoc Solid-Phase Peptide-Synthesis. International Journal of Peptide and Protein Research 1990, 36 (3), 255-266. 28. Drew, D. L.; 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 S-21 pinholin protein. Analytical Biochemistry 2019, 567, 14-20. 29. Sahu, I. D.; McCarrick, R. M.; Lorigan, G. A., Use of Electron Paramagnetic Resonance To Solve Biochemical Problems. Biochemistry 2013, 52 (35), 5967-5984. 30. Dixit, G.; Sahu, I. D.; Reynolds, W. D.; Wadsworth, T. M.; Harding, B. D.; Jaycox, C. K.; Dabney-Smith, C.; Sanders, C. R.; Lorigan, G. A., Probing the Dynamics and Structural Topology of the Reconstituted Human KCNQ1 Voltage Sensor Domain (Q1- VSD) in Lipid Bilayers Using Electron Paramagnetic Resonance Spectroscopy. Biochemistry 2019, 58 (7), 965-973. 31. Davis, J. H.; Jeffrey, K. R.; Bloom, M.; Valic, M. I.; Higgs, T. P., Quadrupolar Echo Deuteron Magnetic-Resonance Spectroscopy In Ordered Hydrocarbon Chains. Chemical Physics Letters 1976, 42 (2), 390-394. 32. Molugu, T. R.; Xu, X.; Leftin, A.; Lope-Piedraffita, S.; Martinez, G. V.; Petrache, H. I.; Brown, M. F., Solid-State NMR Spectroscopy of Membranes. Modern Magnetic Resonance: 2017. 33. Drechsler, A.; Anderluh, G.; Norton, R. S.; Separovic, F., Solid-state NMR study of membrane interactions of the pore-forming cytolysin, equinatoxin II. Biochimica Et Biophysica Acta-Biomembranes 2010, 1798 (2), 244-251.

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34. Seelig, J., P-31 Nuclear Magnetic-Resonance And Head Group Structure Of Phospholipids In Membranes. Biochimica Et Biophysica Acta 1978, 515 (2), 105-140. 35. McLaughlin, A. C.; Cullis, P. R.; Berden, J. A.; Richards, R. E., P-31 NMR OF PHOSPHOLIPID MEMBRANES - EFFECTS OF CHEMICAL-SHIFT ANISOTROPY AT HIGH MAGNETIC-FIELD STRENGTHS. Journal of Magnetic Resonance 1975, 20 (1), 146-165. 36. Lau, T. L.; Ambroggio, E. E.; Tew, D. J.; Cappai, R.; Masters, C. L.; Fidelio, G. D.; Barnham, K. J.; Separovic, F., -beta peptide disruption of lipid membranes and the effect of metal ions. Journal of Molecular Biology 2006, 356 (3), 759-770. 37. Seelig, J., DEUTERIUM MAGNETIC-RESONANCE - THEORY AND APPLICATION TO LIPID-MEMBRANES. Quarterly Reviews of 1977, 10 (3), 353-418. 38. Lafleur, M.; Bloom, M.; Cullis, P. R., Lipid Polymorphism And Hydrocarbon Order. Biochemistry and Cell Biology-Biochimie Et Biologie Cellulaire 1990, 68 (1), 1-8. 39. Minto, R. E.; Adhikari, P. R.; Lorigan, G. A., A H-2 solid-state NMR spectroscopic investigation of biomimetic bicelles containing cholesterol and polyunsaturated phosphatidylcholine. Chemistry and Physics of Lipids 2004, 132 (1), 55-64. 40. Huster, D., Solid-state NMR spectroscopy to study protein lipid interactions. Biochimica Et Biophysica Acta-Molecular and Cell Biology of Lipids 2014, 1841 (8), 1146- 1160. 41. Kandasamy, S. K.; Larson, R. G., Molecular dynamics simulations of model trans- membrane peptides in lipid bilayers: A systematic investigation of hydrophobic mismatch. Biophysical Journal 2006, 90 (7), 2326-2343. 42. Kim, T.; Lee, K. I.; Morris, P.; Pastor, R. W.; Andersen, O. S.; Im, W., Influence of Hydrophobic Mismatch on Structures and Dynamics of Gramicidin A and Lipid Bilayers. Biophysical Journal 2012, 102 (7), 1551-1560. 43. Muhle-Goll, C.; Hoffmann, S.; Afonin, S.; Grage, S. L.; Polyansky, A. A.; Windisch, D.; Zeitler, M.; Burck, J.; Ulrich, A. S., Hydrophobic Matching Controls the Tilt and Stability of the Dimeric Platelet-derived Growth Factor Receptor (PDGFR) beta Transmembrane Segment. Journal of Biological Chemistry 2012, 287 (31), 26178-26186.

