Production of an Isotopically Labelled G-Protein Coupled Receptor In

Investigations into the production of integral membrane proteins for solid-state NMR spectroscopy

Rachel Munro

A Thesis presented to The University of Guelph

In partial fulfillment of requirements for the degree of Master of Science in Biophysics

Guelph, Ontario, Canada

Investigations into the production of integral membrane proteins for solid-state NMR spectroscopy

Rachel Munro Advisor: Professor L. Brown University of Guelph, 2015

This study seeks to probe the challenges associated with heterologous expression and isotopic labeling of microbial rhodopsins and the GPCR adenosine receptor (2A) (A2aR) through biophysical methods including SSNMR, FTIR and Raman spectroscopy. A2aR was transformed into Pichia pastoris, which has previously been shown to be able to cost-effectively produce isotopically-labelled eukaryotic membrane proteins for SSNMR studies; however poor expression resulted in contamination as detected by both FTIR and SSNMR. Next, for Anabaena sensory rhodopsin (ASR) produced in E. coli, we investigated the effect of a full-length construct on both expression and purification. Finally, we showed that the novel biosynthetic production of an isotopically labelled retinal ligand concurrently with its apoprotein proteorhodopsin in E. coli presents a cost effective alternative to de novo synthesis. By using alternately labeled carbon sources (glycerol) we were able to verify the biosynthetic pathway for retinal and assign several new carbon resonances for proteorhodopsin-bound retinal.

Acknowledgements

Firstly, I would like to thank my advisor, Dr. Leonid Brown, for his guidance, patience and unwavering support over the last four years. Additional thanks belong to Dr. Vladimir

Ladizhansky, for being on my advisory committee and for all his help with NMR. Also, I would like to thank Dr. John Dawson for being on my advisory committee. In addition, I thank Dr.

Hermann Eberl for chairing my examination committee. I would not have reached this point without any one of you.

This thesis could not have been completed without the assistance of Dr. Dyanne Brewer and Dr. Armen Charchoglyan at the Advanced Analysis Center at the University of Guelph. I also thank Shweta Singh and Bernadette Byrne at Imperial College London for the A2aR samples, and

Kwang-Hwan Jung for the BUT5600-PR cells.

Thank you to past members of the lab group, Dr. Shenlin Wang and Dr. Ying Fan. I learned so much from you both and all your assistance in learning how to work with membrane proteins, pack samples and run gels, western blots and FTIR. This thesis would not have happened without you. To all the current members of the research group David Bolton, Daryl Good, Andrew Harris,

David Park, and Meg Ward, thank you for all your help and listening every time I needed to complain.

To my family, who have been there through every tear, set-back, and mysterious illness, there are not enough words to express my gratitude for your support. And lastly, to Alex and our cats, past and present, you brought me dinner and kept me sane. Thank you.

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Statement of Work Performed

All work was performed by me unless otherwise stated here. The plasmids for A2aR were supplied by Hino et al. at the Japan Science and Technology Agency and the Prosser group at

University of Toronto – Mississauga. Initial FTIR experiments and the transformation of FL-ASR into BL21 cells were performed with the assistance of Dr. Ying Fan. SSNMR experiments were conducted by Dr. Vladimir Ladizhansky and Meaghan Ward. Mass spectrometry identification of the gel extracts was conducted at the Advanced Analysis Center at the University of Guelph by

Dr. Dyanne Brewer and Dr. Armen Charchoglyan. Time-trial experiments on FL-ASR were conducted by Craig Flaherty. The βUT5600-PR cells were supplied by Jung et al. and initial M9 expression protocol developed by Jeff Gaudet. All resonance Raman spectroscopy was performed by Dr. Leonid Brown.

Table of Contents

ACKNOWLEDGEMENTS iii

STATEMENT OF WORK PERFORMED iv

TABLE OF CONTENTS v

LIST OF TABLES AND FIGURES viii

CHAPTER 1 - INTRODUCTION 1

1.1 MEMBRANE PROTEINS 1 1.2 TECHNIQUES USED TO STUDY MEMBRANE PROTEINS 2 1.2.1 NUCLEAR MAGNETIC RESONANCE 2 1.2.1.1 Principles of NMR spectroscopy 3 1.2.1.2 Sample Preparation for SSNMR of membrane proteins 6 1.2.1.3 Multidimensional NMR spectroscopy 7 1.2.1.4 Solid-state NMR of Membrane Proteins 8 1.2.1.5 Magic Angle spinning solid-state NMR 9 1.2.1.6 SSNMR interactions 10 1.2.1.7 SSNMR derived structure examples 10 1.2.2 FOURIER-TRANSFORM INFRARED SPECTROSCOPY 12 1.2.3 RESONANCE RAMAN SPECTROSCOPY 15 1.3 SUMMARY OF GOALS 16

CHAPTER 2 - PRODUCTION OF ADENOSINE RECEPTOR (2A) FOR SSNMR 18

2.1 G-PROTEIN COUPLED RECEPTORS 18 2.1.1 GPCR STRUCTURE 20 2.1.2 GPCR SIGNAL TRANSDUCTION MECHANISM 23 2.1.3 GPCR SIGNALLING PATHWAYS 24 2.2 ADENOSINE RECEPTOR TYPE 2A 26 2.2.1 A2AR STRUCTURE 28 2.2.2 OLIGOMERIZATION OF A2AR 30 2.2.3 LIGAND BINDING 31 2.3 HETEROLOGOUS EXPRESSION IN PICHIA PASTORIS 31 2.3.1 COMPARISON TO OTHER EXPRESSION HOSTS 32 2.3.2 HOW DOES PICHIA EXPRESSION WORK (AOX1) 33 2.4 STUDIES ON FERMENTER PRODUCED C-TERMINALLY TRUNCATED A2AR 34 2.4.1 MATERIALS AND METHODS FOR ANALYSIS OF 15N-334-A2AR 34 2.4.1.1 Reconstitution of 15N-334-A2aR 34 2.4.1.2 Preparation of samples for SSNMR 36 2.4.2 FTIR ANALYSIS OF 15N-334-A2AR 36 2.4.3 SSNMR OF 15N-334-A2AR 39 2.5 MATERIALS AND METHODS FOR SHAKER FLASK A2AR PRODUCTION 42 2.5.1 PLASMID 42

2.5.2 TRANSFORMATION INTO YEAST CELLS 43 2.5.3 IN-COLONY PCR SCREENING 44 2.5.4 SCREENING FOR HIGH EXPRESSION MUTANTS 45 2.5.5 LARGE SCALE EXPRESSION OF A2AR 46 2.5.6 PURIFICATION OF A2AR 47 2.5.7 RECONSTITUTION OF A2AR FOR SSNMR STUDIES 48 2.5.8 PREPARATION OF SAMPLES FOR SSNMR 49 2.6 RESULTS AND DISCUSSION 49 2.6.1 IN-COLONY SCREENING 49 2.6.2 EXPRESSION OF A2AR 50 2.6.3 PURIFICATION OF A2AR 51 2.6.4 FTIR STUDIES OF RECONSTITUTED A2AR 53 2.6.5 SSNMR OF TRUNC-A2AR 55 2.6.6 MASS SPECTROMETRY OF GEL EXTRACTS 56

CHAPTER THREE – MICROBIAL RHODOPSINS 61

3.1 RHODOPSINS 61 3.2 BACTERIORHODOPSIN 62 3.3 THE PHOTOCHEMICAL CYCLE 63 3.4 EXPRESSION OF RHODOPSINS IN E. COLI 64

CHAPTER FOUR – FULL LENGTH ASR 67

4.1 ANABAENA SENSORY RHODOPSIN 67 4.2 ASR STRUCTURE AND PHOTOCHEMISTRY 68 4.3 FULL-LENGTH ASR 71 4.4 MATERIALS AND METHODS 72 4.4.1 EXPRESSION OF FL-ASR 72 4.4.2 PURIFICATION OF FL-ASR 73 4.4.3 ANALYSIS OF CONTAMINATING PROTEINS 74 4.5 RESULTS AND DISCUSSION 74 4.5.1 TEMPORAL DEPENDENCE OF EXPRESSION 74 4.5.2 IDENTIFICATION OF CONTAMINATING PROTEINS 75

CHAPTER FIVE - BIOSYNTHETIC PRODUCTION OF ISOTOPICALLY LABELLED RETINAL 80

5.1 RETINAL BIOSYNTHESIS PATHWAY 80 5.2 PLASMIDS 81 5.3 PROTEORHODOPSIN 82 5.4 PROTEORHODOPSIN STRUCTURE 83 5.5 RETINAL LABELLING PREDICTION 86 5.6 RESONANCE RAMAN SPECTROSCOPY OF PROTEORHODOPSIN 91 5.7 MATERIALS AND METHODS 94 5.7.1 MATERIALS 94 5.7.2 EXPRESSION OF ΒUT5600-PR IN 2XYT 94 5.7.3 EXPRESSION OF ΒUT5600-PR IN M9+PWL 95

5.7.4 PURIFICATION OF ΒUT5600-PR 96 5.7.5 RECONSTITUTION OF ΒUT5600-PR 96 5.7.6 RAMAN SPECTROSCOPY 97 5.7.7 SSNMR EXPERIMENTS 97 5.8 RESULTS AND DISCUSSION 97 5.8.2 EXPRESSION OF ΒUT5600-PR IN MINIMAL MEDIA 98 5.8.3 RAMAN SPECTROSCOPY 99 5.8.4 SSNMR SPECTROSCOPY OF ΒUT5600-PR 104

CHAPTER SIX – FINAL CONCLUSIONS AND FUTURE DIRECTIONS 112

REFERENCES 115

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List of Tables and Figures

Figures

FIGURE 1 – ENERGY SCHEMATIC OF NUCLEAR SPIN IN AN APPLIED MAGNETIC FIELD [28]...... 4 FIGURE 2 - EXAMPLE OF A TYPICAL STATIC FTIR SPECTRUM OF A MEMBRANE PROTEIN [56] ...... 13 FIGURE 3 – GPCR STRUCTURES DETERMINED ACCORDING TO THEIR CLASSIFICATION. STRUCTURES ARE IDENTIFIED WITH RED FLAGS. GPCR CLASSES ARE REPRESENTED IN BLUE (CLASS A), RED (CLASS B), YELLOW (CLASS C), PURPLE (CLASS D), AND GREEN (CLASS F) [75]...... 20 FIGURE 4 – SUPERPOSITION OF TWELVE CRYSTALLIZED GPCR FAMILY MEMBERS. RHODOPSIN (ORANGE), Β2- ADRENOCEPTOR (MAGENTA), Β1-ADRENOCEPTOR (CYAN), ADENOSINE A2A (PINK), DOPAMINE D3 (BROWN), CHEMOKINE CXCR4 (GREEN), HISTAMINE H1 (MUSTARD), MUSCARINIC ACETYLCHOLINE M2 (BRICK RED), MUSCARINIC ACETYLCHOLINE M3 (GREY), Δ-OPIOID (RED), Μ-OPIOID (PURPLE), AND S1P1 (BLUE) ARE DEPICTED [78]...... 22 FIGURE 5 – SIGNAL TRANSDUCTION MECHANISM IN Β2-AR [82] SHOWING THE CONFORMATIONAL CHANGE IN THE MOLECULES UPON AGONIST ACTIVATION [23]...... 24 FIGURE 6 – GPCR SIGNALLING OVERVIEW CONTAINING EACH CANONICAL PATHWAY. PATHWAYS ARE COLOURED ACCORDING TO THEIR G-PROTEIN SUBUNIT AND DOWNSTREAM PATHWAY COMPONENTS [91]...... 26 FIGURE 7 – ADENOSINE RECEPTOR SIGNALLING MECHANISM FOR THE FOUR KNOWN SUBTYPES: A1, A3, A2B AND A2A [94]...... 27 FIGURE 8 - STRUCTURE OF A2AR BOUND BY AN NECA MOLECULE (RED) IN THE ACTIVE STATE. PROTEIN IS VISUALIZED USING PYMOL (PDB ID: 3VGA) ...... 29 FIGURE 9 – FORMATION OF OLIGOMERS IN HETEROLOGOUSLY PRODUCED A2AR. A) REDUCTION IN DIMERIZATION OF A2AR DUE TO POINT MUTATION IN HELIX V [107]. B) DETECTION OF A2AR OLIGOMERS AFTER SOLUBILIZATION OF PROTEIN EXPRESSED IN HEK 293 [103]. C) DETECTION OF A2AR HOMO-OLIGOMERS IN HEK 293 CELLS USING FLAG-TAG DETECTION AT THE CELL SURFACE [103]. D) REDUCTION IN OLIGOMERS IN A2AR EXPRESSED IN P. PASTORIS AND PURIFIED THROUGH FLAG-TAG AFFINITY [2] FOLLOWED BY TEV CLEAVAGE AND METAL AFFINITY PURIFICATION [3] [108]...... 31 FIGURE 10 – BASELINE-CORRECTED STATIC FTIR SPECTRA OF A2AR RECONSTITUTED INTO PC/PS/CHOLESTEROL (9:1:2.3) LIPOSOMES AT DIFFERENT P/L RATIOS (1:2, 1:1, 2:1, 4:1). ABSORBANCE MEASURED ON BRUKER IFS66VS MACHINE WITH A TEMPERATURE-CONTROLLED SAMPLE HOLDER (HARRICK) CONNECTED TO A CIRCULATING WATER BATH (FISHER). THE PELLETS WERE RESUSPENDED IN 600 ΜL OF DH2O AND 100 ΜL OF THE SUSPENSION WAS DRIED ONTO THE CAF2 WINDOW...... 37 FIGURE 11 – BASELINE-CORRECTED STATIC FTIR SPECTRA OF A2AR RECONSTITUTED INTO A PC/PS/CHOLESTEROL (9:1:2.3) LIPOSOMES MEASURED AT A TWO WEEK INTERVAL IN 4 °C. ABSORBANCE MEASURED ON BRUKER IFS66VS MACHINE WITH A TEMPERATURE-CONTROLLED SAMPLE HOLDER (HARRICK) CONNECTED TO A TH CIRCULATING WATER BATH (FISHER). APPROXIMATELY 1/10 OF THE SAMPLE (0.56 MG) WAS LOADED ON CAF2 WINDOW FOR MEASUREMENT...... 38 FIGURE 12 – 1D SSNMR SPECTRA OF 15N-334-A2AR. SAMPLES WERE RECONSTITUTED INTO PC/PS/CHOL (9:1:1) LIPOSOMES (GREEN) AND DOPC/PS/CHOL (9:1:1) LIPOSOMES (BLACK) AT A PROTEIN TO LIPID RATIO OF 2:1 (W/W). PC/PS/CHOL WAS SCANNED 16456 TIMES WHILE DOPC/PS/CHOL WAS SCANNED 48000 TIMES...... 40 FIGURE 13 – 1D SSNMR SPECTRA OF 15N-334-A2AR RECONSTITUTED INTO DOPC/PS/CHOL LIPOSOMES. SPECTRA WERE TAKEN BEFORE (BLACK) AND AFTER (PURPLE) D2O INCUBATION...... 41 FIGURE 14 – 1D SSNMR SPECTRA OF 15N-334-A2AR (BLACK), PROTEORHODOPSIN (GREEN), AND LEPTOSPHAERIA RHODOPSIN (RED)...... 42 FIGURE 15 – PLASMID AND CONSTRUCT USED FOR A2AR EXPRESSION IN P. PASTORIS. A) PLASMID USED FOR TRANSFORMATION (PPIC9K). PLASMID MAP GENERATED IN SNAPGENE. B) TRUNCATED A2AR SHOWING ATTACHED FLAG AND 10XHIS TAG ON THE N AND C TERMINI RESPECTIVELY. TOPOGRAPHY MAP GENERATED BY PDBSUM...... 43 FIGURE 16 – IN-COLONY PCR OF A2AR. 1) MARKER. 2-6) TRUNC-A2AR COLONIES. 7-10) FL-A2AR COLONIES...... 50 FIGURE 17 – IMMUNOBLOT FOR TRUNC-A2AR EXPRESSION SCREENING OF 8 COLONIES OF P. PASTORIS. COLONIES GROWN IN 25 ML SHAKER CULTURES BEFORE BEING BROKEN, AND THEIR MEMBRANES WERE TREATED WITH 10 % SDS PRIOR TO CENTRIFUGATION. THE SUPERNATANT WAS RUN BY GEL ELECTROPHORESIS AND DETECTED USING QIAGEN/BIORAD ANTIBODIES...... 51

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FIGURE 18 – SDS-PAGE ANALYSIS OF PURIFICATION OF TRUNC-A2AR BY NICKEL AFFINITY RESIN. A) WESTERNC PROTEIN MARKER (BIO-RAD). B) ELUATE. C) SOLUBILIZED MEMBRANE FRACTION. D) FLOW-THROUGH AFTER RESIN BINDING. E-G) 15 MM IMIDAZOLE WASHES OF PROTEIN-BOUND RESIN...... 52 FIGURE 19 – SDS-PAGE OF FLAG-TAG PURIFICATION OF FL-A2AR AND TRUNC-A2AR. A) HIS-TAG MARKER. B) SOLUBILIZED TRUNC-A2AR MEMBRANE FRACTION. C) FLOW THROUGH AFTER FLAG-AFFINITY RESIN BINDING FOR TRUNC-A2AR. D) TRUNC-A2AR ELUATE AFTER FLAG-TAG PURIFICATION. E) HIS-TAG AFFINITY PURIFICATION ELUATE OF TRUNC-A2AR. F) SOLUBILIZED FL-A2AR MEMBRANE FRACTION. G) FLOW THROUGH AFTER FLAG-AFFINITY RESIN BINDING FOR FL-A2AR. H) FL-A2AR ELUATE AFTER FLAG-TAG PURIFICATION. I) HIS-TAG AFFINITY PURIFICATION ELUATE OF FL-A2AR...... 53 FIGURE 20 – BASELINE-CORRECTED STATIC FTIR SPECTRA OF A2AR RECONSTITUTED INTO A DOPC/PS/CHOLESTEROL (9:1:1) LIPOSOME MEASURED FOR 15N-TRUNC-A2AR (BLUE) AND NATURAL ABUNDANCE TRUNC-A2AR (RED). ABSORBANCE MEASURED ON BRUKER IFS66VS MACHINE WITH A TEMPERATURE-CONTROLLED SAMPLE HOLDER (HARRICK) CONNECTED TO A CIRCULATING WATER BATH (FISHER)...... 54 FIGURE 21 – BASELINE-CORRECTED STATIC FTIR SPECTRA OF A2AR RECONSTITUTED INTO A DOPC/PS/CHOLESTEROL (9:1:1) LIPOSOME MEASURED FOR TRUNC-A2AR (BLUE) AND NATURAL ABUNDANCE 334-A2AR (RED). ABSORBANCE MEASURED ON BRUKER IFS66VS MACHINE WITH A TEMPERATURE- CONTROLLED SAMPLE HOLDER (HARRICK) CONNECTED TO A CIRCULATING WATER BATH (FISHER)...... 55 FIGURE 22 - 1D SSNMR SPECTRA OF 15N-TRUNC-A2AR (RED) AS COMPARED TO 15N-334-A2AR (BLACK)...... 56 FIGURE 23 – MASS SPECTROMETRY RESULTS BASED ON IN-GEL DIGESTION OF PUTATIVE 70 KDA PROTEIN (BEFORE PURIFICATION). HITS ARE UNDERLINED IN BLUE...... 57 FIGURE 24 – STRUCTURAL MODEL OF AOX1. A) SECONDARY STRUCTURE IS COLOURED WITH Β CONTENT IN RED, Α HELICES IN CYAN AND LOOP IN MAGENTA. B) SURFACE HISTIDINES IDENTIFIED IN RED. HOMOLOGY MODEL PRODUCED BY SWISS-MODEL BASED ON THE CRYSTAL STRUCTURE OF ASPERGILLUS FLAVUS FAD GLUCOSE DEHYDROGENASE (PDB ID: 4YNT) AND RENDERED IN PYMOL...... 58 FIGURE 25 – MASS SPECTROMETRY IDENTIFICATION OF PUTATIVE 70 KDA PROTEIN ISOLATED FROM SDS-PAGE OF CELL-EXTRACTS AFTER PURIFICATION. SEQUENCE OF NUAM WITH IDENTIFIED PEPTIDES UNDERLINES IN BLUE...... 59 FIGURE 26 – HOMOLOGY MODEL OF NUAM. A) SECONDARY STRUCTURE IS COLOURED WITH Β CONTENT IN RED, Α HELICES IN CYAN AND LOOP IN MAGENTA. B) SURFACE HISTIDINES IDENTIFIED IN RED. HOMOLOGY MODEL PRODUCED BY SWISS-MODEL BASED ON THE CRYSTAL STRUCTURE OF CHAIN 3 OF THE HYDROPHILIC ARM OF RESPIRATORY COMPLEX I FROM THERMUS THERMOPHILUS (PDB ID: 2FUG) AND RENDERED IN PYMOL...... 59 FIGURE 27 – PROTON TRANSFER CHAIN IN BR. PATHWAY OF THE PROTON TRANSFER TOWARDS THE EXTRACELLULAR SURFACE (LOCATED AT THE BOTTOM) IS OUTLINED [129]...... 63 FIGURE 28 – PHOTOCYCLE OF BR SHOWING THE RETINAL CONFORMATION IN THE PHOTOINTERMEDIATES COLOURED ACCORDING TO THEIR CORRESPONDING WAVELENGTHS [132] ...... 64 FIGURE 29 – SAMPLE PREPARATION OF MICROBIAL RHODOPSIN PR IN E. COLI BASED EXPRESSION SYSTEM FOR SSNMR ...... 65 FIGURE 30 – MODEL OF THE E. COLI MEMBRANE ENVELOPE [135]...... 66 FIGURE 31 –A) REPRESENTATION OF THE ASR SIGNAL TRANSDUCING MECHANISM SHOWING DIRECT INTERACTION OF THE TRANSDUCER WITH DNA [136]. B) STRUCTURE OF THE PHYCOBILISOME SHOWING THE LIGHT HARVESTING PROCESS [137]...... 67 FIGURE 32 – STRUCTURE OF C-TERMINALLY TRUNCATED ASR AS DETERMINED BY SSNMR SPECTROSCOPY. A) OVERALL STRUCTURE WITH BOUND RETINAL IN YELLOW. B) RESIDUES ASSOCIATED WITH RETINAL BINDING IN ASR. C) IMPORTANT POLAR CYTOPLASMIC CLUSTER RESIDUES AND THEIR LOCATIONS RELATIVE TO HELICES IN ASR. [26] ...... 69 FIGURE 33 – PHOTOCHROMISM OF ASR [126]...... 69 FIGURE 34 – STRUCTURE OF THE ASR TRIMER AS DETERMINED BY SSNMR WITH INTERMONOMER CONTACTS SHOWN ON THE RIGHT [26] ...... 70 FIGURE 35 – STRUCTURE OF FL-ASR BASED ON THE SSNMR DERIVED STRUCTURE OF C-TERMINALLY TRUNCATED ASR (PDB ID: 2M3G). RETINAL IS IN YELLOW WITH IMPORTANT RESIDUES IN GREEN. MODEL PRODUCED IN PYMOL WITH A CYTOSOLIC HELIX MANUALLY ATTACHED FOR VISUALIZATION PURPOSES...... 72 FIGURE 36 – FL-ASR PRODUCTION MONITORED THROUGH PINK COLOUR FORMATION SHOWING DEPENDENCE ON POST- INDUCTION INCUBATION TIME LENGTH...... 75 FIGURE 37 – SDS-PAGE OF FL-ASR DURING THE PURIFICATION PROCESS STAINED WITH (A) COOMASSIE BLUE AND (B) WITH INVISION HIS-TAG STAIN. 1) FLOW THROUGH AFTER BATCH BINDING. 2-3) 35 MM IMIDAZOLE