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Chapter 5

Pulsed EPR DEER Spectroscopic Study of Bacteriophage S21 Pinholin Reveals Two Different Structural Conformations Between the Active and Inactive Forms

Daniel L. Drew Jr., Tanbir Ahammad, Brandon J. Butcher, Rachel A. Serafin, Indra D. Sahu, Robert M. McCarrick, Gary A. Lorigan*

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

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5.1 Abstract: The mechanism of bacteriophage host cell lysis is a highly regulated process which begins with the lysis of the cytosolic membrane by holin proteins. A newly discovered lysis pathway uncovered a new class of holins known as pinholin. The protein has been shown to have a unique mechanism of lysis in which the pinholin depolarizes the cytosolic membrane leading to the release of signal-anchored-release (SAR) endolysin from the membrane. The initial step of this depolarization pathway is the externalization of the first of pinholin’s two transmembrane domains from the bilayer. This TMD1 externalization step is a critical step in the functional model of pinholin and is required for membrane depolarization to occur. The current structural model of this externalization indicates a complete externalization of TMD1 from the membrane. This study has utilized site directed spin labeling (SDSL) and four-pulse EPR double electron electron resonance (DEER) spectroscopy to probe the structural conformation of the transmembrane domains for the active (S2168) and inactive (S21IRS) forms of pinholin. Multiple spin- labeled pinholin samples were prepared in which one spin label was placed on TMD1 while the other label was positioned on TMD2 and the distances between spin labels were measured. The inactive form of pinholin, which hypothetically has both TMDs incorporated into the membrane, shows short distances between the two transmembrane domains, ranging from 23 ±3 Å to 26 ±3 Å throughout the TMDs. The active form of the pinholin, which is hypothesized to have TMD1 externalized from the membrane, shows longer distances ranging from 25 ±3 Å to 50 ±3 Å. These distances show a progressive increase as the spin label pairs move from the loop region to the end terminus of each helix. These results have led to a revised structural model of the active form of pinholin with TMD1 partially externalized from the bilayer and laying on the membrane surface. This study reports the first direct experimental evidence of the existence of two different structural conformations of pinholin in the membrane.

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5.2 Introduction The lysis of Gram-negative bacterial host cells by double-strand DNA bacteriophages has been revealed to be a carefully regulated process requiring three steps.1, 2 The first of these steps is the lysis of the cytoplasmic membrane by a family of proteins known as holins. Recently a new pathway for lysis was discovered which uses a pinholin to depolarize the cytoplasmic membrane instead of forming large holes.3-5 The depolarization allows for the externalization of a signal-anchored-release (SAR) endolysin protein from the membrane which can then degrade the peptidoglycan. The SAR- endolysin release is responsible for the endolysin degradation and is dependent on the membrane depolarization of the pinholin. Recent studies in the literature have shown the function of pinholin is dependent on the presence of the second to pinholin’s two transmembrane domains (TMD2) within the bilayer.4 Functional models have been proposed in the literature with evidence indicating the externalization of the first pinholin transmembrane domain (TMD1) from the membrane is required to give rise to pinholin function.6 Unfortunately, the different structural conformations between the active and inactive forms of the pinholin within the membrane have not been well characterized. The current structural model between the active and inactive forms of pinholin has TMD1 incorporated in the bilayer when pinholin is in its inactive form and has TMD1 completely externalized from the bilayer when in its active form (Figure 5.1A).7 The model of complete externalization of TMD1 was proposed based on cysteine cross-linking experiments. To overcome the spontaneous externalization of TMD1 from the membrane an additional five amino acids were added to the N-terminus of pinholin (Figure 5.1B) to form a nonlethal form of the protein.4, 8 The active S2168 form of pinholin will be used to probe the structure of pinholin after the externalization of TMD1 from the membrane. This study will probe the structural conformations of the active and inactive forms of pinholin using SDSL coupled with DEER spectroscopy to elucidate the extent of TMD1 externalization from the membrane.

5.3 Experimental Methods DEER measurements were carried out through probing of several doubly spin- labeled active S2168 and inactive S21IRS pinholin proteins. Spin label positions were

96 generated by attaching the nitroxide S-(1-oxyl-2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrol- 3-yl)methyl methanesulfonothioate spin label (MTSL) containing a stable unpaired electron to a cysteine sidechain though a disulfide bond formation. Pinholin is naturally cys-less allowing for experimental control of spin labeling positions through site directed substitution of a specific amino acid to cysteine.9, 10 DEER spectroscopy when coupled with SDSL is a very powerful biophysical technique that can be used to measure distances between 15 to 80 Å in order to study protein structure and conformation.11-16 Previously, the Young group has conducted mutational analysis experiments of the active S2168 and inactive S21IRS forms of the pinholin.6, 8, 17 They indicated that the overall function of pinholin is sensitive to the mutation of certain amino acid positions and that certain positions can initiate the externalization of TMD1 from the membrane for the inactive S21IRS form of pinholin. This study guided our work to choose amino acid positions that were safe for cysteine substitution without affecting the structural/functional relationship of the protein. This was critical in making sure the substitutions would not induce the externalization of TMD1 in the inactive S21IRS form of pinholin .8 This led to the three inactive S21IRS substitution pairs of W27C/A38C, located near the loop region of pinholin, and the G14C/L53C and S8C/L53C positions, located at the terminal ends of each helix. The pairs W27C/A38C and S8C/L53C were also used in the active S2168 pinholin to allow for a direct comparison between the two forms of pinholin. Positions A17C/A38C, A17C/V46C, and S16C/G48C were chosen for the active S2168 form of pinholin as well to determine intermediate distances between the hypothesized shortest and longest DEER pair distances. Each pinholin sample with cysteine substitutions was created using solid phase peptide synthesis (SPPS) and was purified using reverse phase high pressure liquid chromatography (RP-HPLC).18 The amino acid numbering system between S2168 and S21IRS has been established in a previously published work.18 Spin labeling reactions were run in 5x molar excess per cysteine labeling site in DMSO for 24 hours. Upon further purification to remove the excess spin label, pure EPR active pinholin was incorporated into DMPC liposomes at a protein to lipid ratio of 1:1000. 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.6, 17 The four-pulse DEER

97 experiments were conducted at the Ohio Advanced EPR Laboratory 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 coupled with a EN5107D2 resonator. Thus, it was impossible to directly compare modulation depths for the different samples collected at different power levels and pulse widths. DEER samples were run at spin concentrations ~100 µM with 30% (w/w) glycerol added to each sample as a cryoprotectant. 70 µL of sample was loaded into 3 mm quartz EPR tubes and loaded into the resonator cavity. Experimental data was collected using the 15 four-pulse DEER sequence [(π/2)ν1 – τ1 – (π)ν1 – t - (π)ν2 – (τ1 + τ2 – t) - (π)ν1 - τ2 – echo] at Q-band with a probe pulse width of 8/16 ns and pump pulse width of 70 ns. The frequency difference that was used between the pump and probe pulses was ramped from 35 MHz to 120 MHz. 16-step phase cycling at a temperature of 80 K collected out to ~2.0 µs for data acquisition overnight. The DEER data analysis was conducted using the MATLAB DEER Analysis 2018 Program.19 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 2018.20 The background correction was performed using a two-dimensional homogeneous model for liposomes. The best fit of the time domain data was used for optimizing the regularization parameter in the L-curve.

5.4 Results and Discussion Figure 5.2A shows the model of inactive S21IRS pinholin with both TMDs spanning the membrane bilayer and the dual spin-labeled sites used for the study connected by red lines. The DEER time domain data and resulting distance distributions are shown in Figure 5.2B. The double substitution W27C/A38C at the top of the helices, near the loop region, show a major DEER distance peak of 23 ±3 Å. The two DEER substitutions at the bottom of the helices, G14C/L53C and S8C/L53C, also show similar short distances at 26 ±3 Å and 25 ±3 Å, respectively. The presence of these short distances as the DEER substitutions move throughout the helices shows that the space between these helices is not changing as they span the membrane. This data suggests that both helices are incorporated within the lipid bilayer and are packing in close proximity to each other. These results also indicate that the addition of the five amino acids to the N-terminus of