WASHES. 4) FLOW THROUGH AFTER CYTOPLASM INCUBATION. 5) 35 MM IMIDAZOLE WASH AFTER CYTOPLASM BINDING. 6) FINAL ELUATE AFTER CYTOPLASM INCUBATION. FL-ASR MONOMER INDICATED WITH *. OMPF BAND DENOTED WITH **...... 76 FIGURE 38 – MASS SPECTROMETRY ANALYSIS OF 30 KDA BAND ISOLATED FROM SDS-PAGE AS COMPARED TO THE SEQUENCE OF OMPF. IDENTIFIED FRAGMENTS ARE UNDERLINED IN BLUE...... 77 FIGURE 39 – STRUCTURE OF OMPF WITH BETA-BARREL ARCHITECTURE IN GREEN AND INNER HELIX IDENTIFIED IN RED [149]...... 77 FIGURE 40 – COLICIN N TRANSLOCATION MECHANISM INTO MEMBRANES CONTAINING OMPF OLIGOMERS [150]...... 79 FIGURE 41 – SUSPECTED OMPF BINDING MECHANISM AS A RESULT OF FL-ASR INVERSION IN E. COLI INNER MEMBRANE...... 79 FIGURE 42 – A) BIOSYNTHESIS OF RETINAL DERIVATIVES FROM BETA-CAROTENE. B) SYNTHESIS OF BETA-CAROTENE AND RETINAL VIA THE TWO METABOLIC PATHWAYS (MVA AND MEP) AS DERIVED FROM GLUCOSE OR GLYCEROL SOURCES [155]...... 81 FIGURE 43 – A) GENES ENCODED BY PORANGE WHICH CONTROL SYNTHESIS OF BETA-CAROTENE. B) PORANGE VECTOR MAP [158]. C) PKJ900 VECTOR MAP [158]...... 82 FIGURE 44 – HOMOLOGY MODEL OF GREEN-ABSORBING PR BASED ON TEMPLATE OF BPR (PDB ID: 4JQ6) AND COMPILED BY SWISS-MODEL. MODEL REPRESENTATION IS PRODUCED IN PYMOL...... 85 FIGURE 45 – PREDICTED RETINAL LABELLING SCHEME USING 1,3-13C-GLYCEROL (ORANGE) AND 2-13C-GLYCEROL (TEAL). LABELLING SCHEME WAS DERIVED FROM THE MEP PATHWAY. IN PR, OXYGEN WILL BE REPLACED BY NITROGEN OF LYS SIDECHAIN, WHICH IS EXPECTED TO BE 15N LABELED IN BOTH SCENARIOS...... 87 FIGURE 46 – PREDICTED LABELLING SCHEME FOR AMINO ACIDS FROM CELLS GROWN IN 1,3-13C-GLYCEROL (BLUE) AND 2-13C-GLYCEROL (RED) [186]...... 88 FIGURE 47 - INTERACTIONS IN THE RETINAL BINDING POCKET AS PREDICTED BASED ON THE PR HOMOLOGY MODEL BASED ON BPR (FIGURE 44). RETINAL COLOURED ACCORDING TO PREDICTED LABELLING SCHEME AS DESCRIBED IN FIGURE 45. MODEL WAS GENERATED IN PYMOL...... 89 FIGURE 48 – RESONANCE RAMAN BANDS WITH CORRESPONDING MAJOR VIBRATION IDENTIFICATIONS IN BACTERIORHODOPSIN [63]...... 92 FIGURE 49 – RAMAN SPECTRA OF THE RETINAL CHROMOPHORE IN GREEN ABSORBING PR (GPR) AND IN BLUE- ABSORBING PR (BPR) [192]...... 93 FIGURE 50 – ΒUT5600-PR CELLS EXPRESSED IN 2XYT UNDER DIFFERENT INDUCTION CONDITIONS. A) NO INDUCER ADDED. B) 0.2 % L-ARABINOSE AND 1 MM IPTG. C) 7.5 µM ALL-TRANS-RETINAL AND 1 MM IPTG. D) 0.2 % L- ARABINOSE...... 98 FIGURE 51 – ΒUT5600-PR EXPRESSED WITH ADDITION OF A) 30 MM W, B) 20 MM W, C) 10 MM W, D) 5 MM W, AND E) 2 MM W IN THE PRESENCE OF 1 MM P AND L IN GLYCEROL-BASED M9 MEDIUM...... 98 FIGURE 52 – RESONANCE RAMAN SPECTRUM OF NATURAL ABUNDANCE ΒUT5600 PR IN DMPC/DMPA PROTEOLIPOSOMES...... 99 FIGURE 53 – NORMALISED RESONANCE RAMAN SPECTRA OF NATURAL ABUNDANCE (RED) AND 1,3-13C-GLYCEROL- DERIVED (BLUE) PR IN DMPC/DMPA PROTEOLIPOSOMES. MAJOR PEAKS IN THE LABELLED SAMPLE ARE IDENTIFIED IN TABLE 6...... 101 FIGURE 54 – RESONANCE RAMAN SPECTRA OF NATURAL ABUNDANCE (CYAN) AND 2-13C-GLYCEROL-DERIVED (PURPLE) PR IN DMPC/DMPA PROTEOLIPOSOMES. MAJOR PEAKS IN THE LABELLED SAMPLE ARE IDENTIFIED IN TABLE 7...... 103 FIGURE 55 – NORMALISED 15N 1D SSNMR SPECTRA OF ΒUT5600-PR (RED) AND BL21-PR (BLACK). BOTH SAMPLES WERE PRODUCED USING 2-13C-GLYCEROL...... 105 FIGURE 56 – 13C 1D SSNMR SPECTRA OF ΒUT5600-PR (BLACK) AND BL21-PR (RED). BOTH SAMPLES WERE PRODUCED USING 2-13C-GLYCEROL...... 106 FIGURE 57 - 13C-13C DARR SPECTRA OF ΒUT5600-PR (BLACK) AND BL21-PR (RED) SHOWING THE REGION OF 165.9 AND 121.8 PPM CORRELATION IDENTIFIED AS C13-C14 CROSSPEAK. BOTH SAMPLES WERE PRODUCED USING 2- 13C-GLYCEROL. SPECTRA WERE MEASURED AT 800 MHZ WITH A MIXING TIME OF 50 MS USED FOR BOTH SAMPLES...... 107 FIGURE 58 – 13C-13C DARR SPECTRA OF ΒUT5600-PR (BLACK) AND BL21-PR (RED), SHOWING THE REGION OF 139.7 AND 37 PPM CORRELATION IDENTIFIED AS C6-C1 CROSSPEAK. BOTH SAMPLES WERE PRODUCED USING 2-13C- GLYCEROL. SPECTRA WERE MEASURED AT 800 MHZ WITH A MIXING TIME OF 50 MS USED FOR BOTH SAMPLES...... 108

FIGURE 59 - 13C-13C DARR SPECTRA OF ΒUT5600-PR (BLACK) AND BL21-PR (RED), SHOWING THE REGION OF 147 AND 132 PPM CORRELATION IDENTIFIED AS C9-C10 CROSSPEAK. SPECTRA WERE MEASURED AT 800 MHZ WITH A MIXING TIME OF 50 MS USED FOR BOTH SAMPLES...... 108 FIGURE 60 - 13C-13C DARR SPECTRA OF ΒUT5600-PR (BLACK) AND BL21-PR (RED), SHOWING THE REGION OF 140 AND 137 PPM CORRELATION IDENTIFIED AS C6-C5 CROSSPEAK. SPECTRA WERE MEASURED AT 800 MHZ WITH A MIXING TIME OF 50 MS USED FOR BOTH SAMPLES...... 109 FIGURE 61 - 13C-13C DARR SPECTRA OF ΒUT5600-PR (BLACK) AND BL21-PR (RED), SHOWN IN THE REGION OF 100 TO 0 PPM. PUTATIVE LOCATION OF THE C1-C2 CROSS PEAK IS SHOWN BY BOX. SPECTRA WERE MEASURED AT 800 MHZ WITH A MIXING TIME OF 200 MS FOR ΒUT5600-PR AND 50 MS FOR BL21-PR...... 110

Tables

TABLE 1 – CHARACTERISTIC ABSORPTION WAVELENGTHS IN IR SPECTROSCOPY. SIGNAL STRENGTH IS REPRESENTED BY S (STRONG), M (MEDIUM), OR V (WEAK) [59]...... 14 TABLE 2 – LIPOSOMES PREPARED FOR RECONSTITUTION AND PROTEIN TO LIPID RATIOS (P/L) IN PREPARED LIPOSOMES ...... 35 TABLE 3 – PREDICTED ATOM PROXIMITIES WITHIN 4 Å DETECTABLE BY SSNMR WITHIN THE RETINAL BINDING POCKET OF PR. P, W, AND L WILL NOT BE DETECTABLE UNLESS PROVIDED IN THE ISOTOPICALLY LABELLED FORM AND ARE LABELLED WITH ASTERISKS ...... 89 TABLE 4 – CHEMICAL SHIFTS OF ALL-TRANS-RETINAL AND 13-CIS-RETINAL IN SOLUTION AND IN BR FOR ALL 13C- LABELLED CARBON ATOMS. RETINAL AND BR SHIFTS ADAPTED FROM HARBISON ET AL. [189] AND [190] RESPECTIVELY. PR PEAKS WERE ADAPTED FROM PFLEGER ET AL. [179], MEHLER ET AL. [191], RECKEL ET AL. [174], AND SHI ET AL. (UNPUBLISHED DATA) ...... 91 TABLE 5 – EXPECTED SHIFTS IN THE BR SPECTRUM GIVEN THE PREDICTED RETINAL LABELLING SCHEME. SHIFTS CALCULATED FROM SMITH ET AL. [63]. SEE FIGURE 48 FOR THE ASSIGNMENTS OF MAJOR CORRESPONDING VIBRATIONS...... 94 TABLE 6 – RESONANCE RAMAN PEAKS IN 1,3-13C-GLYCEROL-DERIVED PR SAMPLE. EXPECTED SHIFTS ARE CALCULATED FROM TABLE 5...... 101 TABLE 7 - RESONANCE RAMAN PEAKS IN 2-13C-GLYCEROL-DERIVED PR SAMPLE. EXPECTED SHIFTS ARE CALCULATED FROM TABLE 5...... 103 TABLE 8 – CHEMICAL SHIFTS OF ALL-TRANS-RETINAL AND 13-CIS-RETINAL IN SOLUTION AND IN BR FOR ALL 13C- LABELLED CARBON ATOMS. RETINAL AND BR SHIFTS ADAPTED FROM HARBISON ET AL. [189] AND [190] RESPECTIVELY. PR PEAKS WERE ADAPTED FROM PFLEGER ET AL. [179], MEHLER ET AL. [191], RECKEL ET AL. [174], AND SHI ET AL. (UNPUBLISHED DATA). ΒUT5600-PR RETINAL CHEMICAL SHIFTS ADDED FOR COMPARISON...... 111

Chapter 1 Introduction

1.1 Membrane Proteins

Cell membranes are complex and integral components of biological systems. In all organisms the membrane has two main components: proteins and lipids. Both these types of molecules are dynamic and reflect the different functions that cell membranes must perform including transport, surface adhesion, sensory transduction, and metabolism [1]. Over time the general model of the cell membrane has been adapted to include the vast variety of proteins and lipids present that can move laterally within the membrane. Static images of the cell membrane have been obtained using techniques such as electron microscopy, X-ray crystallography, and atomic force microscopy

(AFM) [2]. Techniques are currently being developed to study the cell membrane and its specific components in a more dynamic way which includes solid-state Nuclear Magnetic Resonance

(SSNMR) [3-5].

The membrane itself is a phospholipid bilayer with other lipids such as cholesterol affecting the fluidity of the membrane [2]. In order to study these membranes, it is essential to understand both the structure of its components and how they interact. Much of the functionality of cell membranes is facilitated by membrane proteins. This includes transport of essential molecules, signal transduction, and cell-to-cell interactions [1, 6]. As such, these proteins are significant targets for drug development and structural studies [7-9]. Membrane proteins consist of two main families: integral and peripheral [10]. Integral membrane proteins consist of at least one membrane spanning region of hydrophobic residues [10]. These membrane spanning regions can consist of

α-helices, or beta-strands [10]. Alpha-helical proteins are found in a wide variety of biological membranes, serving functions such as: receptors, channels, passive and active transporters, and

1 enzymes [10]. Comparatively, beta-barrels are found only in the outer membranes of Gram- negative bacteria and the outer membrane of mitochondria and chloroplasts [11]. These proteins generally act as channels for passive transport [11]. Peripheral membrane proteins are only associated with the membrane but may contain anchor residues [12]. There is a high correlation between structure and function in proteins. Therefore, in order to truly understand the function of these important biological molecules, it has become essential to obtain high resolution structures

[13].

1.2 Techniques used to study membrane proteins

Despite representing 35-40 % of the human genome, membrane proteins have been underrepresented in structural databases with only 569 unique protein structures in the protein data bank, the majority of which were determined via X-ray crystallography [14]. Most structural studies use purified protein, in either detergent micelles or proteoliposomes due to the hydrophobic core that membrane proteins possess [15]. This need for membrane mimics makes studying them so difficult. Other difficulties include low expression, poor solubilisation, and reduced stability in solution [16]. Additionally, cell membranes are complex and the many components make assigning definitive data points to the protein of interest difficult. This makes in vitro studies by X-ray crystallography and NMR less viable [17]. As such, numerous techniques are being developed to study membrane proteins in a native-like lipid environment [18].

1.2.1 Nuclear Magnetic Resonance

Nuclear Magnetic Resonance (NMR) is a widely used form of spectroscopy with numerous functions for studies of both inorganic and biological molecules. NMR was first measured in 1938 by Polish-born physicist Isidor Rabi within molecular beams [19]. This was further expanded in

1946 by Felix Bloch and Edward Mills Purcell for use on liquids and solids. They observed that

2 magnetic nuclei, such as 1H, could absorb radio frequency (RF) energy at a nuclei specific frequency. Nuclei under these conditions are termed in resonance giving rise to the term NMR

[19]. Furthermore, the discovery that the resonance frequency of a specific nucleus is dependent upon its chemical environment resulted in NMR being adopted by structural biologists and chemists [20]. Due to the continuous improvements in technology and theory, this technique has gained notoriety as an analytical tool in both simple inorganic substances and complex biological molecules [20].

In the past fifty years this technique has evolved from small molecules to larger protein complexes. As of 2014, over 8500 protein structures have been solved through NMR [21].

However, one of the most pharmaceutically relevant classes of proteins, integral membrane proteins, are underrepresented in the protein databank (PDB). In particular, the G-protein coupled receptors (GPCRs) which comprise fifty percent of drug targets, have only 21 solved structures

[22]. The majority of these were solved using X-ray crystallography, using advantageous thermostabilizing mutations or in the presence of stabilizing antibodies [23]. However, X-ray crystallography can introduce non-native conformations into the protein, especially into the loop regions of membrane proteins, as a result of the crystallization process [16]. Previous success with bacterial rhodopsins, 7-transmembrane helical proteins analogous to GPCRs, as well as with one

GPCR, seems to indicate that NMR, in particular solid-state NMR (SSNMR), has the potential to study these proteins under more physiologically relevant conditions [24-26].

1.2.1.1 Principles of NMR spectroscopy

All nuclei with an odd mass number have half-integer spin due to the presence of an unpaired proton or neutron. Protons, carbons, and nitrogens are the most commonly found nuclei in biological molecules. Whereas protons have spin -½, the most abundant isotope 12C is spinless

3 and does not produce NMR signal, and the 14N isotope of nitrogen has spin 1 and has large quadrupolar moment. The “NMR-friendly” 13C and 15N spin ½ isotopes can be incorporated into a recombinantly expressed protein by growing cells on the minimal medium that contains isotopic carbon and nitrogen sources. When placed in a magnetic field, the nuclei found in biological molecules such as 1H, 13C, 15N have two possible spin states: m = ± ½ (Figure 1) which are under thermal equilibrium as described by the Boltzmann distribution. Under an increasing magnetic field applied along the z-axis, the energy required to induce a transition from spin down to spin up increases proportionally (Figure 1). Therefore, by applying an RF field specific to the transition of the nuclei one can bring about a resonance condition. This energy is quite low resulting in a very small population difference between the two energy levels. As such, this technique has low sensitivity requiring sample concentrations in the 1 mM range in the case of solution NMR.

However, the lifetime of the excited state is quite long, from millisecond to seconds, allowing for multidimensional experiments, molecular motion investigations and narrow line-widths [27].

Figure 1 – Energy schematic of nuclear spin in an applied magnetic field [28].

In order to detect nuclear spin transitions, a secondary oscillating magnetic field, B1, is applied to the sample. This is done orthogonally to Bo. The B1 field can be applied continuously

4 or in pulses giving rise to two different techniques. However, in terms of protein samples, pulsed

NMR is much more powerful. A simple pulsed NMR experiment consists of an initial short RF pulse followed by detection of the resulting signal. The resultant excited states in the nuclei produce an oscillating magnetic field that induces a current in the receiver coil. This induced current can be measured as a function of time and referred to as the free induction decay (FID).

The Fourier transformation of this FID is then what is referred to as an NMR spectrum. [27]

During evolution or mixing times, the time that follows a pulse, a phenomenon known as relaxation occurs in the excited population. There are two characteristic parameters related to this

NMR relaxation; longitudinal relaxation time (T1) and transverse relaxation time (T2). T1 is the time required for the spin-flipped population to return to thermal equilibrium in the plane of Bo.

This time is dependent on the field strength of the static magnetic field. However, the presence of paramagnetic labels can decrease this relaxation time. T2 is relaxation associated with mutual spin- flips due to interacting nuclei in a sample [27].

Electrons, like nuclei, are charged and rotate with a spin to counter the magnetic field produced by the nucleus. The shell of electrons which surround the nucleus can then exhibit a shielding effect that affects the magnetic field at the nucleus reducing the specific NMR frequency of said nucleus. This coupling of the electron cloud and the external magnetic field causes what is commonly named the chemical shift (δ). Chemical shift is the frequency of the resonance expressed with reference to a standard set as 0 ppm (Eq. 1). The unit, parts per million (ppm), is chosen for ease and does not represent the frequency of the spectrometer. Since the dividend of this equation is in Hz while the divisor is in MHz, ppm is the resultant measurement unit. As such, if the observed frequency in an NMR signal is dependent on the electron density distribution, the chemical shift is indicative of local chemical structure. In general, high shielding results in a lower

5 chemical shift whereas lower shielding results in a higher NMR frequency. It is this property which has made NMR so attractive to chemists and structural biologists [29].

휔−휔 훿 = 푟푒푓 [1] 휔푟푒푓

1.2.1.2 Sample Preparation for SSNMR of membrane proteins

Sample preparation is a critical step in any protein structural study. As natural isotope abundance proteins contain very low amounts of the NMR-active 13C and 15N nuclei, samples have to be isotopically labeled, which can be done either uniformly or selectively. Additionally, samples need to be homogeneous in conformation, to produce narrow linewidth. Also, depending on the experiment and size of rotor, sample sizes of at least a milligram or greater are needed for sufficient signal-to-noise ratios [20]. Expression of membrane proteins in adequate levels for economically feasible NMR studies is challenging mainly due to the cost of isotopically labelled nitrogen and carbon sources, or deuterium in the certain cases [15]. As such, the majority of membrane proteins are expressed in Escherichia coli (E. coli) or cell free systems as they have well established protocols for expression of proteins for NMR [15]. There have been cases of optimized expression of eukaryotic membrane proteins such as GPCRs in E. coli [25]. Strategies include low growth temperature, optimized refolding protocols, or use of fusion proteins to encourage proper insertion into the cell membrane [15]. Nevertheless, many eukaryotic membrane proteins do not express or fold in E. coli. Alternative systems, such as mammalian and insect cells are most native, but have high costs, low yields, and lack protocols for uniform labeling and deuteration. However, some success has been seen in using yeast-based expression systems (mainly Pichia pastoris) for the eukaryotic membrane proteins such as Leptosphaeria rhodopsin (LR) [30] and human aquaporin-

1 [31].

In addition to expression and isotope labeling, a successful purification protocol must be established in conjunction with lipid or detergent optimization depending on the type of NMR experiment to be run. For solution NMR, membrane proteins are generally kept in detergent micelles [32]. In contrast, synthetic lipid bilayers analogous to the cell membrane are used for

SSNMR [33-37]. However, strides are currently being made by some groups to study membrane proteins in the membranes of the expression host which could reveal structure, dynamics, and oligomerization states in a more native environment [18, 38-43].

For most NMR studies, proteins are produced in minimal media. This means that carbon and nitrogen are supplied by single sources such as glucose, or glycerol in the case of carbons [44].

However, this can introduce stress into the host cells, making protein yields lower in comparison to enriched media production [45]. Therefore, it is critical to determine which colonies produce the protein of interest in high amounts and optimize the media contents as needed to improve yields prior to switching to labelled media. By varying the carbon or nitrogen sources, one can introduce labelling schemes within the protein of interest [46]. This can help with assigning chemical shifts to certain atoms depending on how they label, improve spectral resolution, and collect internuclear distance restraints.

1.2.1.3 Multidimensional NMR spectroscopy

In 1971, the first two-dimensional (2D) NMR experiment was recorded by Jeener [29]. For larger molecules, a one-dimensional spectrum is far too complex for interpretation as many of the signals overlap. By introducing another 90° pulse after a defined time interval during which the spins are allowed to evolve, the signal is then detected over a range of times after the second pulse.

This is the simplest case of a 2D experiment resulting in a homonuclear correlation spectrum [20].

This then produces 2D spectra with both diagonal peaks and cross-peaks. Diagonal peaks are

7 located along the central diagonal and correspond to the peaks in a one-dimensional NMR experiment. The cross-peaks are located off the diagonal and indicate couplings between pairs of nuclei. This occurs due to a process called magnetization transfer [20]. The simplest way to determine which couplings a cross peak represents is to find the diagonal peak which is directly above or below the cross peak, and the other diagonal peak which is directly to the left or right of the cross peak. The nuclei represented by those two diagonal peaks are coupled [20]. More complicated experiments include magnetization transfer between different isotopes, for example from 1H to 13C, which are deemed heteronuclear correlations [20]. By adding more pulses, varying evolution times and magnetization transfer steps, one can take a three-dimensional (3D) spectra which can then be used for resonance assignments of larger molecules [20]. Multi-dimensional spectroscopy is essential not only for resonance assignments, where individual peaks are assigned to specific protein atoms, but also for measuring specific structural restraints, which is required for protein structure determination.

1.2.1.4 Solid-state NMR of Membrane Proteins

A significant number of membrane protein structures have been determined using solution

NMR. These include both bacterial porins (β-barrel architecture) [11] and α-helical transmembrane proteins [15]. The first membrane protein determined through NMR in 1993, gramicidin, was very small, with a molecular weight of 4 Da [47]. In recent years, technological improvements have enabled solution NMR structures in proteins in the 30 to 40 kDa range [15]. However, solid-state

NMR (SSNMR) is gaining popularity based on the relative lack of restrictions in terms of sample choice, especially in the study of integral membrane proteins [34-37].

To achieve a usable SSNMR spectra certain experimental procedures can be undertaken to overcome anisotropic interactions. Sometimes, the molecule will automatically orient itself in such

8 a way that will partially average the powder pattern. Macroscopic orientation of membrane proteins reconstituted into lipid bilayers on glass plates in such a way that they are aligned perpendicular to the magnetic field can result in narrower line widths and a distinct orientation dependent pattern will be observed. This is termed oriented SSNMR. It is well-regarded in terms of its ability to probe backbone structure. One can compare theoretical spectra with the experimental to probe predicted topology. One such experiment is PISEMA which consists of two parts: Polarization Inversion (PI) and Spin-Exchange at the Magic Angle (SEMA). PISEMA provides the chemical shift and heteronuclear dipolar coupling interactions, while the SEMA component suppresses the dipole-dipole interaction among protons [34, 35]. Overall, this results in a spectral pattern in oriented samples which provides information on the orientation of the individual N-H bonds with respect to the magnetic field, and can be used to derive the protein backbone structure [48].

1.2.1.5 Magic Angle spinning solid-state NMR

The dipolar interaction occurs when the magnetic field of a nucleus affects another nuclei’s spin. As this interaction is anisotropic (depends on the orientation of the internuclear vector relative to the magnetic field), this can severely broaden the SSNMR spectra in the absence of the averaging effect of tumbling in solution. In 1960 Andrew et al. [49] and Lowe [50] independently demonstrated that the rotation of the sample couple be used to obtain the same averaging as seen in solution NMR. They found that spinning the sample at the magic angle (54.7°) reduces or averages out the anisotropic interactions from nuclei other than protons to zero [51], laying the foundation for what is called Magic Angle Spinning (MAS) SSNMR today. While the samples for

MAS SSNMR still have to be stable for long signal acquisition times as seen for solution NMR, there are no restrictions on solubility, crystallizability, molecular size or even purity. As such,

SSNMR is a powerful tool for larger protein assemblies or aggregates, integral membrane proteins, biominerals, protein-DNA complexes, or even full viruses [17].

1.2.1.6 SSNMR interactions

There are several interactions seen in SSNMR which can be manifested in NMR spectra.

For most cases, assuming a large magnetic field-strength, one can assume that the Zeeman interaction and other external magnetic forces are much greater than internal contributions. In other cases, these perturbations can be accounted for in the Zeeman Hamiltonian. For spin ½ nuclei such as 1H, 13C, and 15N, these forces include chemical shielding, dipolar interactions, and indirect spin- spin coupling [33].

The distance between the two nuclei affects the magnitude of the dipolar interaction. As the distance between two nuclear magnetic moments increases, the interaction between them diminishes as 1/r3, allowing one to estimate the internuclear distances. Chemical shielding anisotropy (CSA) is characterized by a tensor σ, which can be diagonalized to yield three principal components: σiso, Ω and κ [33]. The isotropic chemical shielding (σiso) is the result of averaging by isotropic tumbling in solution. The span (Ω) measures the width of the CSA powder pattern in ppm. The skew (κ) is a measure of the asymmetry of the powder pattern. These values can give rise to shifts in the frequency due to an orientation dependence [33]. In MAS experiments, CSA is averaged out by the rapid spinning of the sample. However, if the spinning frequency is less than the size of the CSA, incomplete averaging will occur, which results in spinning side bands [51].

1.2.1.7 SSNMR derived structure examples

The first G-protein coupled receptor to be structurally solved by SSNMR was CXCR1.

This protein is one of the two high-affinity receptors for the CXC chemokine interleukin-8, an immune response protein [25]. The structure was determined in a liquid crystalline phospholipid

10 bilayer without any modifications and at physiological conditions. This was done using a combination of MAS and oriented NMR known as rotationally-aligned (RA) NMR. RA-SSNMR relies on the inherent rotational diffusion of membrane proteins in a lipid bilayer to average orientation-dependent motional dipolar coupling. The native-like conditions used for this experiment are part of what makes NMR such an attractive tool for investigating membrane proteins. This can be easily manipulated with the addition of different phospholipids or cholesterol to analyze their impact on the dynamics of the protein. Additionally, small molecules such as ligands, can be easily added to the sample allowing direct spectroscopic analysis of their effect both on local residues and the whole protein [25].

Anabaena Sensory Rhodopsin (ASR) is a 7 transmembrane protein with retinal as a prosthetic group. Previous studies have found that light-induced structural changes may alter its interaction with a soluble transducer, which then starts a signal cascade [52-54]. Isotopically labelled ASR expressed in E. coli was reconstituted in DMPC/DMPA liposomes. Initial structural calculations were performed using DARR and heteronuclear correlation experiments [26]. Using a mutant, S26C, the cysteine was paramagnetically labelled enabling Paramagnetic Relaxation

Enhancements (PREs) to be measured. PREs are increasingly becoming a method to provide long- range distance information that can complement DARR restraints, which are limited to distances of less than 5 Å [55]. Distances between the spin label and NMR active nuclei can be determined from the increased T1 or T2 relaxation rates depending on the paramagnetic agent [55]. This method can be used to get distances in the range of 15-24 Å (for a lone spin of a nitroxide stable radical).

The PREs and other experimental data were indicative of trimerization of ASR in the synthetic membranes used for measurement [26, 55]. Due to the nature of SSNMR, the structure of this large complex could be obtained which would not be feasible when using solution NMR.

In some cases, X-ray and solution NMR structures have been shown to conflict. One such case is the M2 protein of the influenza A virus which forms a tetrameric proton channel and has an important role in virus replication. Newly developed resistance to antiviral drugs has made characterization of the drug binding site of interest. Due to the conflicting results from other methods, SSNMR was used to probe the binding site. A series of site-directed isotopic labelling

SSNMR experiments revealed a strong binding site at Ser31 and a weak binding site at the C- terminus. It was determined that the weak binding site is obstructed by an amphipathic helix in the full-length protein confirming the binding site to be Ser31. In the mutant S31N, which was found in recent circulating virus strains, it was established that no chemical shift changes occurred upon addition of the antiviral drug indicative of resistance. As such, a strong correlation between a medically relevant binding site and the data available from SSNMR could eventually prove more useful in future studies [17].

1.2.2 Fourier-Transform Infrared spectroscopy

Another method that can be used to study reconstituted and native membrane proteins is

Fourier-transform infrared (FTIR) spectroscopy. This technique uses vibrations at characteristic group frequencies corresponding to different chemical bond types which can help understand the secondary structure of the protein, sample composition, and chemical environment of certain residues [56]. FTIR was first measured in 1957, covering the wavenumber range of 4000 cm−1 to

660 cm−1. The first commercial spectrometer was produced in 1969 after the computer technology became available. FTIR simultaneously shines multiple frequencies of light at the sample and measures the absorption [57]. The light shines into a Michelson interferometer, a motorized mirror configuration. As mirrors move, each wavelength of light in the beam is periodically blocked, and transmitted by the interferometer, due to wave interference. Different wavelengths are modulated

12 at different rates, so that at each moment, the beam coming out of the interferometer has a different spectrum. Different combinations of light frequencies are then used while the computer determines the exact absorption at each wavelength [57]. The advantages of FTIR spectroscopy compared to the traditional dispersive one include an improved signal-to-noise ratio, an accurate wavenumber scale and higher IR light throughput [57]. Another advantage of this technique is the wide variety of environments in which the protein can be studied including crystals, lipid membranes and organic solvents [56]. Proteins typically present a spectrum as seen in Figure 2.

Figure 2 - Example of a typical static FTIR spectrum of a membrane protein [56]

Infrared active molecular vibrations must be associated with a change in the dipole moment. A molecule can vibrate in many ways, and each way is called a vibrational mode. For molecules with N number of atoms in them, linear molecules have 3N – 5 vibrational modes, whereas nonlinear molecules have 3N – 6 vibrational modes. As an example H2O, a non-linear molecule, will have 3 × 3 – 6 = 3 modes. Simple diatomic molecules have only one bond and only one vibrational band. In the case of FTIR, this includes asymmetrical diatoms. However, if the

13 molecule is symmetrical, the band is not observed in the IR spectrum, but only in the Raman spectrum [58]. Vibrational spectra become increasingly more complex as the molecules gets larger, as seen in a protein or proteoliposome spectrum [56]. The atoms in a CH2X2 group, commonly found in organic compounds and where X can represent any other atom, can vibrate in nine different ways. Six of these involve only the CH2 portion: symmetric and antisymmetric stretching, scissoring, rocking, wagging and twisting. The rocking, wagging, and twisting modes do not exist for H2O, since they are rigid body translations and no relative displacements exist

[58].

Table 1 – Characteristic absorption wavelengths in IR spectroscopy. Signal strength is represented by s (strong), m (medium), or v (weak) [59]. Functional Group Characteristic Absorption(s) (cm-1) Alkyl C-H Stretch 2950 - 2850 (m or s) Alkenyl C-H Stretch 3100 - 3010 (m) Alkenyl C=C Stretch 1680 - 1620 (v) Alkynyl C-H Stretch ~3300 (s) Alkynyl C=C Stretch 2260 - 2100 (v) Aromatic C-H Stretch ~3030 (v) Aromatic C-H Bending 860 - 680 (s) Aromatic C=C Bending 1700 - 1500 (m,m) Alcohol/Phenol O-H Stretch 3550 - 3200 (broad, s) Carboxylic Acid O-H Stretch 3000 - 2500 (broad, v) Amine N-H Stretch 3500 - 3300 (m) Nitrile C=N Stretch 2260 - 2220 (m) Aldehyde C=O Stretch 1740 - 1690 (s) Ketone C=O Stretch 1750 - 1680 (s) Ester C=O Stretch 1750 - 1735 (s) Carboxylic Acid C=O Stretch 1780 - 1710 (s) Amide C=O Stretch 1690 - 1630 (s) Amide N-H Stretch 3700 - 3500 (m)

Infrared spectroscopy is typically used to identify structures since different conformations of functional groups result in characteristic bands in both intensity and position. When proteoliposomes are studied through FTIR the spectra show a carbonyl stretch group peak for lipid esters at 1700-1800 cm-1 [30]. The peak between 1600 and 1700 cm-1 (amide I) corresponds to the

C=O stretching vibrations of the backbone with the peak position dependent on the secondary

14 structure of the protein [6]. In comparing lipid peak amplitude with the amide I peak, it is possible to determine the protein to lipid ratio in the proteoliposomes [30, 60]. The amide II peak corresponds to the backbone C-N stretching coupled with in-plane N-H bending in the range of

1510 to 1580 cm-1 [6], and can be used to probe secondary structure by H/D exchange. FTIR is also extremely useful in the preparation of samples for SSNMR as the proteoliposome stability can be tested. Also, the extent of isotopic labelling of the protein can be estimated from the amide peak shifts. The amide I peak will shift to lower wavenumbers when proteins are 13C-labelled while the amide II peak will downshift when proteins are 15N-labelled [30]. Various protein and lipid combinations can be tested using FTIR to determine the optimum composition of the proteoliposome for each protein.

1.2.3 Resonance Raman spectroscopy

Raman spectroscopy, as named after C.V. Raman, is a spectroscopic technique used to observe vibrational, rotational, and other low-frequency modes in a system [61]. Raman spectroscopy is commonly used in chemistry to provide a fingerprint by which molecules can be identified. It relies on inelastic scattering of monochromatic light, usually from a laser in the visible, near infrared, or near ultraviolet range [61]. The laser light interacts with molecular vibrations, phonons or other excitations in the system, resulting in the energy of the laser photons being shifted up or down. The shift in energy gives information about the vibrational modes in the system. Electromagnetic radiation from the illuminated spot is collected with a lens and sent through a monochromator. Elastic scattered radiation at the wavelength corresponding to the laser line (Rayleigh scattering) is filtered out by either a notch filter or a band pass filter, while the rest of the collected light is dispersed onto a detector [62]. While Raman spectroscopy gives information about the vibrational modes of the molecules similar to FTIR, the selection rules are

15 different (dipole moment change vs. polarizability change), so that information provided by

Raman can be complementary.

Resonance Raman spectroscopy is a useful tool for probing chromophores in biological systems. The wavelength of the laser beam can be set to the electronic transition of the chromophore within a complex biological matrix such as a protein [63]. As such, one can probe specific features of conjugated molecules (e.g., aromatic amino acids, DNA, carotenoids) within larger proteins. Historically, this technique has been used frequently for analysing the retinal molecule inside both visual rhodopsin and members of the microbial rhodopsin family such as bacteriorhodopsin (BR) [64]. 1.3 Summary of Goals

This study seeks to investigate various aspects of the preparation of integral membrane proteins for SSNMR. The proteins included in this study include human adenosine receptor (2A) (A2AR),

Anabaena sensory rhodopsin (ASR), and green-light absorbing proteorhodopsin (PR). Factors such as host selection, truncation, and protein-ligand interactions will be explored. These proteins share two main features: seven transmembrane alpha-helical architecture and the ability to bind certain ligands for signal transduction. Using the biophysical techniques herein identified,

SSNMR, FTIR and Raman spectroscopy, one can characterize how the proteins are expressed and the inherent challenges associated.

Chapter 2 will outline the production of A2AR for SSNMR. In Chapter 3, one will find a brief introduction into microbial rhodopsins, most notably bacteriorhodopsin, to which ASR and PR are homologous. Chapter 4 will outline the study into the expression of ASR in its full length form and the challenges associated with its expression and subsequent purification. Chapter five will

16 explore the biosynthesis of retinal in E. coli for isotopic labelling of the ligand in PR producing cells.

Chapter 2 Production of Adenosine Receptor (2A) for SSNMR

2.1 G-protein Coupled Receptors

G-protein coupled receptors (GPCRs) are integral membrane proteins found in a variety of eukaryotic organisms [7]. GPCRs are a large family of membrane proteins involved in the transduction of signals across the phospholipid bilayer. Processes in which GPCRs are involved include vision, taste, smell, neurotransmission, immune response, and autonomic nervous system responses [65]. As such, the medical community is especially interested in the study of GPCRs as they comprise approximately 30 to 50% of drug targets for human diseases [66]. Approximately half of all medications work on GPCRs including beta blockers, antihistamines and various psychiatric medications. However, structural studies of GPCRs to date have only determined the high resolution structures of 25 proteins of this class (Figure 3).

The first GPCR structure, determined in 2000 by Krzysztof Palczewski, was that of bovine rhodopsin, originally discovered in 1879 by Franz Christian Boll, the protein associated with vision

[67]. Upon illumination, the covalently bound retinal in the transmembrane domain isomerizes triggering conformational change of rhodopsin which activates the downstream visual transduction cascade [67]. In 2012, the Nobel Prize in Chemistry was awarded to Robert J. Lefkowitz and Brian

K. Kobilka for mapping how the β2-Adrenergic Receptor (β2-AR) works based on their shared work from the 1970’s to present day [68]. This was done through a series of x-ray crystallography experiments. Further experiments using selective labelling and solution NMR have resulted in more insights into the mechanism for GPCR function [69].

Typically, a GPCR is classified according to the ligand or signalling molecule which binds the receptor with specificity and high affinity [70]. There are 6 classes of GPCRs [71]:

Class A: Rhodopsin-like Class B: Secretin receptor family Class C: Metabotropic glutamate/pheromone Class D: Fungal mating pheromone receptors Class E: Cyclic AMP receptors Class F: Frizzled/Smoothened

The majority of GPCR structures determined to date belong to the rhodopsin-like class. This is mainly due to the size of this class and their importance to the pharmaceutical industry. The largest class, A, comprises 85% of GPCR genes and are frequently drug targets. This class is further divided into five categories: opsin, amine, peptide, cannabinoid, and olfactory receptors [72]. The largest of them is odorant receptors (ORs) which are generally located at the ciliated surface of olfactory sensor neurons and comprise 3% of the human genome [73]. These proteins display affinity for a range of ligands, and conversely a single ligand molecule may bind to a number of

GPCRs with varying affinities [74].

Given that GPCRs are frequent targets of drug development, it is advisable to fully characterize the function and structure of every available protein. This includes the ligand binding site location, the conformational change induced, and where the G-protein binds. It should be noted that in order to experimentally determine these parameters, one must have a reliable method for producing various mutants of the receptor, as well as the G-protein and the ligand molecule.