98 the pinholin locks TMD1 of the inactive S21IRS pinholin in the membrane and prevents translocation. The structural model of active S2168 pinholin can be seen in Figure 5.3 along with the DEER time domain data and resulting distance distributions. The shortest distance observed is for the W27C/A38C spin label pair at the top of the helices, closest to the loop region, with a distance of 25 ±3 Å. This externalization of TMD1 becomes evident as the spin label pairs move down the helices. The five total dual spin-labeled DEER distance measurements for active S2168 pinholin showed increasing distances as we moved from the loop region to the N- and C-termini of the helices at the other surface of the bilayer. The distance observed for the shortest distance (W27C/A38C) was 25 ±3 Å. The intermediate distances for A17C/A38C, A17C/V46C, and S16C/G48C were 35 ±3 Å, 35 ±3 Å and 47 ±3 Å, respectively. These three distances, to be the intermediate distances between the W27C/A38C and S8C/L53C pairs, follow a progression of increasing distance. More importantly, all three pairs are showing major distance peaks greater than 26 ±3Å, which was the longest observed distance for pinholin in the inactive form. The most compelling piece of data to show the partial externalization of TMD1 is the comparison of dual spin-labeled positions S8C/L53C on the active and inactive forms of pinholin. In the inactive S21IRS pinholin the S8C/L53C positions show a distance of 26 ±3 Å indicating the presence of TMD1 incorporated within the membrane. The same S8C/L53C spin-labeled positions on the active S2168 form of pinholin reveal a much longer distance of 50 ±3 Å. In Figure 5.4 a comparison of preliminary structural simulations for TMD1 externalization of active S2168 pinholin with the S8C/L53C spin-labeled positions is shown. In Figure 5.4A preliminary simulation result indicates a theoretical distance of 74 ±4 Å for the previously hypothesized complete externalization of TMD1. Therefore, the experimental data showing a predominant distance population at 50 ±3 Å does not support the model of a fully externalized TMD1 but instead suggests a partially externalized TMD1 from the membrane. This model is supported by the simulation shown in Figure 5.4B with a partial TMD1 externalization distance of 50 ±4 Å. Based on the data shown in Figure 5.2, the structural model of the inactive S21IRS form of pinholin was proposed with both TMDs of pinholin present in the membrane, shown in Figure 5.1A.

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The DEER distance data shown in Figure 5.3 suggests a new structural model of the active S2168 form of pinholin, shown in Figure 5.5. This model deviates from the current models proposed in the literature as it shows a partial externalization of TMD1 from the membrane instead of a complete externalization.4, 7 In this model the first TMD of pinholin is lying on the surface of the membrane and interacting with the lipid headgroups of the bilayer. This model is consistent with previously published 31P and 2H solid-state nuclear magnetic resonance (SS-NMR) spectroscopy studies.

5.5 Conclusions In conclusion, we have provided direct evidence for two different structural conformations for the inactive S21IRS and active S2168 pinholin within the membrane using DEER spectroscopic measurements. This study increases the understanding of the structure-function relationship of the pinholin as it progresses from a nonfunction to a functional conformation. Additionally, this work expands the application of EPR DEER spectroscopy to study structural and conformational difference of complex membrane protein systems.

5.6 Acknowledgements: We are grateful to the members of the Young group at Texas A&M University for their experimental suggestions. This work was generously supported by a NSF CHE- 1807131 grant and a NIGMS/NIH Maximizing Investigator’s Research Award (MIRA) R35 GM126935 award. Gary A. Lorigan would also like to acknowledge support from the John W. Steube Professorship.

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5.7 Figures:

Figure 5.1 The current functional model for the structural conformations of A) active S21 pinholin, showing the complete externalization of TMD1 from the membrane, and B) inactive S21 pinholin with the five additional N-terminus amino acids locking TMD1 inside the membrane.

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Figure 5.2 A) Ribbon diagram of membrane incorporated inactive S21IRS pinholin with DEER substitution pairs marked and numbered in red. B) The corresponding frequency modulation fit and distance distribution for each DEER pair.

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Figure 5.3 A ribbon diagram of membrane incorporated active S2168 pinholin with DEER substitution pairs marked and numbered in blue. The corresponding frequency modulation fit and distance distribution for each DEER pair which show a progression of increasing distances from the first to the fifth DEER pair.

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Figure 5.4 A preliminary structural model comparison of active S2168 pinholin. A) The previously hypothesized complete externalization of TMD1 from the membrane with a distance of 74 ±4 Å. B) The newly proposed model showing partial TMD1 externalization with a distance of 50 ±4 Å

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Figure 5.5 Newly proposed structural model of active S2168 pinholin in the membrane highlighting the partial externalization of TMD1 laying on the surface of the membrane.

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References:

1. Young, R., Phage Lysis: Three Steps, Three Choices, One Outcome. Journal of Microbiology 2014, 52 (3), 243-258. 2. Young, R., Phage Lysis: Do we have the hole story yet? Curr Opin Microbiol. 2013, 16 (6), 1-8. 3. Park, T.; Struck, D. K.; Dankenbring, C. A.; Young, R., The pinholin of lambdoid phage 21: Control of lysis by membrane depolarization. Journal of Bacteriology 2007, 189 (24), 9135-9139. 4. Park, T.; Struck, D. K.; Deaton, J. F.; Young, R., Topological dynamics of holins in programmed bacterial lysis. Proceedings of the National Academy of Sciences of the United States of America 2006, 103 (52), 19713-19718. 5. 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. 6. 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. 7. Pang, T.; Savva, C. G.; Fleming, K. G.; Struck, D. K.; Young, R., Structure of the lethal phage pinholin. PNAS 2009, 106 (45), 18966-18971. 8. Pang, T.; Park, T.; Young, R., Mutational analysis of the S21 pinholin. Molecular Microbiology 2010, 76 (1), 68-77. 9. Sahu, I. D.; Lorigan, G. A., Site-Directed Spin Labeling EPR for Studying Membrane Proteins. Biomed Research International 2018. 10. Sahu, I. D.; Kroncke, B. M.; Zhang, R. F.; Dunagan, M. M.; Smith, H. J.; Craig, A.; McCarrick, R. M.; Sanders, C. R.; Lorigan, G. A., Structural Investigation of the Transmembrane Domain of KCNE1 in Proteoliposomes. Biochemistry 2014, 53 (40), 6392-6401. 11. Sahu, I. D.; Lorigan, G. A., Biophysical EPR Studies Applied to Membrane Proteins. J Phys Chem Biophys: 2015; Vol. 5. 12. Jeschke, G.; Polyhach, Y., Distance measurements on spin-labelled biomacromolecules by pulsed electron paramagnetic resonance. Physical Chemistry Chemical Physics 2007, 9 (16), 1895-1910. 13. 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. 14. 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. 15. Jeschke, G., DEER Distance Measurements on Proteins. Annual Review of Physical Chemistry, Vol 63 2012, 63, 419-446. 16. 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.

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17. Pang, T.; Park, T.; Young, R., Mapping the pinhole formation pathway of S21. Molecular Microbiology 2010, 78 (3), 710-719. 18. Drew, D. L.; 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 S-21 pinholin protein. Analytical Biochemistry 2019, 567, 14-20. 19. Jeschke, G.; Chechik, V.; Ionita, P.; Godt, A.; Zimmermann, H.; Banham, J.; Timmel, C. R.; Hilger, D.; Jung, H., DeerAnalysis2006 - a comprehensive software package for analyzing pulsed ELDOR data. Applied Magnetic Resonance 2006, 30 (3-4), 473-498. 20. Chiang, Y. W.; Borbat, P. P.; Freed, J. H., The determination of pair distance distributions by pulsed ESR using Tikhonov regularization. Journal of Magnetic Resonance 2005, 172 (2), 279-295.

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Chapter 6

Conclusions and Future Directions

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6.1 Summary of Work: Understanding the dynamic structure of membrane proteins is critical for developing our understanding of their functions and mechanisms.1, 2 The work performed in this dissertation used powerful biophysical techniques to better characterize the pinholin S21 bacteriophage lytic protein in both an active and inactive form. Three pulse ESEEM spectroscopy approach was utilized to study the pinholin secondary structure, while four pulse DEER spectroscopy was used to probe the different structural models of the active and inactive forms pinholin in the membrane. SS-NMR spectroscopy was used to study the dynamic properties of membrane lipid bilayers in the presence of the active and inactive forms of pinholin. In Chapter 2, the synthesis of both the active S2168 and inactive S21IRS forms of pinholin using solid phase peptide synthesis (SPPS) was described in detail. Circular dichroism (CD) spectroscopy showed a predominately α-helical secondary structure for both the active and inactive forms of pinholin with the CD spectra showing a characteristic double minima at 208 nm and 222nm. The CD molar ellipticity ratio ([θ]222/ [θ]208) also provided preliminary data suggesting the presence of oligomeric states of pinholin corresponding to the penultimate step of the pinholin depolarization pathway. Continuous wave (CW) electron paramagnetic resonance (EPR) spectroscopy was used to show successful spin labeling of pinholin. 31P solid-state (SS) nuclear magnetic resonance (NMR) spectroscopy indicated that pinholin was incorporated into both bicelles and proteoliposome systems. These results showed interactions of both the active and inactive forms of pinholin with the lipid bilayer, to varying degrees, through decreases in the 31P CSA width when compared to the empty DMPC MLVs. These differences in the way the active S2168 and inactive S21IRS forms of pinholin interact with the membrane suggest differences in the role each form plays in the bacteriophage lytic pathway. Chapter 3 discussed the application of ESEEM spectroscopy to probe the local protein secondary structure of full-length functional systems such as pinholin for the first time. The local α-helical secondary structure of both TMD1 and TMD2 was confirmed with the presence of deuterium modulation observed at the i-3 and i-4 positions.3-7 The confirmation of the α-helical structure in TMD1 and TMD2 for the active S2168 and inactive S21IRS forms of the pinholin demonstrates the applicability of this technique to probe both