Additionally, knowledge of the primary sequence of the protein and the structure of the ligand would be necessary, as these would allow homology modelling and identification of putative residues of functional importance [13].

Figure 3 – GPCR structures determined according to their classification. Structures are identified with red flags. GPCR classes are represented in blue (Class A), red (Class B), yellow (Class C), purple (Class D), and green (Class F) [75].

2.1.1 GPCR Structure

The general structure of a GPCR consists of seven transmembrane helices with an extracellular N-terminus and an intracellular C-terminus [66]. This is analogous to bacterial rhodopsins which have been well studied in the past. A high level of structural homology can be seen between different GPCRs. Considering their ubiquity in cellular function, this homology implies a common transduction mechanism. The GPCR system consists of three main components: the receptor, the ligand, and the transducing G-protein [73].

Typically, each transmembrane helix is approximately 20 residues in length and the helices bundle together to form the transmembrane domain [73]. Most often, the receptor contains an

20 intramembrane ligand binding cleft and an intracellular G-protein binding domain [73].

Superposition of twelve known x-ray crystallography-derived GPCR structures, shows high conservation in the transmembrane region with variable loops (Figure 4). Frequently, there is an intracellular interfacial eighth helix which adds instability to non-membrane bound GPCRs and is prone to proteolytic cleavage [76].

Unlike bacterial rhodopsins, GPCRs have long loop regions which can affect stability and thereby make GPCRs harder to work with [77]. Extracellular loops are prone to glycosylation which can affect downstream analysis and stability in solution. In addition, there are two highly conserved cysteine residues which form stabilizing disulfide bonds located in the loop regions. A guanine-nucleotide exchange factor (GEF) domain is found primarily formed by intracellular loops

2 and 3. The second extracellular loop has been identified as the ligand binding site ‘lid’ wherein the variability within the loop regions can reflect the ligand which binds the receptor. However, this affects the possibility of homology modelling in these regions, which encourages determining the structure for each GPCR experimentally [65].

Figure 4 – Superposition of twelve crystallized GPCR family members. Rhodopsin (orange), β2-adrenoceptor (magenta), β1-adrenoceptor (cyan), adenosine A2a (pink), dopamine D3 (brown), chemokine CXCR4 (green), histamine H1 (mustard), muscarinic acetylcholine M2 (brick red), muscarinic acetylcholine M3 (grey), δ-opioid (red), μ-opioid (purple), and S1P1 (blue) are depicted [78].

GPCRs are highly affected by post-translational modifications and are commonly found in hetero-oligomeric complexes. This may be due to phosphorylation of the Ser or Thr residues on the C-terminus. Scaffolding protein such as β-arrestins have high affinity to these phosphorylated residues and sterically prohibit homo-oligomerization and recruit additional proteins to a signalling complex [79]. Furthermore, palmitoylation of sites on the C-terminus helps target the protein to cholesterol and sphingolipid-rich domains. This facilitates rapid signal transduction as GPCR associated transducers are also associated with these lipid rafts [80].

2.1.2 GPCR signal transduction mechanism

The GPCR signal transduction mechanism has been well studied in a number of proteins, with β2-Adrenergic receptor being one of the most studied. Previous studies have found that when bound by a ligand, the receptor changes its conformation, resulting in a 14 Å outward movement in helices 5 and 6 in the cytoplasmic domain (Figure 5), which allows it to function as a guanine nucleotide exchange factor [69]. In the case of β2-Adrenergic receptor, when the GDP bound by the G-protein is exchanged for a GTP, the α subunit of the G-protein is released and activates a signal cascade within the cell, typically but not always through cyclic AMP (cAMP). The G- protein exhibits GTPase function, ergo when the GTP is dephosphorylated, the α subunit returns to the receptor and the ligand is released [81]. The active site of the receptor can be desensitized by phosphorylation leading to β-arrestin binding. This prevents interactions with multiple G- proteins [79]. However, depending on the type of receptor and the signalling pathway it interacts with, the components of the G-subunit will vary (Figure 6).

Inactive Active

Figure 5 – Signal transduction mechanism in β2-AR [82] showing the conformational change in the molecules upon agonist activation [23].

2.1.3 GPCR signalling pathways

GPCRs can be found in a wide variety of signalling pathways. There are three main G- protein signalling pathways (Gα, Gβγ, and G-protein independent) mediated by four subclasses of

G-proteins (Gαs, Gαi/o, Gαq/11, and Gα12/13) [83]. These pathways are shown in figure 6. While most GPCRs are capable of activating more than one Gα-subtype, they also show a preference for one subtype over another. Functional selectivity occurs when the subtype activated depends on the type of bound ligand. However, the ligand may be capable of stabilizing multiple conformation of the GPCR’s GEF domain. This could result in interactions with different subclasses of G-proteins but this is subject to availability of the G-protein isoform. Furthermore, feedback pathways may result in receptor modifications (e.g., phosphorylation) that alter the G-protein preference.

However, GPCRs are generally classified according to their preferred binding partner under experimental or physiological conditions [83].

The Gα pathway can activate three different types of effectors. Gαs and Gαi/o act on adenylate cyclase in a process which will be discussed further later [84]. Gαq/11 acts on phospholipase C-β (PLCβ), which catalyzes the cleavage of membrane-bound phosphatidylinositol 4,5-biphosphate (PIP2) into the second messengers inositol (1,4,5) trisphosphate (IP3) and diacylglycerol (DAG). Elevated intracellular calcium levels in turn affect allosterically activated enzymes such as Ca2+/calmodulin-dependent kinases [85]. Finally,

Gα12/13 affects the three RhoGEFs, which in turn activate Rho, a small cytosolic GTPase which in turn can affect Rho-kinase proteins [86].

Gβγ signalling is generally found to affect ion channels. These include G-protein-regulated inwardly rectifying K+ channels (GIRKs) [87], P/Q- and N-type voltage-gated Ca2+ channels [88], as well as some isoforms of AC and PLC, along with some phosphoinositide-3-kinase (PI3K) isoforms [89].

Finally, there are some signalling pathways affected by GPCRs which are independent of

G-proteins. Proteins associated with GPCRs such as scaffolding proteins [79] can acts as signal transducers especially in the case of subcellular localization [90]. Frequently, the effecter seen is a member of the MAPK family [80].

Figure 6 – GPCR signalling overview containing each canonical pathway. Pathways are coloured according to their G-protein subunit and downstream pathway components [91].

2.2 Adenosine Receptor Type 2A

Human adenosine receptors are a type of GPCR that regulate intracellular levels of cyclic adenosine monophosphate (cAMP) [92]. Adenosine, a purine nucleoside, is involved in numerous cell functions including cell proliferation, vascular regeneration and immune functions [92]. To date, there have been four identified types of human adenosine receptor: A1R, A2aR, A2bR and

A3R [93]. A1R and A3R exhibit inhibition on adenylate cyclase through the activation of the Gi subunit (Figure 7). A2aR and A2bR activate adenylate cyclase through the Gs subunit of the G- protein (Figure 7). As such, each receptor depending on tissue localization will have very different effects on the cell. These receptors have been shown to mediate the variety of anti-inflammatory effects of adenosine [93].

Figure 7 – Adenosine receptor signalling mechanism for the four known subtypes: A1, A3, A2B and A2A [94].

The A2a receptor (A2aR), the protein being expressed for this study, is abundant within the basal ganglia, vasculature, T lymphocytes and platelets [95]. It is frequently found in membrane receptor complexes with dopamine or glutamate receptors, leading to integrated functionality [96].

In response to stress, extracellular concentrations of adenosine rise, activating A2aR in response.

When bound by an adenosine molecule, the receptor changes its conformation and the bound Gs protein is released, activating adenylate cyclase which in turn increases the endogenous cAMP level which, through a signal cascade, results in various site-specific effects. These include reduction of inflammation, regulation of dopamine and glutamate levels, and vasodilation of coronary arteries [95]. This in turn inhibits immune responses against tumors by preventing cell- mediated toxicity and suppressing interferon-γ production [92]. Current studies are also analysing

A2aR as a potential drug target for Huntington’s disease and Parkinson’s disease [7].

A2aR is highly unstable in solution; this has been attributed to the inherent structural flexibility of the protein, resulting in multiple conformations which may interact to form aggregates. The C-terminal eighth helix found in the cytosolic domain of the protein has been

27 found to be very unstable. This has led to previous work being done on a truncated construct as opposed to the full length protein. Additionally, the use of detergents removes some lipids which provide stability to the protein. Numerous strategies have been employed to account for this, including inverse agonist binding, lipid or lipid-like substances such as cholesteryl hemisuccinate or glycerol, and site-directed mutagenesis. These keep A2aR in its inactive conformation and stable [97].

2.2.1 A2aR Structure

A2aR has previously been characterized via x-ray crystallography studies similar to those used for β2-Adrenergic receptor. First, a truncated construct fused to T4 Lysozyme was used for

X-ray crystallography. The structure was originally obtained in 2008 for a truncated construct in the presence of the inverse agonist ZM241385. Since, 6 more crystallographic structures have been obtained for A2aR. These have been obtained from truncated constructs expressed in E. coli [97],

P. pastoris [98], S. cerevisiae [99], Sf 9 cells [100] and Sf 21 cells [101]. Current structures show

A2aR having seven transmembrane helices with an eighth helix on the cytoplasmic side between residues Arg296 and Leu308 of the protein (Figure 8).

Several features have been elucidated from these studies. These include the lack of palmitoylation sites which are typically seen in GPCRs. Instead helix VII is stabilized by interactions with helix I [100]. Deviations in the helix structure relative to β2-Adrenergic receptor of RMSD 2A are seen. These differences are pronounced at the extracellular portions of helices I,

II, III, and V. These differences seemed linked to the ligand-binding pocket [100]. The presence of a Trp residue near the ligand binding pocket is purported to be key in shifting between the active and inactive states of GPCRs. This residue (Trp246) has a large contact area with ZM241385,

28 which could be linked to the strength of the ligand binding [100]. A kink in the extracellular loop

3 formed by disulfide linkages, constrains the loop at the top of the ligand binding site. The ligand binding pocket is located closer to the extracellular surface than previously seen in rhodopsin or

β2-Adrenergic receptor. Furthermore, His250 has been identified as a key residue in ligand binding, specifically of ligands containing the furan ring feature such as ZM241385 [100]. As such, it is clear that ligand binding sites in GPCRs are key distinctive structural features.

Similar to β2-AR, X-ray crystallography has elucidated some conformational changes in the protein upon agonist activation. In helix III, there is a highly conserved E/DRY motif which forms an ionic lock with the N-terminus of helix VI. This lock is broken in the activated receptor resulting in an outward movement of the helix VI [101]. This was observed when activated by an agonist (NECA), wherein the receptor formed a 5-6 Å cleft between helices 5 and 6 where the C- terminus of the G-protein can bind [102].

Figure 8 - Structure of A2aR bound by an NECA molecule (red) in the active state. Protein is visualized using Pymol (PDB ID: 3VGA) 29

2.2.2 Oligomerization of A2aR

In its native environment, A2aR has been found to form both homo- and hetero-oligomers.

Using FRET based assays, the existence of higher-order A2aR oligomers in the plasma membrane, the recognized site of action of A2aRs, has been proven [103]. In addition to forming homodimers,

A2a receptors are known to associate in heteromeric complexes, notably with adenosine A1 [93], dopamine D2 [104], cannabinoid CB1 [105], and glutamate mGlu5 receptors [106].

It has been suggested that this oligomerization is related to the fifth transmembrane helix

(TM5). When M193 located on TM5 is replaced with an alanine residue, there is a significant reduction in dimerization [107], indicative of this residue being a crucial component of A2aR association (Figure 9 A). In previous purification studies homo-oligomers have been seen in both mammalian cell lines as well as Pichia pastoris expressed A2aR (Figure 9 B-D). Depending on the construct used and the mode of purification, A2aR has been found to form higher order oligomers [108]. However, when purified in a two-step process including both Flag-tag affinity purification and TEV protease cleavage followed by His-tag metal affinity purification, this oligomerization was significantly reduced (Figure 9 D).

A B C D

Figure 9 – Formation of oligomers in heterologously produced A2aR. A) Reduction in dimerization of A2aR due to point mutation in helix V [107]. B) Detection of A2aR oligomers after solubilization of protein expressed in HEK 293 [103]. C) Detection of A2aR homo-oligomers in HEK 293 cells using Flag-tag detection at the cell surface [103]. D) Reduction in oligomers in A2aR expressed in P. pastoris and purified through Flag-tag affinity [2] followed by TEV cleavage and metal affinity purification [3] [108].

2.2.3 Ligand Binding

Ligand binding is a crucial component of GPCR function. Various antagonists and agonists have been found for A2aR. Adenosine, a natural agonist, and NECA, a synthetic agonist, have been used for structural studies using a mutant form of A2aR to increase thermostability for structural studies [102]. Other agonists include: DPMA and Limonene. Other antagonists include: theophylline, caffeine, and ZM-241385 [109]. By introducing high affinity ligands, one can ‘lock’ the protein in stable conformations during structural parameter acquisitions [69]. In this way detailed structural information potentially useful to the pharmaceutical community can be determined.

2.3 Heterologous Expression in Pichia pastoris

Eukaryotic membrane proteins pose more of a difficulty for functional expression than prokaryotic membrane proteins [110]. Eukaryotic proteins have the increased complexity of post-

31 translational modifications and complex folding mechanisms which are not supported by the E. coli expression system [111]. This is exemplified by the poor expression of GPCRs in E. coli based systems [112]. A viable alternative to E. coli expression systems for eukaryotic membrane proteins is a methylotrophic yeast expression system Pichia pastoris. This system has been successfully used to study various soluble eukaryotic proteins as well as produce membrane proteins for crystallization [113]. The ability of P. pastoris to grow to high cell densities and its tightly regulated promoter make it an ideal system for the expression of eukaryotic proteins [110].

Previous work conducted at the University of Guelph has shown the feasibility of P. pastoris for the expression of isotopically labelled Leptosphaeria rhodopsin (LR), a fungal rhodopsin, for

SSNMR [30]. LR, which is structurally similar to GPCRs, showed high expression in P. pastoris and remained stable and functional after reconstitution into liposomes [30]. This showed great promise for the use of P. pastoris as an expression system for other eukaryotic membrane proteins such as aquaporin (AQP) [31, 108] and human adenosine receptor [108].

2.3.1 Comparison to other expression hosts

P. pastoris has some key features compared to other typically used expression hosts which make it a viable host for A2aR. Since yeast is eukaryotic, it contains the machinery to perform post-translational modifications to the protein. In E. coli, without these modifications, misfolding is typically seen resulting in a non-functional protein [111]. As such, despite the relatively low cost and ease of E. coli cultures in the case of isotopic labelling, a eukaryotic host is preferable to produce A2aR [30]. Higher order eukaryotic hosts such as mammalian or insect cells could be used to produce proteins such as A2aR. However, lower protein yields have been seen as compared to Pichia. Additionally, they require complex rich media which is expensive for isotopic labelling

[114].

Yeast is one of the simplest eukaryotic host organisms. P. pastoris is one of two regularly used yeast strains for heterologous expression [114]. Saccharomyces cerevisiae (Baker’s yeast) is the other. While S. cerevisiae is widely used and would result in a functional protein, it would not be ideal for producing NMR samples. P. pastoris can grow to higher cell densities and can use methanol as its sole carbon source. This results in a much larger protein yield for relatively less expensive media. This would allow isotopic labelling under economically feasible conditions which is needed for NMR studies. Multiple clones can be generated and screened in a matter of a few weeks making this similar to E. coli expression in terms of speed [113].

2.3.2 How Does Pichia Expression Work (AOX1)

Usually the gene for the desired protein is introduced under the control of the AOX1 promoter. Consequently, the presence of methanol induces the production of the protein of interest.

In most Pichia expression vectors, the protein of interest is produced as a fusion product to the secretion signal of the α-mating factor from S. cerevisiae. This causes the protein to be secreted into the growth medium, which greatly facilitates subsequent protein purification. However, for membrane proteins, this α-factor has been shown to improve targeting of the protein to the endoplasmic reticulum for insertion [115]. This α-factor is later cleaved during post-translational processing [116].

Pichia pastoris has two alcohol oxidase genes, Aox1 and Aox2, which have a strongly inducible promoter. AOX is the first enzyme in the methanol utilization pathway, as such these genes allow Pichia to use methanol as both a carbon and energy source [117]. There exists different strains of Pichia in which the regulation of these genes is modified for heterologous expression.

In the case of a Mut +, both the AOX1 and AOX2 genes are present and functional. Methanol metabolism is identical to the wild-type strain. The Mut s has the AOX1 gene disrupted and ergo

33 the AOX2 gene is solely responsible for methanol metabolism. This results in a significantly slower growth rate, about 5% to 10% of the activity compared to AOX1. In the case of a Mut – strain, Pichia is unable to metabolize methanol [117].

One such Mut+ strain, which has been used extensively for the expression of GPCRs is the

SMD1163 strain (his4 pep4 prb1). The histidinol dehydrogenase gene (HIS4) has been altered in this strain to allow for the selection of transformants with the His4 gene restored by the inserted plasmid. Additionally, SMD1163 is also a protease deficient strain, lacking the active genes coding for proteinase A (PEP4) and proteinase B (PRB1) [118]. Due to reduced proteolytic activity, previous work has shown that use of this strain has resulted in higher expression levels of GPCRs

[7].

2.4 Studies on fermenter produced C-terminally truncated A2aR

2.4.1 Materials and Methods for Analysis of 15N-334-A2aR

2.4.1.1 Reconstitution of 15N-334-A2aR

Previously, 15N-labeled and natural abundance A2aR as produced in its C-terminally truncated (at the position of Val334, the full length being 412 amino acids) form in Pichia cultures was provided by the group of B. Byrne, Imperial College London. The coding sequence was preceded by FLAG-tag, 10xHis-tag, and TEV cleavage site in the pPIC9K vector, transformed into SMD1163 strain. Expression was produced in fermenter cultures which require the constant addition of a carbon source which is cost prohibitive for 13C-labelling. The concentration of the protein samples provided by the Imperial College London group (S. Singh) was verified by

Bradford protein assay (Bio-rad DC protein assay) prior to lipid reconstitution for SSNMR studies.

The liposomes outlined in Table 2 were prepared using natural abundance 334-A2aR. Dry powder lipids were first dissolved and mixed in warm chloroform at the designated weight ratios. The

34 chloroform was removed through vacuum evaporation to form a thin lipid film. The lipids were then thoroughly dehydrated under vacuum desiccation for 5 hours. The lipids were rehydrated with reconstitution buffer (5 mM NaCl, 10 mM Tris, pH 8) to form a 12.1 mg/mL suspension. The reconstitution buffer in this experiment included the addition of ZM241385 (Tocris) at a concentration of 100 nM to maintain the inactive conformation of A2aR.

Table 2 – Liposomes prepared for reconstitution and protein to lipid ratios (P/L) in prepared liposomes Lipids Lipids Ratio P/L PC/PS/Cholesterol 9:1:2.3 1:2 9:1:2.3 1:1 9:1:2.3 2:1 9:1:2.3 4:1 9:1:1 2:1 4:1:1 2:1 4:1:0.5 2:1 DOPC/PS/Cholesterol 9:1:1 2:1 DOPC/Cholesterol 9:1 2:1

Samples for both the natural abundance 334-A2aR and 15N-334-A2aR were prepared according to the same procedure. The solubilized proteins and lipids in 0.05 % DDM were combined at a protein to lipid to Triton X-100 ratio of 1 :1 :0.5 (w/w/w) and incubated at room temperature for 1 hour. To withdraw the detergent and form the proteoliposomes, Bio-beads SM-

2 (Bio-rad) were added at 0.6 g per mL of mixture and incubated at 4 °C with mixing for 24 hours.

The proteoliposomes were collected using a syringe and the beads were washed in 200 mM NaCl until the turbidity of the solution after incubation was negligible. The collected proteoliposome suspension was centrifuged at 150,000 ×g, 4 °C for 50 minutes. The pelleted proteoliposome was washed several times in NMR buffer (10 mM NaCl, 25 mM TrisCl, pH 8) and stored at 4 °C.

For the natural-abundance samples, FTIR was used to optimize the protein to lipid ratio, cholesterol content, and lipid type. After the lipid optimization, the 15N-A2aR was reconstituted

35 into the PC/PS/Cholesterol (9:1:2.3) liposomes at a ratio of 1:1 (P/L). This sample was then used for both FTIR and SSNMR. FTIR experiments were used to assess the stability of the sample over time and how the protein responds to different storage conditions (4 °C or -20 °C). Based on the

SSNMR spectra of PC/PS/Chol based proteoliposomes, further samples of 15N-A2aR were reconstituted at a protein to lipid ratio of 2:1 in DOPC/PS/Cholesterol (9:1:1) liposomes.

2.4.1.2 Preparation of samples for SSNMR

The washed isotopically-labelled proteoliposomes were centrifuged at 900,000 ×g for 3 hours with 20 µL of NMR buffer. The samples were then center-packed into a 3.2 mm NMR rotor.