109 peripheral and integral membrane proteins. In addition, the comparison of ESEEM frequency domain intensities between the active S2168 and inactive S21IRS forms indicates there is no change in helical structure after TMD1 externalizes from the membrane. The presences of a small deuterium ESEEM peak for the i-2 position in TMD2 of the active S2168 pinholin could indicate TMD2 intermolecular interactions resulting from the oligomeric state of pinholin.8 This data, along with the data indicating α-helical secondary structure of TMD1 and TMD2 for inactive and active pinholin, would suggest there is no need for a refolding or conformational change of either TMD during the oligomerization step of the pinholin pathway. In chapter 4, 31P and 2H SS-NMR spectroscopy compared the dynamic interactions of the active S2168 and inactive S21IRS forms of pinholin with the phospholipid bilayer. 31P nuclei were used to probe the interaction of active and inactive pinholin with the lipid headgroups. The 31P CSA width values of 1 and 2 mol% active S2168 are smaller than the inactive S21IRS CSA values at the same mol% and temperature. The decrease in 31P CSA width indicates a higher degree of perturbations at the surface of the membrane for the active S2168 pinholin when compared to that of the inactive S21IRS form of pinholin. The 31P SS-NMR spectra also show an isotropic peak with an increasing line width as the concentration of pinholin increases. A greater isotropic linewidth is observed in the active S2168 pinholin, when compared to the inactive S21IRS. This could be a result of displacement of the lipids in the bilayer coming from the partial externalization of TMD1 of the active S2168 pinholin.9, 10 The 2H SS-NMR quadrupolar splitting data showed an overall decrease in the spectral width as the concentration (mol%) of pinholin was increased. Unlike 31P CSA trends, the inactive S21IRS showed a greater decrease in the 2H quadrupolar splitting. This trend indicates a greater degree of interaction with the acyl chains of the lipid coming from the inactive form of the pinholin. These interactions are originating from the presence of TMD1 remaining incorporated in the membrane, while pinholin is in an inactive form. This conclusion is further supported by the additional loss in resolution of the doublet peaks observed when comparing 1 mol% inactive S21IRS to the spectra for 1 mol% active S2168 pinholin.11-13 The order parameters determined from the quadrupolar splitting of the dePaked 2H SS-NMR spectra give additional support for the conclusions drawn from the

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31P and 2H data. The decrease in the order parameters of carbon 2-6 in the presence of 1 mol% active S2168 pinholin when compared to empty bilayers supports the idea of a partial TMD1 externalization from the membrane. This study highlights the application of SS-NMR spectroscopy to distinguish between the two different conformations of the pinholin protein and the resulting interactions with the lipid bilayer. Finally, Chapter 5 investigated the structural models of membrane incorporated active S2168 and inactive S21IRS pinholin. This was accomplished through site directed spin labeling (SDSL) coupled with four pulse DEER spectroscopy. Spin labeling site on each TMD were strategically chosen, based on previously published mutational analysis studies, as to not alter the structure or function of pinholin.14 The results from this study supported the current structural model of pinholin in an inactive conformation with both TMDs spanning the width of the lipid bilayer.15 Inactive S21IRS DEER spin-labeled pairs, both near the loop region of pinholin and at the terminus of each helix, all showed distances less than 26 ± 3 Å indicative of the helices packing in close proximity to each other. Alternatively, the results from the active S2168 form of the pinholin led to the proposal of an altered structural model for the translocation of TMD1 from the membrane. A series of five DEER spin label pairs, positioned from the loop region to the terminal end of each helix, showed a progression of increasing distance distributions ranging from 25 ± 3 Å to 50 ± 3 Å. A direct comparison between the active and inactive forms of pinholin is possible at positions S8C/L53C. The DEER data shows an inactive S21IRS distance of 26 ± 3 Å, indicative of helices in close proximity, while these same spin-labeled positions in the active S2168 pinholin show a distance of 50 ± 3 Å. The comparison of the active S2168 S8C/L53C data to the theoretical distance of 74 ± 4Å, generated through molecular dynamic simulations, indicated that a complete externalization of TMD1 from the membrane was not possible. This led to an altered structural model of active pinholin with TMD1 partially externalized from the membrane. This partial externalization is also supported by the SS-NMR study probing the interactions of active pinholin with the bilayer outlined in Chapter 4. The work shown in this dissertation has utilized a wide variety of powerful biophysical techniques and has better characterized the structure and dynamic interactions of the S21 bacteriophage membrane lytic pinholin protein with the membrane.