Experiments were run on Bruker Avance III spectrometer at the frequency of 800.230 MHz. For

D2O incubation, the sample in the rotor was soaked overnight at 4 °C in D2O buffer.

2.4.2 FTIR Analysis of 15N-334-A2aR

A2aR was reconstituted into PC/PS/Cholesterol (9:1:2.3) liposomes at four different theoretical protein to lipid weight ratios: 1:1, 1:2, 2:1 and 4:1. FTIR absorption spectra of the proteoliposomes displayed lipid ester peaks at the vibration frequency of 1732 cm-1 (Figure 10).

The amide I peaks for the samples were located at 1653 cm-1 (as expected for mainly α-helical protein) and the amide II peaks were located at 1541 cm-1 (Figure 10). By comparing the relative amplitudes of the lipid ester peak to the amide I peak, it was estimated that the experimental protein to lipid ratios are close to the theoretical ratios (Figure 10). The relative amplitudes of the amide I peak to the amide II peak in each of the samples were consistent at 3:1 (Figure 10). As the protein content of the proteoliposomes increased, relative decrease of the lipid phosphate peaks at 950-

1250 cm-1 was evident (Figure 10). These series demonstrated that A2aR can be reconstituted into synthetic lipids at high protein-to-lipid ratios required for SSNMR on par with those used

36 previously used for microbial rhodopsins without any effect on the secondary structure of the protein.

0.16

0.14

0.12

0.10

0.08

0.06

0.04

0.02

0.00

Figure 10 – Baseline-corrected static FTIR spectra of A2aR reconstituted into PC/PS/Cholesterol (9:1:2.3) liposomes at different P/L ratios (1:2, 1:1, 2:1, 4:1). Absorbance measured on Bruker IFS66vs machine with a temperature-controlled sample holder (Harrick) connected to a circulating water bath (Fisher). The pellets were resuspended in 600 μl of dH2O and 100 μl of the suspension was dried onto the CaF2 window.

As SSNMR experiments can run for many days, protein stability at an experimental temperature (4 °C) was tested. The sample of A2aR reconstituted at a protein to lipid ratio of 1:1 was initially measured prior to storage at 4 °C and measured after a two week interval. The relative amplitudes of the amide I peak and the amide II peak did not differ between the two samples stored at 4 °C (Figure 11). Overall, the loss of amplitude in the protein after storage at 4 °C was less than

15% (Figure 11). There were some shifts in the amide I and II bands suggesting a decrease in helicity, but overall the structure seemed to remain stable (Figure 11). Therefore, A2aR was a good candidate for SSNMR studies given the relative stability at the experimental SSNMR temperature.

Figure 11 – Baseline-corrected static FTIR spectra of A2aR reconstituted into a PC/PS/Cholesterol (9:1:2.3) liposomes measured at a two week interval in 4 °C. Absorbance measured on Bruker IFS66vs machine with a temperature-controlled sample holder (Harrick) connected to a circulating water bath (Fisher). th Approximately 1/10 of the sample (0.56 mg) was loaded on CaF2 window for measurement.

Natural abundance A2aR was reconstituted into additional six types of liposomes (Table 2).

The average C=O stretching vibration for the lipid esters was 1730 cm-1. The amide I peak for the samples averaged at a vibration frequency of 1653 cm-1. The average vibration frequency for the amide II peak was 1544 cm-1. Samples in PC/PS/Cholesterol (9:1:1) liposomes showed lower lipid content and exhibited low spectral signal in FTIR. This is most likely attributable to A2aR having a high cholesterol affinity which affects the packing efficiency when cholesterol is decreased to

9% from 20%. Additionally, improved lipid retention occurred with DOPC based proteoliposomes, presumably due to the increased saturation of the fatty acyl chains.

2.4.3 SSNMR of 15N-334-A2aR

Initially, the 15N-A2aR sample was reconstituted into PC/PS/Cholesterol (9:1:2.3) liposomes at a protein to lipid ratio of 1:1. After being packed into a 3.2 mm NMR rotor, the sample did not have sufficient protein content for good spectral quality. Another 1 mg sample of

15N-A2aR was reconstituted at a protein to lipid ratio of 2:1. This improved the spectral quality with visible peaks relating to specific amino acids on the protein backbone peak located at 120 ppm (Figure 12). Next, we found that the protein backbone peak was narrower and had more resolved structure in the DOPC/PS/Cholesterol sample (Figure 12). Comparatively,

DOPC/PS/Cholesterol (9:1:1) based proteoliposomes showed an improvement in both the signal by 10-15% and the line width by 20% (Figure 12). This is indicative of more homogeneous structural conformation within the sample. This could be due to the multiple unsaturated fatty acids in DOPC as compared to PC. These saturated fatty acids have reduced mobility in membranes as compared to unsaturated fatty acyl chains. Consequently, the protein sample will be less likely to have slight conformational changes between monomeric units.

Figure 12 – 1D SSNMR spectra of 15N-334-A2aR. Samples were reconstituted into PC/PS/Chol (9:1:1) liposomes (green) and DOPC/PS/Chol (9:1:1) liposomes (black) at a protein to lipid ratio of 2:1 (w/w). PC/PS/Chol was scanned 16456 times while DOPC/PS/Chol was scanned 48000 times.

In attempt to improve spectral resolution, the DOPC/PS/Chol (9:1:1) proteoliposomes were incubated overnight with D2O. D2O incubation results in the exchange of H for D on all exchangeable sites, notably in the amide hydrogens in solvent accessible regions. This should improve the spectral quality by dilution of the proton bath and by the weakening of signals from the exchangeable residues. Improved spectral resolution in peaks possibly relating to Cys and Ala, the amino acids identified at members of β-structure by previous X-ray structures, was seen at

123.3 ppm and 125.6 ppm (Figure 13). Improved resolution pertaining to α-helical structures was seen between 100 and 120 ppm (Figure 13).

Figure 13 – 1D SSNMR spectra of 15N-334-A2aR reconstituted into DOPC/PS/Chol liposomes. Spectra were taken before (black) and after (purple) D2O incubation.

In comparison with previously obtained 15N-spectra of PR and LR, there is comparatively lower peak resolution in 334-A2aR (Figure 14). However, there is enough resolution to justify spectroscopy in the second and third dimension. There is agreement in the location of the amide backbone between the three proteins which confirms the seven-transmembrane alpha-helical architecture within the 334-A2aR sample (Figure 14). Overall, it appears that A2aR is an excellent candidate for SSNMR studies. However, the development of a suitable protocol to produce uniformly 13C, 15N labelled A2aR in an economically feasible manner is critical and we undertook a study to express A2aR in shaker flasks based on the construct previously used for X-ray crystallography [98].

Figure 14 – 1D SSNMR spectra of 15N-334-A2aR (black), Proteorhodopsin (green), and Leptosphaeria rhodopsin (red).

2.5 Materials and Methods for Shaker Flask A2aR Production

2.5.1 Plasmid

The gene containing either the truncated form of A2aR (Trunc-A2aR) or the full-length construct (FL-A2aR) were inserted in the plasmid pPic9K (Figure 15). This plasmid contains the gene encoding resistance to Geneticin, a kanamycin derivative. As such, successful transformants will grow on media containing this antibiotic. For Trunc-A2aR, the gene was truncated at residue

316, removing the majority of the C-terminal tail which has been shown to exhibit instability.

Previous studies were done on a construct truncated at residue 334 and provided by Imperial

College London. However, since this construct was optimized in bioreactor cultures, it was unfeasible for uniform carbon and nitrogen labelling. Additionally, fermenter cultures have been shown to increase the incidence of mis-folding in the recombinant protein as compared to shaker flask cultures. As such, the plasmid containing the construct truncated at residue 316 previously

42 used by Hino et al. [98] was obtained. This construct was produced in shaker flask cultures to high yields resulting in a crystal structure after purification. Numerous steps were used make this plasmid optimal for expression in Pichia including the N-terminal alpha-factor and Flag-tag sequence, C-terminal 10xHis-tag (Figure 15B), and codon optimization [98]. Therefore, this additional truncation of 18 residues may be advantageous for the purpose of isotopic labelling.

A B

FLAG

10XHIS

Figure 15 – Plasmid and construct used for A2aR expression in P. pastoris. A) Plasmid used for transformation (pPIC9K). Plasmid map generated in SnapGene. B) Truncated A2aR showing attached Flag and 10XHis tag on the N and C termini respectively. Topography map generated by PDBSum.

2.5.2 Transformation into yeast cells

P. pastoris (strain SMD1163) were grown in 25 mL of yeast extract peptone dextrose

(YPD) media (10 g yeast extract, 20 g peptone, 100 mL 20% dextrose for 1 L of media) in a sterile

250 mL baffled flask at 300 rpm, 30 °C in a shaker. In a sterile 2 L baffled flask, 0.5 mL of cells inoculated 250 mL of YPD and shook at 300 rpm, 30 °C for 5-8 hours until the OD600 reached 1.3.

The cells were collected at 1500 ×g, 4 °C for 5 minutes and the pellet was resuspended in 300 mL of ice-cold sterile water. The cells were then centrifuged and resuspended a second time before being spun down and resuspended in 40 mL of sterile ice-cold sorbitol.

The DNA plasmid was linearized with PmeI and purified by the QIAquick nucleotide extraction kit (Qiagen) and desalted by the QIAquick nucleotide removal kit (Qiagen). Ethanol precipitation was used to concentrate the DNA and resuspended to a concentration 1 mg/mL. The linearized DNA (5 μL) was added to 40 μL of cell suspension in an electroporation cuvette. The mixture was incubated for 5 minutes in an ice bath before electroporation (MicroPulser, Bio-Rad) at the Pic setting (2 kV charge voltage, 5.6-5.7 ms pulse duration). Immediately after electroporation, 1 mL of sterile ice-cold 1 M sorbitol was added to the cuvette and mixed.

Electroporation was used on 2 aliquots of cells and 100-200 μL aliquots were spread on RDB

(regeneration dextrose medium, 1 M sorbitol, 2 % dextrose, 1.34 % yeast nitrogen base, 0.00004

% biotin, and 1X amino acids (Glu, Met, Lys, Leu and Ile)) plates and grown at 30 °C until colonies appeared to select for the loss of His auxotrophy. The resulting colonies which grew due to His prototrophy were then resuspended in sterile water and 100-200 μL aliquots were spread on YPD plates with 0.25, 0.5, 0.75, 1, and 2 mg/mL G418 and grown at 30 °C until colonies appeared.

2.5.3 In-colony PCR screening

PCR analysis of Pichia transformants was carried out to get the right colonies with the integrated gene of interest. Colony PCR was applied without isolating DNA from the yeast cells instead of the standard PCR protocol (5 μL of 10×Taq polymerase buffer, 1 μL of 20 mM dNTPs,

20 μM of each primer, 10 pg to 1 μg template DNA, 1 to 2 units of Taq DNA polymerase and sterile water to bring up the total volume to 50 μL) (Sambrook & Russell, 2001). A small yeast colony was taken and suspended in 20 μL of 5% DMSO in a PCR tube, mixed, boiled for 5 minutes

44 and then spun down in a microcentrifuge. The reaction mixture for PCR was set up in the specified order:

1 μL 10 ×Taq polymerase buffer 0.2 μL 10 mM dNTPs 10 pmol each primer 2 μL yeast cells suspension in DMSO solution 0.2 μL Taq DNA polymerase (5 units/ μL) and topped up with sterile H2O to the total volume of 20 μL.

Thermal cycle conditions for PCR amplifications were typically 35 cycles at 94ºC for 30 s, 56ºC for 1 min, and 72ºC for 2 min. The gene of interest was amplified using common primers for Pichia expression vectors, primers 5’Aox1 (5’GAC TGG TTC CAA TTG ACA AGC3’) and 3’Aox1

(5’GCA AAT GGC ATT CTG ACA TCC3’), which are also the sequencing primers for Pichia expression vectors. These primers flank the gene including the alpha-factor. After the amplification, the PCR products were analyzed by agarose gel electrophoresis on a 1 ×TAE buffer,

0.8% gel, run at 120 V, for 40 min.

2.5.4 Screening for high expression mutants

To screen for the best A2aR production in the transformants, eight colonies were selected for small scale production. The colonies inoculated 5 mL buffered-minimal dextrose (BMD) (100 mL 20% glycerol, 100 mL 10X yeast nitrogen base (YNB), 2 mL 0.02% (w/v) biotin for 1 L culture) media and grown in shake-flask cultures at 300 rpm at 30 °C overnight. An additional 20 mL of BMD was added to the cultures which were then shaken at 300 rpm at 30 °C for 24 hours until the OD600 reached 10. The cells were then spun down at 1500 ×g for 10 minutes before the pellets were stored at -20 °C.

A pellet volume of buffer A (7 mM NaH2PO4 at pH 6.5, 7 mM ethylenediaminetetraacetic acid (EDTA), 7 mM dithiothreitol (DTT), and 1 mM protease inhibitor cocktail (Roche) and 15

μg of lyticase to digest the cell wall (from Arthrobacter luteus, Sigma) were added to the pellet

45 and incubated at room temperature for 3 hours. The slurries were spun down at 1500 ×g for 10 minutes and resuspended in a pellet volume of buffer A with a half-pellet volume of ice-cold acid- washed glass beads (Fisher) (420-600 μm diameter). The slurries were disrupted 10 times with 1 minute pulses with vigorous vortexing and then spun down at 700 ×g for 5 minutes at 4 °C and the supernatants were collected.

The broken membranes were run on a 0.5 M Tris/SDS, 12.5% (w/v) acrylamide gel with 2 markers (Precision Plus Kaleidoscope Standards, Precision Plus Unstained Protein Standards, Bio- rad). The proteins were transferred to a Polyvinylidene Fluoride (PVDF) membrane through electrolysis and incubated overnight in 20 mL of Tris-Buffered Saline –Tween (TBS-T) with 0.6 g of bovine serum albumin (BSA) and 0.6 g of gelatine at 4 °C to eliminate non-specific binding.

The membrane was then washed three times with TBS-T for 20 seconds each followed by three washes at 5 minutes each in TBS-T at room temperature. The membrane was incubated at room temperature with the primary antibody (4 μL 6xHis Primary antibody, 20 mL TBS-T, 0.01% (v/v)

NaN3, 1 g BSA) followed by the previously used wash procedure. The diluted second antibody (2

μL Secondary antibody, 2 μL Precision Protein StrepTactin-HRP (Bio-Rad), 20 mL TBS-T) was incubated on the membrane for one hour at room temperature followed by three washes in TBS-T at 20 seconds each followed by three washes in SDS/TBS-T at ten minutes each. The membrane was incubated in 1.5 mL of ECL detection reagents for 5 minutes before film was developed using a Kodak development system.

2.5.5 Large Scale expression of A2aR

The selected colony from small scale screening was used to inoculate 50 mL of BMG (for

15 15 N labelled samples use 0.8% NH4Cl) media in a sterile 250 mL baffled flask. The cells were grown in a shaker at 300 rpm and 30 °C. After 18-24 hours the cells were transferred to a sterile 2

L baffled flask and an additional 200 mL of BMG was added. The cells were shaken at 275 rpm,

29 °C for 18-24 hours before being spun down at 1500 ×g, 4 °C for 10 minutes. The cells were resuspended in 800 mL of buffered-minimal methanol (BMM) (100 mL 1 M NaH2PO4 pH 5.0,

100 mL 10X YNB, 2 mL 0.02% biotin, 7 mL 100% filtered methanol for 1 L culture) media in a sterile 2.5 L Fernbach flask and grown for 24 hours in the shaker at 240 rpm, 21 °C. Cells were initially grown at this stage at 30 °C, however, this affected downstream purification. Lowering the temperature improved purification efficiency probably due to slower cellular processes. The cells were collected through centrifugation at 1500 ×g, 4 °C for 10 minutes and washed twice with

MilliQ water before storing the pellet at -20 °C.

The cell pellet was resuspended in one pellet volume of buffer A in the presence of EDTA- free protease inhibitor (Roche) and incubated with slow shaking with 15 mg of lyticase for 3-4 hours. The cells were then centrifuged at 1500 ×g, 4 °C for 10 minutes prior to being resuspended in a pellet volume of buffer A. A half pellet volume of ice-cold acid-washed beads were added to the suspension and the slurry was vigorously mixed on the vortex in five 1 minute pulses before being spun down at 700 ×g, 4 °C for 5 minutes and the supernatant collected. This process was repeated 8 times until the cells were completely broken. The supernatants were combined and were then centrifuged at 40,000 ×g, 4°C for 30 minutes and the pellets were stored at -20 °C.

2.5.6 Purification of A2aR

The pellets were resuspended in solubilization buffer (62.5 mL per L of culture, 1% (w/v)

DDM, 4 mM antagonist theophylline, 0.2 % cholesteryl hemisuccinate (CHS, Sigma), 20 mM

KH2PO4, pH 7.5, 20% (v/v) glycerol) and stirred overnight at 4 °C. The solution was centrifuged at 150,000 ×g, 4 °C for 30 minutes and the supernatants were collected. The batch method with 6-

His tag affinity resin (Ni2+-NTA agarose, Qiagen) was used to extract the protein from the

47 supernatant. We used approximately 5 mL of resin slurry per 800 mL culture and stirred the solution slowly at 4 °C overnight to allow binding. The resin was collected in an empty PD-10 column (GE Healthcare) while the other solubilised proteins flowed through.

The resin was washed in buffer (0.05% (w/v) DDM, 4 mM theophylline, 0.2 % CHS, 50 mM KH2PO4, pH 7.5, 400 mM NaCl, 20 mM imidazole, 5 mM PMSF, 10% (v/v) glycerol) about two times the resin volume. This was mixed for 2 minutes before flowing through the column. The protein was eluted with elution buffer (0.05% DDM, 4 mM Theophylline, 0.2 % CHS, 50 mM

KH2PO4, pH 7.5, 400 mM NaCl, 10% (v/v) glycerol) three times using 500 mM imidazole after two minutes of mixing each. The protein concentration was monitored using Bradford protein assay (Bio-Rad DC protein assay) with PR as a protein standard. The elution fractions were combined and concentrated to a volume of 1-2 mL. The eluate and crude extract were analysed through SDS-PAGE to determine protein purity. The gel was incubated in 50 % methanol, 10% acetic acid for one hour, shaking at 100 rpm. After washing with 200 mL of MilliQ water 2 X for

10 minutes each, it was stained with InVision His-tag stain for one hour, shaking at 100 rpm. The gel was visualized using UVP ChemiDoc-It Imager using ultraviolet light.

2.5.7 Reconstitution of A2aR for SSNMR studies

The concentration as determined by Bradford assay for the protein eluates was used to determine the proper reconstitution volume. The proteins and lipids were combined for a protein to lipid to Triton X-100 ratio of 1 :1 :0.5 (w/w/w) and incubated at room temperature for 1 hour.

To withdraw the detergent, Bio-beads SM-2 (Bio-rad) were added at 0.6 g per mL of mixture and incubated at 4 °C with mixing for 24 hours. The proteoliposomes were collected using a syringe and the beads were washed in 200 mM NaCl until the turbidity was negligible. The collected proteoliposome suspension was centrifuged at 150,000 ×g, 4 °C for 50 minutes. The pelleted

48 proteoliposomes were washed several times in NMR buffer (10 mM NaCl, 25 mM TrisCl, pH 8) and stored at 4 °C.

2.5.8 Preparation of samples for SSNMR

The washed isotopically-labelled proteoliposomes were centrifuged at 900,000 ×g for 3 hours with 20 μL of NMR buffer. The samples were then center-packed into a 3.2 mm NMR rotor.

Experiments were run on Bruker Avance III spectrometer at frequency of 800.230 MHz.

2.6 Results and Discussion

2.6.1 In-colony screening

To ensure the presence of the target gene in our transformants, colonies which grew on

YPD plates supplemented with 0.25 mg/mL G418 were chosen for in-colony PCR screening. Cells were treated with 5% DMSO, boiled for five minutes and membranes spun down before the supernatants were used for amplification. The gene encoding Trunc-A2aR is 978 bp. The primers are located adjacent to the inserted gene within the AOX1 sequence; consequently the amplified product will contain parts of the plasmid. Including the vector, this corresponding amplified product should be 1.6 kDa. As SMD1163 is a Mut+ strain of Pichia, the AOX1 gene is intact and is approximately 2.2 kDa. Therefore there should be two bands present in the successful integrants:

1.6 kDa and 2.2 kDa. The gene encoding FL-A2aR, including the Flag-tag, His-tag, and vector fragment is 1.9 kDa. As such, successful FL-A2aR transformants should have two bands of size

1.9 kDa and 2.2 kDa. The band weights in colonies 5 and 6 of Trunc-A2aR (1.2 kDa and 2.1 kDa) and colony 9 of FL-A2aR (1.6 kDa and 2.1 kDa) mostly agreed with these predicted values indicating the successful transformations (Figure 16). Colonies 5 and 6 of Trunc-A2aR were re- plated on YPD and screened for expression.