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We have, for the first time, successfully synthesized both the active and inactive forms of pinholin using SPPS. Using CD and ESEEM spectroscopy we reported the first experimental data probing the global and local α-helical secondary structure of the inhibitory and functional helices of pinholin. The results from the SS-NMR data probed the differences in the way the active and inactive forms of pinholin interact with the bilayer and revealed the first piece of evidence indicating TMD1 interacts with the lipid headgroups of the bilayer. DEER spectroscopic studies supported the current structural model of inactive pinholin with both TMDs spanning the membrane and led to a new structural model of the active pinholin. The active S2168 DEER data supported a partially externalized model of active pinholin with TMD1 laying on the surface of the membrane.

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6.2 Future Directions:

6.2.1 ESEEM Spectroscopy Studies Three-pulse EPR ESEEM could be used to probe the hole formation of the active S2168 form of pinholin. In the literature, the second transmembrane domain (TMD2) of pinholin is known as the functional domain due to its ability to oligomerize and form pinholes in the membrane. This work would involve site directed spin labeling (SCDL) of the functional domain of active S2168 pinholin followed by incorporation of pinholin into a proteoliposome lipid mimetic system at varying protein to lipid ratios. The rehydration of this system would be performed using a deuterated buffer. At the lowest protein to lipid ratio the concentration of active S2168 pinholin would be below the critical concentration needed for hole formation. In this case, the spin label would be located in the hydrophobic core of the lipid bilayer with the deuterated buffer unable to pass through the bilayer. This would result in no observed deuterium modulation in the ESEEM spectra. As the concentration of pinholin increases above the critical concentration holes would being to form in the membrane. The placement of the spin label on TMD2 of pinholin would allow for the spin label to interact with the deuterated buffer as it passed through the newly generated hole in the membrane. This would result in the presence of deuterium modulation in the ESEEM spectra. Moving the spin labeling site through TMD2 could help to probe which face of the functional helix is facing the hole or interacting with the bilayer.

6.2.2 SS-NMR Spectroscopic Studies This study would probe the incorporation of the active S2168 and inactive S21IRS into proteoliposomes containing alternative lipids including lipid mixtures. Lengthening the acyl chains to have 16-18 CH2 groups and changing head group charges, from phosphocholine to phosphoethanolamine for example, will allow for a better mimic of the pinholin natural membrane environment. Additionally, DMPC, which was used in this study, has the propensity to phase separate at higher temperatures if left for extended 31 periods of time. This limited our ability to perform P T1 relaxation experiments. Using different lipids would overcome this limitation and allow for an additional way to probe the interactions of active and inactive pinholin within the head groups of the bilayer.

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Due to the strong magnetic field strength of NMR spectroscopy when compared to EPR spectroscopy it is also possible to perform alignment studies of the pinholin. This would involve incorporating pinholin into bicelles instead of vesicles. Addition of paramagnetic lanthanide metals to the bicelles would give experimental control over the perpendicular vs parallel alignment of the bilayer.11, 16

6.2.3 DEER Spectroscopy Studies Using the mutational analysis work of the Young group, the effect of individual amino acids on the externalization of TMD1 can be performed using DEER spectroscopy. In Chapter 5 the spin label pairs S8C/L53C and G14C/L53C were shown to be reliable positions to probe the inactive S21IRS pinholin without effecting TMD1 externalization. This study would keep one of those two DEER pairs constant for each sample in the study while substituting selected TMD1 amino acids to probe given amino acid characteristics, like charge, size, or hydrophobicity. Monitoring the resulting DEER distance will determine if the mutational substitution influenced the translocation of TMD1. Additionally, the oligomerization state of the active S2168 pinholin can be probed through DEER spectroscopy through placing single spin-labeled positions in TMD2. Mixtures of these single spin-labeled active pinholin could be mixed with wildtype versions of the protein to counteract detection of excessive intermolecular distances.