(bp) 1 2 3 4 5 6 7 8 9 10

10,000 8,000 6,000 5,000 4,000 3,000 2,500 2,000 1,500 1,000 750 500

250

Figure 16 – In-colony PCR of A2aR. 1) Marker. 2-6) Trunc-A2aR colonies. 7-10) FL-A2aR colonies.

2.6.2 Expression of A2aR

Expression of A2aR in the truncated form was monitored through Western blot analysis.

A2aR-Trunc has a molecular weight of 36.5 kDa. The FL-A2aR construct encodes a protein of size 47 kDa. Membrane proteins typically run at lower apparent molecular weights [55]. High intensity bands of approximate molecular weight 25, 40, and 80 kDa were seen in the membrane fraction, thought to correspond to the monomer, dimer, and tetramer of Trunc-A2aR (Figure 17).

This oligomerization pattern was previously observed in Pichia expressed A2aR (Figure 9D).

Colony C was chosen for large scale purification.

A B C D E F G H

80 kDa

40 kDa

25 kDa

Figure 17 – Immunoblot for Trunc-A2aR expression screening of 8 colonies of P. pastoris. Colonies grown in 25 mL shaker cultures before being broken, and their membranes were treated with 10 % SDS prior to centrifugation. The supernatant was run by gel electrophoresis and detected using Qiagen/Biorad antibodies.

2.6.3 Purification of A2aR

Trunc-A2aR was purified through metal affinity chromatography and progress was monitored through InVision His-tag Stain of SDS-PAGE gel. The presence of an approximately

70 kDa band, assumed to be A2aR in an oligomeric state, was monitored (Figure 18). After binding overnight to Ni2+-NTA-agarose resin and subsequent washing steps, losses of approximately 10 % were estimated based on band intensity (Figure 18). Bands from the solubilised fraction and the final eluate of apparent molecular weight 70 kDa were excised and analysed through mass spectrometry for identification.

A B C D E F G

75 kDa

25 kDa

Figure 18 – SDS-PAGE analysis of purification of Trunc-A2aR by nickel affinity resin. A) WesternC protein marker (Bio-Rad). B) Eluate. C) Solubilized membrane fraction. D) Flow-through after resin binding. E-G) 15 mM imidazole washes of protein-bound resin.

Purification via the N-terminal Flag tag was attempted for both Trunc-A2aR and FL-A2aR.

Purification progress was monitored via InVision His-tag staining of SDS-PAGE gels. An approximately 80 kDa band was purified after Flag-tag purification and subsequent elution (Figure

19). Band intensity was greater in the eluted samples from Trunc-A2aR indicating increased stability (Figure 19). Additionally, the expected size difference between the Trunc-A2aR and FL-

A2aR (10 kDa) was not visible on the gel (Figure 19). This implies either contamination of a histidine-rich protein of approximately 80 kDa in both cases or degradation of the C-terminal tail of FL-A2aR.

A B C D E F G H I

80 kDa

30 kDa

Figure 19 – SDS-PAGE of Flag-tag purification of FL-A2aR and Trunc-A2aR. A) His-tag marker. B) Solubilized Trunc-A2aR membrane fraction. C) Flow through after Flag-affinity resin binding for Trunc- A2aR. D) Trunc-A2aR eluate after Flag-tag purification. E) His-tag affinity purification eluate of Trunc-A2aR. F) Solubilized FL-A2aR membrane fraction. G) Flow through after Flag-affinity resin binding for FL-A2aR. H) FL-A2aR eluate after Flag-tag purification. I) His-tag affinity purification eluate of FL-A2aR.

2.6.4 FTIR studies of reconstituted A2aR

The extent of labelling of Trunc-A2aR in 15N media was measured through FTIR. The sample was reconstituted into DOPC/PS/Chol (9:1:1) liposomes at a protein to lipid ratio of 2:1

(w/v). A shift in the amide II peak to the right was seen of 12 cm-1 (Figure 20). Previously, shift of 17 cm-1 was seen in the amide II band as a result of 15N-labelling in LR [30]. This shift is similar which shows high labelling efficiency of the sample.

1.80

1.60

1.40

12 cm-1 1.20

1.00

0.80

0.60

0.40

0.20

0.00 1800 1700 1600 1500 1400

Figure 20 – Baseline-corrected static FTIR spectra of A2aR reconstituted into a DOPC/PS/Cholesterol (9:1:1) liposome measured for 15N-Trunc-A2aR (blue) and natural abundance Trunc-A2aR (red). Absorbance measured on Bruker IFS66vs machine with a temperature-controlled sample holder (Harrick) connected to a circulating water bath (Fisher).

By comparing spectra of our produced samples with those of previously provided Trunc-

A2aR, one can use FTIR to identify the extent of contamination and/or differences in protein conformation. In comparison with the spectra for 334-A2aR, the sample of Trunc-A2aR produced downshifted amide I and amide II bands (Figure 21). Additional broadening of both peaks can be seen with a distinctive shoulder in the amide I peak indicative of beta or random coil content. This shift is characteristic of the increased beta content being seen in the produced sample as compared to the sample provided by Imperial College, consistent with contamination by the proteins identified by mass spectrometry to be discussed later.

Figure 21 – Baseline-corrected static FTIR spectra of A2aR reconstituted into a DOPC/PS/Cholesterol (9:1:1) liposome measured for Trunc-A2aR (red) and natural abundance 334-A2aR (blue). Absorbance measured on Bruker IFS66vs machine with a temperature-controlled sample holder (Harrick) connected to a circulating water bath (Fisher).

2.6.5 SSNMR of Trunc-A2aR

A 1D 15N-SSNMR spectrum was taken for 15N-Trunc-A2aR. In comparing this spectrum with those previously obtained for 334-A2aR, an upshift of the main amide backbone peak of 5 ppm was observed (Figure 22). Generally, residues which form alpha-helical structures reside between 100 to 120 ppm while random coils and beta-strands are between 110 and 150 ppm [119].

It is residues of beta and random coil structures, based on the general chemical shift of the amide backbone, which contribute to the shift in the Trunc-A2aR spectra (Figure 22). Broadening is seen in the Arg peaks, with NE being upshifted as well arguing for a different chemical environment

(Figure 22). Also, Trunc-A2aR has lower peak resolution showing poor homogeneity in the sample

(Figure 22). It is obvious that the sample either did not contain A2aR or contamination was of such magnitude that no reliable conclusions can be made about Trunc-A2aR. Alternatively, this could potentially indicate denaturation of the protein.

Protein Backbone

129 ppm 124 ppm

Arg Arg

NE NH

89.5 ppm 92 ppm 78 ppm

Figure 22 - 1D SSNMR spectra of 15N-Trunc-A2aR (red) as compared to 15N-334-A2aR (black).

2.6.6 Mass spectrometry of gel extracts

Given the apparent secondary structure alteration within the sample, identification of the protein was undertaken. In-gel digestion and subsequent mass spectrometry was used to identify the isolated 70 kDa protein band (before the affinity purification). The protein was identified as alcohol oxidase I, a protein native to P. pastoris (Figure 23). The gene controlling the expression of this protein is the methylotrophic promoter also used for heterologous expression in P. pastoris.

As such, the high yield of AOX1 relative to the protein of interest is quite probable assuming low

A2aR production. Protein production in yeast is proportional to the number of copies of the linearized plasmid which is taken up and integrated by the cells. Thereby cells of lower plasmid copy number will conversely lead to low protein yields. Additionally, the use of minimal media, which is necessary for isotopic labelling, is prohibitive in terms of the addition of DMSO and external histidine which have been shown to improve A2aR production in the past [7]. DMSO was attempted to be added to the minimal media but resulted in high cell death in BMM.

Figure 23 – Mass spectrometry results based on in-gel digestion of putative 70 kDa protein (before purification). Hits are underlined in blue.

AOX1 contains key structural properties which make it distinguishable from A2aR spectroscopically (Figure 24A). Notably, the Rossman folds which are beta-sheet domains. AOX1 consists of 3.9 % histidines a majority of which can be found at the protein surface, which are the amino acids targeted by metal affinity purification (Figure 24B). In the event of low A2aR production, AOX1 most likely binds the nickel resin with great affinity and as such is purified in conjunction with A2aR. Estimated secondary structure content based on the homology model is

32 % α-helical and 22 % β-strands (Figure 24A). This mixture of secondary structure is consistent with the shifts seen in both FTIR and SSNMR relative to the 334-A2aR sample.

A B

Figure 24 – Structural model of AOX1. A) Secondary structure is coloured with β content in red, α helices in cyan and loop in magenta. B) Surface histidines identified in red. Homology model produced by Swiss-Model based on the crystal structure of Aspergillus flavus FAD glucose dehydrogenase (PDB ID: 4YNT) and rendered in PyMol.

In the metal affinity purified sample, the mass spectrometry of the gel extract of the 70 kDa band identified NUAM, a subunit of the mitochondrial NADH:ubiquinone oxidoreductase

(Complex I) (Figure 25). There are 14 core subunits in complex I, which act as the entry for the electron transport chain in mitochondria [120]. The assembly forms an L-shaped structure with a hydrophilic arm which extends into the mitochondrial matrix. NUAM is homologous to a subunit found in Yarrowia lipolytica which has been characterized previously [120]. This protein is composed of hydrophilic residues as it is found in the hydrophilic arm of complex 1. Similar to

AOX1, there are many surface histidines in NUAM (Figure 26B), which may explain its presence despite the imidazole washing procedure. Estimated secondary structure content based on the homology model is 25 % α-helical and 15 % β-strands (Figure 26A). This protein structure, similar to AOX1, agrees with the secondary structure content implied by the shift in the SSNMR spectra

58 and FTIR of Trunc-A2aR. Additionally, the low lipid retention seen by FTIR in the sample is most likely due to precipitation of the protein in lieu of the formation of proteoliposomes.

Figure 25 – Mass spectrometry identification of putative 70 kDa protein isolated from SDS-PAGE of cell- extracts after purification. Sequence of NUAM with identified peptides underlines in blue.

A B

Figure 26 – Homology model of NUAM. A) Secondary structure is coloured with β content in red, α helices in cyan and loop in magenta. B) Surface histidines identified in red. Homology model produced by Swiss-Model based on the crystal structure of chain 3 of the hydrophilic arm of respiratory complex I from Thermus thermophilus (PDB ID: 2FUG) and rendered in PyMol.

These contaminants in the sample must be addressed in further attempts to produce A2aR samples. The ideal solution requires re-transformation of the cells to introduce higher copy numbers of the plasmid. The higher expression would then allow purification to proceed with

A2aR hopefully being purified instead of AOX1 and NUAM as the metal cation affinity resin

59 would be more likely to form strong associations with the 10xHis-tag on the protein. Also, this would prevent contamination in FLAG-tag binding as the protein will no longer be unavailable for binding. Furthermore, finding methods to prevent proteolysis of the C-terminal tail would allow studies into the full-length construct which could be more relevant medically than the truncated version [121].

Chapter Three Microbial Rhodopsins

3.1 Rhodopsins

Rhodopsins are a distinct class of seven transmembrane α-helical proteins that perform photosensory and ion-transport functions [122]. Analogous to the GPCR visual rhodopsin present in the human eye, these proteins bind retinal, a vitamin A derivative, as a chromophore which is essential to their function [122]. Several classical studies of membrane proteins have used microbial rhodopsins to develop new techniques for structural investigation [24]. The most comprehensively studied rhodopsin is bacteriorhodopsin (BR), a light-activated haloarchaeal proton pump [123].

After the discovery of four archaeal-type rhodopsins in the membrane of Halobacterium salinarum, a large family of microbial rhodopsins was found in diverse species, including archaea, eubacteria, cyanobacteria, fungi and algae [122, 124, 125]. Rhodopsin function is highly variable and organism-dependent. For microbial (type I) rhodopsins their functions include photo-sensors, ion channels and pumps. Type II rhodopsins are found in animals and perform functions related to vision and other light-dependent processes (e.g., circadian). Microbial rhodopsins can be divided into two groups according to their function: ion transporters and photosignal transducers. In general, the major differences between these two groups are the photocycle rates, which are usually slow for sensors, and the amino acid residues found at the proton donor and acceptor positions

[126].

While archaeal rhodopsins are known since 1972, it is only within the last fifteen years that rhodopsins were discovered in eubacteria as well [127]. While similar in structure to their archaeal homologs, there are some eubacterial rhodopsins with unique architecture. Anabaena sensory

61 rhodopsin (ASR) was the first photosensor identified in eubacteria in 2003 [126]. Several structural studies have occurred to date on the truncated form of this protein, with the structure being solved with SSNMR [26]. Another unique rhodopsin is proteorhodopsin (PR) which acts as a proton pump similar to BR. Despite the similarity in function, PR is not as well characterized as BR [24]. These two eubacterial rhodopsins are described in greater detail in the next chapters.

3.2 Bacteriorhodopsin

Bacteriorhodopsin is a well characterized microbial rhodopsin due to its stability and ability to form trimers in membranes [128]. Its structure was determined using electron crystallography in

1990 and has been a template structure for other rhodopsins and GPCRs since [122]. It is found in

Archaea, more notably Halobacteria. It is produced in response to oxygen starvation. Its primary function is to act as a proton pump which under illumination forms a proton gradient which is then converted to chemical energy by ATP synthase. The proton is passed to extracellular side of the membrane through a series of proton hops (Figure 27).

Upon excitation via light, the bound retinal isomerizes from an all-trans conformation to 13- cis-retinal which lowers the effective pKa of the retinal Schiff base (Figure 27). This results in a proton transfer from the Schiff base (Lys 216) to the primary proton acceptor (Asp 85). This process allows a proton from the primary proton donor Asp 96 to be transferred to the Schiff base

(Figure 27). The primary proton acceptor is eventually deprotonated leading to a proton being released into the external medium at acidic pH (at alkaline pH the proton release from proton- releasing complex of glutamates precedes the deprotonation). The primary proton donor takes up another proton from the cytosol while retinal undergoes isomerisation back to the all-trans state

(Figure 27).

Figure 27 – Proton transfer chain in BR. Pathway of the proton transfer towards the extracellular surface (located at the bottom) is outlined [129].

3.3 The Photochemical Cycle

The retinal molecule bound in microbial rhodopsins reacts to light isomerising from an all- trans configuration to 13-cis [122]. However, there are a number of reaction intermediates through which the retinal binding pocket (Figure 28) and the rest of the protein changes; each has an associated absorption wavelength: K610, L550, M410, N560, and O640 [130]. As such, one can isolate their presence with techniques such as laser spectroscopy. Additionally, the length of the photocycle is dependent on whether the rhodopsin is a sensor or a proton pump. Proton pumps tend to have faster photocycles, due to demand for efficient transport. Sensory rhodopsins can have associated transducer molecules which require longer times to remain dissociated from the signalling complex [131].

Figure 28 – Photocycle of BR showing the retinal conformation in the photointermediates coloured according to their corresponding wavelengths [132]

3.4 Expression of rhodopsins in E. coli

In this study, transformed E. coli was used to express two proteins: proteorhodopsin (PR) and Anabaena sensory rhodopsin (ASR). The nutritional diversity of E. coli is valuable for the expression and downstream analysis of the desired protein [133]. This exemplifies the desired characteristics of a good host due to its ability of to grow in both enriched and minimal media with a relatively short generation time [133]. This nutritional flexibility is critical for isotopic labelling of the protein being over-expressed for downstream analysis [44]. As previously discussed, the ability to control the sole sources of carbon and nitrogen in the growth media is critical for isotopic labelling. As such, one has the ability to introduce differential labelling schemes within the produced protein [44]. Additionally, the protein yield per liter of a culture should be sufficient to perform SSNMR at acceptable signal-to-noise ratios. Optimized protocols can yield as much as 15 milligrams of PR [24] or 5 milligrams of ASR [55] per litre of shake-flask culture in E. coli following the protocol outlined in Figure 29.

Grow E. coli in minimal media and induce rhodopsin Solubilize protein in over-expression detergent and purify using IPTG and on Ni2+-NTA retinal agarose resin

Reconstitute protein Study sample with into DMPC/DMPA FTIR or SSNMR (9:1) liposomes at and assign peaks protein to lipid ratio of 2:1

Figure 29 – Sample preparation of microbial rhodopsin PR in E. coli based expression system for SSNMR

E. coli are a type of gram negative bacteria [134]. Gram negative bacteria have two cell membranes: the inner membrane (IM) and the outer membrane (OM) (Figure 30). The OM consists of mainly of β-barrel proteins with the outermost membrane layer consisting of lipopolysaccharide

(LPS) (Figure 30). Proteins more α-helical in structure are generally found in the IM [135]. As such, PR and ASR would be typically found in the inner membrane when expressed in E. coli [18].

This poses the difficulty of isolating the inner membrane when studying membrane proteins in vivo as the OM and IM are both linked to the peptidoglycan layer in the periplasmic space (Figure

30). As such, purification procedures involving membrane solubilization are generally used to isolate rhodopsins for structural analysis. However this does not preclude the possibility of interactions between the inner and outer membrane and their components and possibly co- purification with the protein of interest.

Figure 30 – Model of the E. coli membrane envelope [135].

Chapter Four Full-Length Anabaena Sensory Rhodopsin

4.1 Anabaena Sensory Rhodopsin

Anabaena sensory rhodopsin (ASR) is a membrane-bound light sensor protein which was discovered in cyanobacteria in 2003 [52]. Despite numerous studies using both biochemical and biophysical assays, it is only in the past year that the photosensory transduction mechanism of

ASR was clarified. There is an associated soluble transducer protein (ASRT) which, upon light- induced release from ASR [54], directly interacts with the promoters of genes cpc and pec, which correspond to the synthesis of phycocyanin and phycoerythrocyanin (Figure 31A). These proteins are associated with the phycobilisome (Figure 31B), or the light harvesting complex of cyanobacterial chlorophyll photosynthesis [136].

A B

Figure 31 –A) Representation of the ASR signal transducing mechanism showing direct interaction of the transducer with DNA [136]. B) Structure of the phycobilisome showing the light harvesting process [137].

4.2 ASR Structure and Photochemistry

The structure of ASR has been determined using both SSNMR [26] and x-ray crystallography [138]. It shares the seven transmembrane alpha-helical architecture as seen in all members of the rhodopsin family (Figure 32). There is a bound retinal molecule in the transmembrane domain which is covalently attached to Lys210. ASR consists of 261 amino acid residues (26 kDa) [52]. When produced in E. coli, ASR exhibits a red colour which is indicative of retinal binding. ASR is photochromic. The predominantly all-trans-retinal (ASRAT)in its dark- adapted state transitions to a mixture dominated by 13-cis species upon illumination (Figure 33).

The light-adapted state results in a transition from predominantly 13-cis retinal (AST13C) to all- trans in the dark (Figure 33). As such, the photostationary ASRAT/ASR13C ratio depends on the wavelength of illumination. Therefore, ASR is capable of sensing light of a single pigment [126].

This is opposite to what is known in BR but similar to the photochromism seen in invertebrate visual rhodopsins [126]. The photocycle of all-trans-ASR is similar to that of BR in the early stages, but all the steps related to proton translocation to and from the retinal Schiff base are different, because of the absence of a second carboxylate group at the ASR photoactive site and lack of the cytoplasmic proton donor [139-141]. This deprotonation of the Schiff base and ensuing conformational change of ASR provide the trigger for the dissociation of ASRT from ASR, which then diffuses through the cytoplasm to the DNA [136].

Figure 32 – Structure of C-terminally truncated ASR as determined by SSNMR spectroscopy. A) Overall structure with bound retinal in yellow. B) Residues associated with retinal binding in ASR. C) Important polar cytoplasmic cluster residues and their locations relative to helices in ASR. [26]

Figure 33 – Photochromism of ASR [126].

ASR has a relatively slow photocycle, and its Schiff base proton acceptor (Asp75) is conserved compared to other rhodopsins, but it lacks the proton donor present in proton-pumping rhodopsins. Instead, it carries a serine residue (Ser86). One unique feature of ASR is the presence of a proline (Pro206) instead of an aspartate in one helix turn below the Schiff base lysine residue

[52]. Further studies using SSNMR have found ASR to form tightly associated trimers in membranes with inter-monomer contacts on helices B, D and E (Figure 34). This was determined via combination of PREs and biochemical methods [55]. This is similar to the oligomer configuration of BR, suggesting formation of 2D crystals of trimers in membranes [128]. Indeed, crystallinity of ASR was proven recently [18] and this 2D crystal structure is extremely useful for

SSNMR studies since it ensures homogeneity of the sample.

Figure 34 – Structure of the ASR trimer as determined by SSNMR with intermonomer contacts shown on the right [26]

4.3 Full-Length ASR

Early studies of the first discovered type 1 proteins, the haloarchaeal proton pump bacteriorhodopsin and phototaxis receptors sensory rhodopsins I and II (SRI and SRII), showed that they were functional after the truncation of their flexible C-terminal tail. This is advantageous as truncation of the C-terminus greatly increased heterologous expression levels [142]. However, in the case of ASR, there was a 20% reduction in photocycle time as compared to the full-length construct [143]. Interestingly, this difference has not been observed for purified full-length and truncated ASR [140, 143].

Signal transduction by ASR is unidirectional. This means that its photo-signalling mechanism only works in the periplasm to cytoplasm direction. This is in line with the direction of light-induced Schiff base proton transfer towards the cytoplasmic surface (Asp217), which causes further conformational changes of ASR and activation of ASRT [141]. Surprisingly, previous photoelectric studies have found that C-terminal truncation of ASR dramatically affects the vectoriality of the proton transfer [143]. It appears that in whole E. coli cells the truncation inverts the direction of proton movement associated with the Schiff base deprotonation suggesting that the photoactive site and the C-terminal tail are in communication[143].

This communication may be due to interaction between ASR and some cytoplasmic components, which in the case of ASRT has been shown to involve the C-terminal end of ASR

[136, 143]. Indeed, the direct effect of ASR C-terminus, which contains 7 positively charged arginines, on gene expression has been shown, suggesting its interaction with DNA [144] (or some transcription factors). The ASR structures which have been determined thus far have used a truncated form [26, 138]. But, in order to understand activation of the transducer and achieve a

71 more complete picture of ASR dynamics, it is imperative to investigate the full-length form of

ASR (FL-ASR). This necessitates the production of FL-ASR for SSNMR.

Figure 35 – Structure of FL-ASR based on the SSNMR derived structure of C-terminally truncated ASR (PDB ID: 2M3G). Retinal is in yellow with important residues in green. Model produced in Pymol with a cytosolic helix manually attached for visualization purposes.

4.4 Materials and Methods

4.4.1 Expression of FL-ASR

BL21 cells, transformed with a plasmid encoding N-terminally 6xHis-tagged FL-ASR under an IPTG-inducible promotor, were cultured in M9 minimal medium with D-glucose and ammonium chloride as the sole carbon and nitrogen sources respectively. A 2 mL culture was induced with 20 μL of cells stored at -20 °C and grown for 24 hours at 30 °C and 240 rpm. The 2

72 mL culture was used to inoculate a 25 mL culture which was grown overnight at 30 °C and 240 rpm. The 25 mL culture was subsequently used to inoculate 1 L of M9 media and grown at 30 °C and 275 rpm. The cells were induced at an optical density of 0.4 with 7.5 μM all-trans-retinal and

1 mM Isopropyl-β-D-thiogalactoside (IPTG). The cells were collected 6 hours after induction at

5000 × g and 4 °C.

4.4.2 Purification of FL-ASR

The cells were resuspended in 60 mL of lysis buffer (150 mM NaCl, 50 mM Tris-HCl, 1 mM MgCl2, 2 μg/mL DNAse I, 0.2 mg/mL lysozyme, pH 7.2) and shaken at room temperature for

3 hours at 500 rpm. The cells were freeze fractured by storing at -20 °C overnight and thawing.

The cells were broken by sonication and centrifuged at 5000 × g to check for unbroken membranes and to remove debris. The supernatant was subsequently centrifuged at 150,000 × g for 50 min at

4 °C and the resultant supernatant discarded. The membrane fraction was solubilized in 30 mL of solubilization buffer (5 mM Tris-HCl, pH 7.5, 1% (w/v) n-Dodecyl β-D-maltoside (DDM)) at 4

°C for 4 hours. The sample was then spun down at 40,000 × g for 1 hour to remove the insoluble fraction. The protein was purified via the batch procedure using Ni2+-NTA agarose resin (Qiagen) allowing the resin to bind the protein overnight at 4 °C. The protein-bound resin was washed 8 times in washing buffer (300 mM NaCl, 50 mM Tris-HCl, 40 mM Imidazole, pH 8, 0.05% (w/v)

DDM) on a 500 mL Nalgene vacuum filter. The protein was then eluted through a step-wise addition of 25 mL of elution buffer (300 mM NaCl, 50 mM Tris-HCl, 500 mM imidazole, pH 8,

0.05% (w/v) DDM) and the resin removed through centrifugation at 5000 × g for 10 min at 4 °C.

The eluted fraction was syringe-filtered to remove residual resin. The protein concentration was determined through the retinal absorption peak at 530 nm with an extinction coefficient of 48,000 cm-1. The protein yield for FL-ASR was approximately 1 mg for 1 L culture.

4.4.3 Analysis of contaminating proteins

The samples were denatured in 5% SDS and boiled until the colour disappeared. 5xSDS- loading dye was added prior to running the samples on a 12.5% acrylamide/bis-acrylamide, 0.1%

SDS gel at pH 8.8 for 1.5 hours at 120 V. The gel was incubated in 50 % methanol, 10% acetic acid for one hour, shaking at 100 rpm. After washing with 200 mL of MilliQ water 2 X for 10 minutes each, it was stained with InVision His-tag stain for one hour, shaking at 100 rpm. The gel was visualized using UVP ChemiDoc-It Imager using ultraviolent light. The gel was then incubated with Coomassie blue stain overnight. The gel was washed in 20 % Methanol and visualized. The bands identified as containing contaminating proteins were excised and sent to mass spectrometry for identification.

4.5 Results and Discussion

4.5.1 Temporal dependence of expression

For C-terminally truncated ASR, after induction by IPTG/all-trans-retinal, cells can be grown for 24 hours, resulting in good ASR yields of up to 6 mg/L of culture after purification.

However, in the case of FL-ASR, a brown colour was observed to form indicative of cell death and lacking the purple colour associated with ASR expression (Figure 36). This could be interpreted as toxicity of FL-ASR to BL21 cells and imply interaction of the C-terminus with some cellular components. Time trials were conducted, resulting in an optimal 6 hour induction time, after which the brown colour develops.

Post-induction Culture Number incubation time 1 2 3 4 5

3 hrs

6 hrs

9 hrs

18 hrs

Figure 36 – FL-ASR production monitored through pink colour formation showing dependence on post- induction incubation time length.

4.5.2 Identification of contaminating proteins

During purification of ASR harvested at the optimal time point of induction, significant losses were seen from the resin bound FL-ASR after incubation with 35 mM imidazole. Under the same conditions, the truncated ASR does not exhibit any significant losses at this stage. This suggests that there is some form of interference occurring with His-tag binding on the N-terminus.

To identify the interacting partners, the purified protein was run on a 12.5 % acrylamide/bis- acrylamide gel, resulting in multiple bands (Figure 37). Additionally, the purified protein was re- incubated with the cytoplasm to find possible cytoplasmic interaction partners in E. coli and run on the same gel. Membrane proteins run at a lower molecular weight on gels than their actual mass, as such the monomer of 29 kDa is observed at 20 kDa [145] (Figure 37). Not all of these bands were attributable to FL-ASR or its oligomers (previously seen in CT-ASR at 20, 37 and 50 kDa

[55]) as seen through the comparison of His-tag stained gel versus the Coomassie blue.

Specifically, the band at approximately 30 kDa in lanes 4-6 loses intensity after re-incubation with the cytosol and subsequent washing when monitored through His-tag staining. In contrast, the

Coomassie stained gel, shows very little change in intensity in lanes 4-6, indicative of a contaminant at this location (Figure 37).

A B

(kDa) 1 2 3 4 5 6 1 2 3 4 5 6 200 150 100 75

37 ** ** 25

20 * *

Figure 37 – SDS-PAGE of FL-ASR during the purification process stained with (A) Coomassie Blue and (B) with InVision His-tag stain. 1) Flow through after batch binding. 2-3) 35 mM Imidazole washes. 4) Flow through after cytoplasm incubation. 5) 35 mM Imidazole wash after cytoplasm binding. 6) Final eluate after cytoplasm incubation. FL-ASR monomer indicated with *. OmpF band denoted with **.

The major contaminant band of approximately 30 kDa was excised and mass spectrometry identification of the protein was performed through digest and peptide mass identification (Figure

38). It was identified as OmpF, a 39.3 kDa outer membrane protein abundantly found in E. coli.

OmpF is a bacterial porin which allows diffusion of small molecules through the outer membrane

[146]. This protein forms a 1.2 nm pore which allows very fast rates of diffusion as compared to other bacterial porins [146]. The protein forms a beta-barrel with an alpha- helix being formed in the pore (Figure 39). Porin formation is part of the E. coli response to changing environments and its overexpression is often observed upon cell stress, such as growth on minimal medium and expression of toxic proteins [147, 148]. Porins, like OmpF, are produced under different

76 conditions. Variations in pH, osmolarity, temperature, the concentration of certain toxins, and growth phase have been proven to affect its production [146]. It seems that the conditions induced by M9 media and ASR overexpression promote porin formation.

Figure 38 – Mass spectrometry analysis of 30 kDa band isolated from SDS-PAGE as compared to the sequence of OmpF. Identified fragments are underlined in blue.

Figure 39 – Structure of OmpF with beta-barrel architecture in green and inner helix identified in red [149].

OmpF is typically found in trimers within Gram negative bacteria [146]. Oligomerization of OmpF has been shown to participate in the function Colicin N, a toxin which affects Gram negative bacteria. Colicin N can use OmpF trimers as part of its translocation mechanism into the

E. coli outer membrane. The pore-forming domain of Colicin N binds the cleft between OmpF

77 monomers, filling the pore (Figure 40). The remaining helices are sufficient to span the periplasm and form a functional toxic pore, thereby killing the bacterium [150]. It is this cleft formed by

OmpF oligomers which could participate in binding with the C-terminal tail of FL-ASR (Figure

41). This would form a relatively stable complex, which would co-purify. It should be noted that this interaction could also occur after solubilisation, wherein the detergent-bound membrane proteins can interact within solution. Additionally, the poor yields seen by FL-ASR, when expressed in E. coli, could also be linked to the interruption of cellular functions by blocking transport through OmpF. To interact with OmpF by its C-terminus, FL-ASR would have to flip its native orientation in the inner membrane. This could explain the inversion of direction of the photoelectric signals between the full-length and C-terminally truncated ASR observed in the whole E. coli cells previously and explain the lack of any difference in photochemistry of the purified proteins at the same time. However, the cell death could also be a result of the cationic tail of FL-ASR directly interacting with the DNA, disrupting cellular processes. More investigation into the orientation of FL-ASR in the E. coli membrane is necessary to make any further conclusions.

Figure 40 – Colicin N translocation mechanism into membranes containing OmpF oligomers [150].

Figure 41 – Suspected OmpF binding mechanism as a result of FL-ASR inversion in E. coli inner membrane.

Chapter Five Biosynthetic production of isotopically labelled retinal

5.1 Retinal Biosynthesis Pathway

Retinal is derived from β-carotene in a reaction catalysed by a dioxygenase. β-carotene is an important vitamin in mammalian diets as retinal is a key component of sight [151]. As previously discussed, retinal is also the chromophore used in microbial rhodopsins. The ability to produce this ligand isotopically labeled within the expression host could be critical for structure determination of microbial and animal rhodopsins. Until now, these structural studies used either natural abundance retinals, or retinals 13C-labeled at only one or two positions [26, 151-153], which gave a very limited amount of structural information. In vitro synthesis of multiply or fully labeled retinals is prohibitively expensive [154], so that biotechnological approaches look very attractive as an alternative.

The biosynthetic pathway for beta-carotene is well studied and has been introduced as an exogenous pathway into many organisms, most famously golden rice [155, 156]. Two independent non-homologous metabolic pathways are known: the mevalonate (MVA) pathway in eukaryotes and archaea and the methylerythritol phosphate (MEP) pathway in bacteria and several photosynthetic eukaryotes [155]. The pathways are outlined in figure 42. The MVA pathway uses the pyruvate precursor from glycolysis and undergoes the citric acid cycle before mevalonate is produced ultimately resulting in IPP, the five-carbon building block of carotenoids (Figure 42B).

The MVA pathway is also found in a few bacteria, what has been explained in previous works by recent acquisition by horizontal gene transfer (HGT) from eukaryotic or archaeal donors. Previous studies have found that E. coli uses only the MEP pathway [157], which is ideal when trying to introduce differential labelling in retinal.

Figure 42 – A) Biosynthesis of retinal derivatives from beta-carotene. B) Synthesis of beta-carotene and retinal via the two metabolic pathways (MVA and MEP) as derived from glucose or glycerol sources [155].

5.2 Plasmids

The strain of cells being used for this study is E. coli UT5600 which were transformed with two plasmids: pOrange and pKJ900. This strain has been used previously to characterize color- tuning mutations of proteorhodopsin using selection of colonies with different color after random mutagenesis [158]. pOrange contains the genes encoding CrtE, B, I and, Y responsible for synthesis of β-carotene from FPP (Figure 43A-B) under a constitutively active promoter. Thus, the cells are constantly producing β-carotene, resulting in an orange colour. This plasmid also contains the gene encoding chloramphenicol resistance (Figure 43B). A second plasmid, pKJ900,

81 contains the genes for PR and Mus musculus 15’-β-dioxygenase (Figure 43C). PR is regulated by the IPTG-inducible lac operon promoter. β-dioxygenase is regulated by L-arabinose induction.

Upon the addition of IPTG and L-arabinose, the cells will synthesize PR with internally produced retinal, allowing for isotopic labeling of both the opsin and the chromophore. This system can potentially be later applied to other rhodopsins such as ASR to determine distances within the retinal binding pocket. Isotopically labeled retinal can be further extracted from PR or ASR and incorporated into other proteins with the desired pattern of labeling done independently in other strains of E. coli or yeast.

A B C

Figure 43 – A) Genes encoded by pOrange which control synthesis of beta-carotene. B) pOrange vector map [158]. C) pKJ900 vector map [158].

5.3 Proteorhodopsin

In 2000, metagenomic analysis of marine bacterioplankton revealed the existence of proteorhodopsins [159]. Proteorhodopsins act as light-driven proton pumps, as observed in both marine bacterial membranes [159] and in E. coli expressed recombinant proteorhodopsins [160].

The exact purpose of proteorhodopsins for their host organisms is still unknown, however their

82 production seems to be linked to stress [161]. While proteorhodopsins can generate a proton- motive force, increasing ATP production and thereby increasing flagellar motility, the impact on survival is undetermined [162-164]. Genes encoding proteorhodopsins have been found not only in eubacteria, but also giant viruses [165], archaea [166], and marine protists [167]. Additionally, proteorhodopsin genes have been identified in more than just marine environments, but also fresh water [168], Siberian permafrost, sea ice [169] and in high mountain regions [170].

Proteorhodopsins have a wide range of absorption maxima. Despite 80% homology among proteins of this class, maxima range from 490 nm (blue-absorbing proteorhodopsin, BPR) to 525 nm (green-absorbing proteorhodopsin, GPR). This is considered to be important in view of the availability of light sources in different environments. BPR is found generally in deeper ocean levels where blue light can more easily penetrate. In contrast, GPR is found in surface organisms where green light is more readily available [171]. This study will focus on GPR, heretofore referred to as PR.

5.4 Proteorhodopsin Structure

Similar to all rhodopsins, PR contains seven transmembrane α-helices and a covalently bound retinal molecule in the transmembrane domain. However, unlike ASR, the structure of green-light absorbing PR has not been determined as of yet. The structure of three mutants of BPR were determined via X-ray crystallography in 2011 [172] and structure of another distant homolog in 2013 [170]. However, this success has not been repeated for GPR. X-ray crystallography of membrane proteins is quite challenging due to the amphipathic nature of the molecules. This makes well-diffracting crystals very difficult to obtain [173]. Also, the crystalline state is often not an accurate representation of a native-lipid environment [173]. A solution NMR structure does exist of PR, but there is an RMS difference of 4.219 Å compared to BR [174] and doubts were cast at

83 whether this structure reflects the native conformation of PR [170]. Comparatively, the BPR structure has a RMS difference of 1.150 Å when compared to BR. This is indicative of large structural variations within the solution NMR structure, likely due to micellar environment, low pH, and high temperature. As such, determining the structure of PR within a native lipid environment would assist in revealing the true structural characteristics of PR.

PR shares homology with BR which is associated with several similarities in their photocycles. While both have a Lys Schiff base (Lys 231 in PR), and an Asp primary proton acceptor, the primary proton donor in PR is Glu 108 (vs. Asp96 in BR). These residues follow a similar proton pathway following the isomerisation of retinal from all-trans to 13-cis in the presence of light. One significant difference is that the primary proton acceptor of PR (Asp97) is complexed with a unique histidine residue (His75), which in turn participates in the intermonomer interactions [172, 175]. Additionally, in BR there is a pair of glutamates, Glu 194 and Glu 204, which release the proton into the extracellular media. There exists no homologous pair in PR which means that the proton release mechanism is still unknown. Xanthorhodopsin (XR), another eubacterial proton-pumping rhodopsin, also does not have the proton releasing complex. Instead, a large cavity on the extracellular side of the protein is formed by the displacement of the B-C and

F-G interhelical loop regions of XR [176]. H/D exchange SSNMR experiments in PR have suggested a similar cavity [177]. However, the lack of a determined structure severely limits further functional knowledge of PR.

Figure 44 – Homology model of green-absorbing PR based on template of BPR (PDB ID: 4JQ6) and compiled by Swiss-Model. Model representation is produced in PyMol.

Chemical shift assignments have been previously made by our group using SSNMR for approximately two-thirds of the 238 residues in PR [24, 178]. SSNMR has also been used to probe secondary structures within the protein. A small α-helical structure was found within the E-F loop and a short β-turn in the B-C loop was identified [178]. The conformation of the retinal chromophore has been confirmed through SSNMR, showing the all-trans configuration in PR’s ground state. Additionally, SSNMR has been used to identify changes in the protein induced by a point-mutation in the E-F loop resulting in a 20 nm redshift [179]. SSNMR is a valuable tool in probing the dynamics of the protein. Exchange rates and solvent accessibility have been investigated through H/D exchange experiments [177]. PR has several mobile loops (A-B, C-D, and F-G) which can be coupled to ion transport [178]. Other factors investigated include temperature, hydration and membrane elasticity [180].

Despite the significant number of assignments and distance constraints, we have not been able to determine structure of PR by SSNMR. Additional distance constraints within the retinal binding pocket might prove useful for structure determination. To do so, one requires an isotopically labelled retinal molecule bound within the protein. However, organic synthesis of isotopically labelled retinal is costly, as mentioned above. An alternative is to produce retinal biosynthetically, in the simplest case within the cells expressing PR. As such, growth of PR in labelled media would result in both labelled protein and the labelled retinal chromophore.

5.5 Retinal Labelling Prediction

The sole carbon source to be used in the media for growth of UT5600 (which is a K12 derivative) is glycerol (glucose represses the arabinose-induced pBAD promoter) [181, 182]. As such one can employ alternatively labelled 13C-glycerol to achieve more precise assignments and more distance constraints in the retinal binding pocket. The alternate labeling should allow for measuring longer distances due to the removal of dipolar truncation and simplification of the protein spectra [183]. Two of these glycerols are 1,3-13C-glycerol and 2-13C-glycerol. Glycerol undergoes glycolysis and following the MEP pathway forms DMAPP and IPP which combine to form GPP (Figure 42B). Under this pathway, there is a reductoisomerase which catalyses the DXP to MEP reaction which results in the rearrangement of carbons yielding the derived labelling scheme (Figure 45). A similar pathway is seen in Val synthesis resulting in a similar labelling pattern, in which two central carbon atoms, Cα and Cβ, originate from the second position in glycerol (Figure 46) [184]. Another addition of IPP forms GGPP (Figure 43A). These two GGPP molecules combine to form phytoene which isomerizes to lycopene. Finally, β-carotene is formed and subsequently cleaved by β-carotene dioxygenase to form two retinal molecules (Figure 42A).

Following this pathway one can determine which carbons will be 13C-labeled depending on the

86 glycerol source. If one uses 1,3-13C-glycerol, the carbons outlined in orange in Figure 45 will be

13C. Using 2-13C-glycerol would result in the teal residues being 13C-labelled (Figure 45). As such we can determine which residues will interact with the labelled retinal carbons based on homology modelling of PR with the retinal chromophore included.

16/17

2 16/17 3 1 4 6 5 7 9 11 13 15 8 10 12 14

18 19 20

Figure 45 – Predicted retinal labelling scheme using 1,3-13C-glycerol (orange) and 2-13C-glycerol (teal). Labelling scheme was derived from the MEP pathway. In PR, oxygen will be replaced by nitrogen of Lys sidechain, which is expected to be 15N labeled in both scenarios.

Additional considerations must be applied to the biosynthesis of the amino acids within the protein. Under differential labelling schemes, fully selective or partial labelling can occur (Figure

46). Using 1,3-13C-glycerol, the carbons outlined in blue will be 13C. Using 2-13C-glycerol would result in the red residues being 13C-labelled (Figure 46). Amino acids which are synthesized from products of the citric acid cycle are subject to partial labelling (Figure 46). Also, unlabelled amino acids added to the growth medium (UT5600 needs exogenous Pro, Trp, and Leu [185]) would not normally contribute to the NMR spectra, even though they can be added in their isotopically labelled forms if needed otherwise. Therefore, when identifying cross-peaks, the type of glycerol used must be taken into account.

Figure 46 – Predicted labelling scheme for amino acids from cells grown in 1,3-13C-glycerol (blue) and 2-13C- glycerol (red) [186].