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References:

1. Cournia, Z.; Allen, T. W.; Andricioaei, I.; Antonny, B.; Baum, D.; Brannigan, G.; Buchete, N. V.; Deckman, J. T.; Delemotte, L.; del Val, C.; Friedman, R.; Gkeka, P.; Hege, H. C.; Henin, J.; Kasimova, M. A.; Kolocouris, A.; Klein, M. L.; Khalid, S.; Lemieux, M. J.; Lindow, N.; Roy, M.; Selent, J.; Tarek, M.; Tofoleanu, F.; Vanni, S.; Urban, S.; Wales, D. J.; Smith, J. C.; Bondar, A. N., Membrane Protein Structure, Function, and Dynamics: a Perspective from Experiments and Theory. Journal of Membrane Biology 2015, 248 (4), 611-640. 2. Elofsson, A.; von Heijne, G., Membrane protein structure: Prediction versus reality. Annual Review of Biochemistry 2007, 76, 125-140. 3. Kubota, T.; Lacroix, J. J.; Bezanilla, F.; Correa, A. M., Probing alpha-3(10) Transitions in a Voltage-Sensing S4 Helix. Biophysical Journal 2014, 107 (5), 1117-1128. 4. Liu, L. S.; Lorigan, G., Probing the Secondary Structure of Membrane Proteins with the Pulsed EPR ESEEM Technique. Biophysical Journal 2014, 106 (2), 192A-192A. 5. Liu, L. S.; Sahu, I. D.; Bottorf, L.; McCarrick, R. M.; Lorigan, G. A., Investigating the Secondary Structure of Membrane Peptides Utilizing Multiple H-2-Labeled Hydrophobic Amino Acids via Electron Spin Echo Envelope Modulation (ESEEM) Spectroscopy. Journal of Physical Chemistry B 2018, 122 (16), 4388-4396. 6. Zhang, R. F.; Sahu, I. D.; Gibson, K. R.; Muhammad, N. B.; Bali, A. P.; Comer, R. G.; Liu, L. S.; Craig, A. F.; McCarrick, R. M.; Dabney-Smith, C.; Sanders, C. R.; Lorigan, G. A., Development of electron spin echo envelope modulation spectroscopy to probe the secondary structure of recombinant membrane proteins in a lipid bilayer. Protein Science 2015, 24 (11), 1707-1713. 7. Bottorf, L.; Rafferty, S.; Sahu, I. D.; McCarrick, R. M.; Lorigan, G. A., Utilizing Electron Spin Echo Envelope Modulation To Distinguish between the Local Secondary Structures of an alpha-Helix and an Amphipathic 3(10)-Helical Peptide. Journal of Physical Chemistry B 2017, 121 (14), 2961-2967. 8. 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. 9. Seelig, J., P-31 Nuclear Magnetic-Resonance And Head Group Structure Of Phospholipids In Membranes. Biochimica Et Biophysica Acta 1978, 515 (2), 105-140. 10. Lau, T. L.; Ambroggio, E. E.; Tew, D. J.; Cappai, R.; Masters, C. L.; Fidelio, G. D.; Barnham, K. J.; Separovic, F., Amyloid-beta peptide disruption of lipid membranes and the effect of metal ions. Journal of Molecular Biology 2006, 356 (3), 759-770. 11. Minto, R. E.; Adhikari, P. R.; Lorigan, G. A., A H-2 solid-state NMR spectroscopic investigation of biomimetic bicelles containing cholesterol and polyunsaturated phosphatidylcholine. Chemistry and Physics of Lipids 2004, 132 (1), 55-64. 12. Huster, D., Solid-state NMR spectroscopy to study protein lipid interactions. Biochimica Et Biophysica Acta-Molecular and Cell Biology of Lipids 2014, 1841 (8), 1146- 1160. 13. Abu-Baker, S.; Lorigan, G. A., Phospholamban and its phosphorylated form interact differently with lipid bilayers: A (31)P, (2)H, and (13)C solid-state NMR spectroscopic study. Biochemistry 2006, 45 (44), 13312-13322.

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14. Pang, T.; Park, T.; Young, R., Mutational analysis of the S21 pinholin. Molecular Microbiology 2010, 76 (1), 68-77. 15. Pang, T.; Savva, C. G.; Fleming, K. G.; Struck, D. K.; Young, R., Structure of the lethal phage pinholin. PNAS 2009, 106 (45), 18966-18971. 16. Nakazawa, Y.; Suzuki, Y.; Saito, H.; Asakura, T., The Interaction of A beta(1-40) Peptide with Lipid Bilayers and Ganglioside As Studied by Multinuclear Solid-State NMR. Nmr Spectroscopy of Polymers: Innovative Strategies for Complex Macromolecules 2011, 1077, 299-+.

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