Using this labelling scheme and the predicted structure within the retinal binding pocket, one can derive the interactions which would occur within PR and be detectable by SSNMR.

Prediction was based on residues being within 4 Å of the retinal molecule in the BPR-based homology model (Figures 44, 47). These interactions are predicted based on carbon-carbon interactions and carbon-nitrogen interactions resulting in the different cross-peaks which would arise due to the different labelling schemes (Table 3). This list is not exhaustive as nuclei with separation longer than 4 Å may give dipolar correlations as well. It should be noted that using

13C13,13C20-labeled retinal our group has previously identified proximities of C13 to V102 and long-range correlations of C20 with I71, M134, and L135, which could not be explained by homology modeling, but may originate from relay transfer of magnetization (Shi et al, unpublished).

Table 3 – Predicted atom proximities within 4 Å detectable by SSNMR within the retinal binding pocket of PR. P, W, and L will not be detectable unless provided in the isotopically labelled form and are labelled with asterisks PR Residues Residue Atom 1,3-13C-glycerol 2-13C-glycerol W98* CD1 C12 C14 NE1 C12 C14 V102 CG2 C12, C20 C13 L105* CD2 C20 M134 CG C18 CE C7, C8, C19 C5, C6, C9 G138 CA C3 C2 W159* CB C18 W197* CE3 C19 Y200 C C16 CB C7, C8, C16 C9, C10 CG C8 C9, C10 CD1 C10 CD2 C11 C9, C10 CE1 C12 CE2 C11, C12 C10, C13 CZ C11, C12 C13 P201* CB C4 C5 Y204 CD2 C2 K231 CG C15 CD C15 CE C15 C14 NZ C15 C13, C14

W98 G138

V102

M134

Y204 L105 K231 Y200

P201 W197

W159

Figure 47 - Interactions in the retinal binding pocket as predicted based on the PR homology model based on BPR (Figure 44). Retinal coloured according to predicted labelling scheme as described in figure 45. Model was generated in PyMol.

Using these predicted cross-peaks and the chemical shifts associated with 13C-labelled retinal, one can derive distances within the retinal binding pocket of PR, using previously obtained assignments for the protein resonances. Chemical shift assignments for 13C-labelled retinal atoms have previously been found for free retinal and its Schiff base in solution, and in solid-state for BR in both isomeric states (all-trans-BR568 and 13-cis-BR548), and PR (Table 4). The differences in the chemical environment between BR and PR is clearly visible in the differences in the chemical shift of a retinal carbon between different proteins. However, it has been found that PR may contain some 13-cis-retinal under certain conditions [187, 188]. This may result in peak doubling within the spectra, but the weaker peaks originating from 13-cis- species will be likely undetectable.

While there are some differences in the binding pocket of PR as compared to BR, these chemical shifts should be fairly distinct in the SSNMR spectra. As such, the retinal peaks can be easily identified assuming no scrambling occurred during retinal synthesis. Likely, some peaks will be obscured in the 1D spectrum by protein resonances, requiring more dimensions for assignments.

In the 2D spectra, carbon-carbon correlation peaks can then be assigned and distances within the retinal binding pocket refined.

Table 4 – Chemical shifts of all-trans-retinal and 13-cis-retinal in solution and in BR for all 13C-labelled carbon atoms. Retinal and BR shifts adapted from Harbison et al. [189] and [190] respectively. PR peaks were adapted from Pfleger et al. [179], Mehler et al. [191], Reckel et al. [174], and Shi et al. (Unpublished data) Position All-trans-retinal 13-cis-retinal BR568 BR548 PR 1 33.7 34.3/ 34.9 2 40.2 39.5/ 43.5 3 20.4 4 32.4 34.4/ 35.6 5 128.5 126.7/ 136.8 144.8 144.8 6 134.2 136.8/ 139.5 135.4 134.9 7 130.2 129.5 130.7 8 135.15 138.9 132.7 131.6 9 141.4 142.2/ 141.1 146.4 148.4 10 130.2 133.0 129.7 130.0 11 133.9 139.1 135.4 138.5 12 134.0 134.3 124.2 13 155.5 155.2/ 153.7 169.0 165.3 166.5 14 129.8 122.0 110.5 120.2 15 189.4 191.85/ 190.1 163.2 165.3 161.7 16 30.9 29.7/ 30.5 17 29.7 28.4/ 29.2 18 21.7 23.6/ 20.8 22.0 22.0 19 13.1 13.25/ 12.6 11.3 11.3 20 13.1 21.60 13.3 13.3 14.5

5.6 Resonance Raman spectroscopy of Proteorhodopsin

Raman spectroscopy is a powerful tool in probing chromophores in a large molecular complex such as PR. Retinal has previously been probed in BR [63, 64] and PR [192] (among others) through this technique. Site-directed isotope labelling in the retinal molecule allowed for

Raman bands to be assigned to the specific bonds and their vibrational modes in BR (Figure 48).

The predominant band located at 1527 cm-1, was identified to be mainly derived from the double bond stretching between C11 and C12, and C9 and C10. Other notable peaks include the methyl deformation band at 1448 cm-1, the single C-C stretching bonds between carbons 10-11, 14-15, 8-

9 and 12-13 located between 1122 cm-1 and 1255 cm-1 (so called fingerprints). The band at 1008 cm-1 is consistent among retinal compounds and has been shown to be related to symmetric in plane rocking of C19 and C20 methyl groups.

Figure 48 – Resonance Raman bands with corresponding major vibration identifications in bacteriorhodopsin [63].

Raman bands are sensitive to minute differences in the Schiff base conformation, retinal isomeric state and planarity, as well as protein environment. PR has previously been analysed through Raman as well in both PR and BPR forms (Figure 49). Mutation analysis in conjunction with Raman spectroscopy managed to elucidate key residues which differentiate the two homologs in light absorption (Leu105 in PR vs Gln205 in BPR) [192]. In comparing the bands in PR with

BR, there are some notable differences. The ethylenic stretch group peak at 1527 cm-1 in BR is shifted to 1533 cm-1 in PR, consistent with the well-known correlation to the absorption in the visible [193]. The methyl deformation peak at 1458 cm-1 in BR also shifts to 1448 cm-1 in PR.

However, the 1008 cm-1 peak pertaining to the methyl groups C19 and C20 is consistent between these two proteins. Carbon-carbon single bonds stretches are different with PR lacking the 1211

92 cm-1 band seen in BR and having much lower amplitude of the 1170 cm-1 band. However, the similarities are such that the extent of labelling can be analysed by this method using isotopic shifts of the vibrational bands as determined under 13C-labelling of retinal in BR.

Figure 49 – Raman spectra of the retinal chromophore in green absorbing PR (GPR) and in blue-absorbing PR (BPR) [192].

Previous work on BR has systematically determined the spectral shifts associated with the substitution of retinal carbons with their 13C isotope [63]. Given that each band is related to a specific vibrational mode within the molecule, the shift can be calculated for each of the predicted isotope labelling schemes. In reality, the situation is more complex, as the most bands are combinations of several different vibrations and some are products of resonance of several vibrations, which will shift upon change in frequency of any of the components. BR bands which can help determine the labelling efficiency of the protocol include 1639 cm-1 (C=N stretch) and

1007 cm-1 (Table 5). Since the methyl groups and C15 are not supposed to be labelled in the 2-

93 glycerol labelling scheme, their associated bands are not expected to be shifted as compared to the

1,3-glycerol derived retinal (Table 5). It should be noted, though, that the Schiff base nitrogen will be labeled in both schemes, producing a downshift of the C=N stretch.

Table 5 – Expected shifts in the BR spectrum given the predicted retinal labelling scheme. Shifts calculated from Smith et al. [63]. See figure 48 for the assignments of major corresponding vibrations. BR Band 1,3-13C-Glycerol 2-13C-Glycerol 1639 1609 1626 1527 1507 1504 1456 1456 1456 1255 1250 1245 1248 1244 1235 1214 1210 1195 1201 1193 1186 1169 1159 1145 1007 1003 1007

5.7 Materials and Methods

5.7.1 Materials

5.7.2 Expression of βUT5600-PR in 2XYT

UT5600 E. coli cells containing pOrange and pKJ900 were plated on 2XYT media and incubated overnight at 30 °C. An orange colony (producing β-carotene) was chosen for PR expression screening. Cells were grown overnight in four 2 mL 2XYT shaking at 275 rpm at 30

°C with 100 µg/mL ampicillin and 50 µg/mL chloramphenicol. Cells were then resuspended in four 250 mL baffled flasks in 25 mL of 2XYT with ampicillin and chloramphenicol to an OD of

0.4. Cells were either not induced or induced with 1 mM IPTG and 7.5 µM all-trans-retinal, or 1 mM IPTG and 0.2 % L-arabinose, or 0.2 % L-arabinose. Under different induction conditions, the cells will exhibit different colours according to the genes encoded by the plasmids pOrange and pKJ900 (Figure 43B-C). L-arabinose will induce the production of beta-carotene 15’-15’- dioxygenase, which catalyzes the transformation of beta-carotene into two retinal molecules. Thus, when in the presence of only this inducer, the cells will become white due to the loss of beta-

94 carotene. IPTG will induce the production of PR. Ergo, when IPTG is added in the presence of either L-arabinose or exogenous all-trans-retinal, the cells will exhibit a pink colour indicative of the presence of retinal-bound PR. However, without L-arabinose, these cells will not remove the excess beta-carotene resulting in some orange colour as well. The cells were incubated for 21 hours after induction, shaking at 275 rpm at 30 °C. Cells were spun down at 5000 x g for 15 min and colours were observed.

5.7.3 Expression of βUT5600-PR in M9+PWL

The orange colony that produced pink cells under IPTG and L-arabinose induction was chosen for expression in minimal media. Previous studies indicated that the cells needed proline, leucine and tryptophan in order to produce β-carotene. Cells were grown in M9 media with the addition of 1 mM Pro, 1 mM Leu, and 1 mM Trp. However, no growth was observed. Five cultures of varying Trp concentrations (2, 5, 10, 20, and 30 mM) were grown overnight in 2 mL M9 cultures with 1 mM Pro, and 1 mM Leu. Cells were then resuspended to an OD of 0.1 and allowed to grow until they reached an OD of 0.4 and were induced with 1 mM IPTG and 0.2 % (w/v) L-arabinose.

A large scale culture with 1 mM P, L and 30 mM W was cultured in M9 minimal medium with glycerol and ammonium chloride as the sole carbon and nitrogen sources respectively. A 2 mL culture was induced with 20 μL of cells stored at -20 °C and grown for 24 hours at 30 °C and

275 rpm. The 2 mL culture was used to inoculate a 25 mL culture which was grown overnight at

30 °C and 275 rpm. The 25 mL culture was subsequently used to inoculate 1 L of M9 media and grown at 30 °C and 275 rpm. The cells were induced at an optical density of 0.4 with 0.2 % L- arabinose and 1 mM Isopropyl-β-D-thiogalactoside (IPTG). The cells were collected 21 hours after induction at 5000 × g and 4 °C.

5.7.4 Purification of βUT5600-PR

Proteorhodopsin was purified in the same manner as FL-ASR with some alterations. The broken membrane pellet was solubilized in 1% Triton X-100 overnight at 4 °C with gentle stirring.

The first four washes were performed in 0.05 % Triton X-100 before switching to 0.05 % DDM for all subsequent steps.

5.7.5 Reconstitution of βUT5600-PR

Dry powder lipids, 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) and 1,2- dimyristoyl-sn-glycero-3-phosphate (DMPA), were first dissolved and mixed in chloroform in a ratio of 9:1 (w/w) (DMPC/DMPA, Avanti Polar Lipids). The chloroform was removed through vacuum evaporation to form a lipid film. The lipid film was then thoroughly dehydrated using vacuum desiccation for 5 hours. The lipids were rehydrated with reconstitution buffer (5 mM

NaCl, 10 mM Tris, pH 8) to form a 11.1 mg/mL suspension.

Prior to reconstitution, unlabelled retinal was added to the protein mixture to ensure all PR present was in its chromophore-bound state. The protein and lipids were combined at a ratio of

2 :1 :0.5 (w/w/w) (protein :lipid :Triton X-100) and incubated at 4 °C for 6 hours. Bio-beads SM-

2 (Bio-rad) were added at 0.6 g per mL of mixture and incubated at 4 °C with mixing for 24 hours for detergent removal. The proteoliposomes were collected using a syringe and the beads were washed in 200 mM NaCl until the turbidity was negligible. The collected proteoliposome suspension was centrifuged at 150,000 × g, 4 °C for 50 minutes. The pelleted proteoliposomes were washed several times in NMR buffer, pH 9 and spun down for analysis.

5.7.6 Raman Spectroscopy

Proteoliposomes were spun down and resuspended in the minimal volume of NMR buffer, pH9. The resulting thick suspension was transferred into a glass cuvette, sealed to prevent drying, and FT-Raman spectra were measured in Bruker FRA106/s accessory to the IFS66vs spectrometer, with excitation at 1024 nm, at 2 cm-1 resolution.

5.7.7 SSNMR experiments

Proteoliposomes were spun down at 900,000xg for 9 hours and center-packed into a 3.2 mm rotor (Bruker). 1D spectra were taken for both 13C and 15N. DARR experiments were performed with mixing times of 50 ms and 200 ms. Experiments were run on Bruker Avance III spectrometer at the frequency of 800.230 MHz.

5.8 Results and Discussion

5.8.1 Expression of PR in enriched media

In 2XYT media, the cells were expressed under the control of different inducers. When cells were allowed to grow without any inducer, they remained yellow signifying the presence of beta-carotene which is produced continuously in the cells (Figure 50A). Under L-arabinose and

IPTG induction, the cells showed the characteristic pink colour seen in PR producing cells with a bound retinal, indicative of the presence of the synthesized retinal and proper over-expression of

PR (Figure 50B). Induction in the presence of exogenous all-trans-retinal and IPTG resulted in an orange-red colour (Figure 50C). Finally, L-arabinose induction resulted in white cells (Figure

50D), as expected from production of retinal from beta-carotene (see section 5.7.2 above).

A B C D

Figure 50 – βUT5600-PR cells expressed in 2XYT under different induction conditions. A) No inducer added. B) 0.2 % L-arabinose and 1 mM IPTG. C) 7.5 µM all-trans-retinal and 1 mM IPTG. D) 0.2 % L-arabinose.

5.8.2 Expression of βUT5600-PR in minimal media

When expressed in M9 media, βUT5600-PR requires the addition of three amino acids:

Pro, Leu and Trp. UT5600, the E. coli strain used, exhibits auxotrophy for these amino acids and as such cannot produce P, W and L independently [185]. Also, the constant production of β- carotene introduces a large amount of stress on the cells increasing their nutritional requirements.

Specifically, the sufficient amount of the amino acids P, W, and L need to be added to achieve proper expression. While P and L can be added in low amounts (1 mM), it was found that W must be added in concentrations of 10 mM or higher (Figure 51). Also, it seems the cells readily shed the pOrange plasmid and therefore must be re-plated prior to expression. If all of the above requirements were satisfied, the red-colored cells were produced without addition of external retinal and isotopically labeled PR could be obtained with yield of a few mg per liter of culture.

A B C D E

Figure 51 – βUT5600-PR expressed with addition of A) 30 mM W, B) 20 mM W, C) 10 mM W, D) 5 mM W, and E) 2 mM W in the presence of 1 mM P and L in glycerol-based M9 medium.

5.8.3 Raman Spectroscopy

Raman spectroscopy was used to investigate the isotope labeling pattern of retinal, using purified PR reconstituted into proteoliposomes for better signal-to-noise ratio and purity. The resulting spectrum of natural abundance PR was very close to the one previously reported (Figure

49 above). There was a large peak related to double bond stretches between carbons 11 and 12, and carbons 9 and 10 at 1536 cm-1 (Figure 52). The methyl deformation peak at 1448 cm-1 and the

1008 cm-1 peak pertaining to the methyl groups C19 and C20 are both present (Figure 52).

Furthermore, excellent resolution between 1162 cm-1 and 1198 cm-1, the fingerprints region, can be seen (Figure 52). The peak at 1654 cm-1 is correlated to the C=N stretching of the Schiff base.

Through comparison of this natural abundance spectra with that of the labelled retinal one can estimate the extent and pattern of labelling within the molecule.

Figure 52 – Resonance Raman spectrum of natural abundance βUT5600 PR in DMPC/DMPA proteoliposomes.

The first labelled sample was produced with 1,3-13C-glycerol as the sole carbon source.

Figure 53 shows a comparison of Raman spectra of the labeled and natural abundance PR, with the spectral quality of the former being much lower because of the small amount of the sample used (Figure 53). Thus, the Raman analysis is restricted to the major bands only in this case. A large shift is expected in the 1654 cm-1 C=N stretching peak due to the labelling of both 15N and

13C (C15) of the Schiff base, however, given the amount of noise in this region no solid conclusion can be made. However, from the appearance of the region, there appears to be fractional labelling, as at least two downshifted bands (albeit unresolved) are present. The main ethylenic stretch peak is divided into two (downshifted and unshifted) in the labelled sample indicative of partial labelling

(Figure 53). The shifted peak (1509 vs 1536 cm-1) does conform to the expected isotope shift based on the labelling prediction (Table 6). However, this divided peak is expected considering that the labelling scheme predicts two fifths of the ethylenic regions will contain 13C=13C bonds (Figure

45). In general, the shifts expected for the C-C single bond stretches in the fingerprint regions conform to their expected values, even though it looks that significant fraction of the peaks is unshifted again (Table 6). Additionally, the methyl rocking band associated with C19 and C20 does exhibit the anticipated shift implying that no scrambling occurred in methyl group labelling

(Table 6). In summary, results from Raman spectroscopy conform to the predicted labeling pattern but suggest that there may be incomplete labeling, possibly due to the spin dilution by products of unlabeled Trp added in large excess.

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Figure 53 – Normalised Resonance Raman spectra of natural abundance (red) and 1,3-13C-glycerol-derived (blue) PR in DMPC/DMPA proteoliposomes. Major peaks in the labelled sample are identified in Table 6.

Table 6 – Resonance Raman peaks in 1,3-13C-glycerol-derived PR sample. Expected shifts are calculated from Table 5. Vibrational Mode Natural 13C-labelled Peak Expected Observed Abundance Peak (cm-1) Shift Shift (cm-1) C=NH 1654 1653a 30 1 C11=C12 + C9=C10 1536 1536 23 0 1509 23 27 CH3 deformation 1453 1454 0 -1 Lysine and C12-C13 1253 1249 9 4 C8-C9 1234 1225 10 9 C14-C15 1201 1188 18 13 C10-C11 1173 1157 18 16 C19 & C20 Methyl Rocking 1007 1002b 4 5 aSeveral downshifted peaks are observed in addition but not resolved. bRepeated higher resolution measurement (not shown) showed 1000 cm-1 position.

A second labelled sample with a better yield was produced with 2-13C-glycerol as the sole carbon source resulting in higher spectral resolution. The comparison of the Raman spectra of the

101 labelled and natural abundance PR can be seen in figure 54. The shift in the C=N peak due to the labelling of just the Schiff base nitrogen is visible. However, the presence of both 1656 cm-1 and

1638 cm-1 bands in the labelled sample indicates scrambling of nitrogen labelling. The main ethylenic stretch peak in the labelled sample is divided with upshifted and downshifted components at 1548 cm-1 and 1504 cm-1 respectively (Figure 54). In 2-glycerol labelling, three- fifths of the ethylenic region is expected to be labelled. As such, a divided peak is to be expected.

The downshifted peak does agree with the predicted shift (Table 7). The upshifted peak is counterintuitive, but the enhancement of the upshifted ethylenic modes has been observed in BR upon single 13C labeling [63] and our data may represent the extreme case of such enhancement in the case of multiple labeling. In the single C-C bond region, numerous shifts are expected due to the predicted 13C-C labelling scheme. The majority of these are seen in the labelled spectrum

(Table 7). There is an unanticipated shift in the C19-20 methyl rocking peak, but it is consistent with the shift observed in BR upon 13C13 labeling [63] (Table 7). Overall, this spectrum seems to concur with the predicted labelling scheme while also confirming the incomplete labelling suggested by 1,3-13C-glycerol-derived sample data.

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Figure 54 – Resonance Raman spectra of natural abundance (cyan) and 2-13C-glycerol-derived (purple) PR in DMPC/DMPA proteoliposomes. Major peaks in the labelled sample are identified in Table 7.

Table 7 - Resonance Raman peaks in 2-13C-glycerol-derived PR sample. Expected shifts are calculated from Table 5. Vibrational Mode Natural 13C-labelled Peak Expected Observed Abundance Peak (cm-1) Shift Shift (cm-1) C=NH 1654 1656 13 -2 1638 16 C11=C12 + C9=C10 1536 1548 23 -12 1504 22 CH3 deformation 1453 1451 0 -2 C8-C9 1234 1223 13 11 C14-C15 1201 1186 19 15 C3-C4 1173 1149 24 24 C19 & C20 Methyl Rocking 1007 1003 0 4

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5.8.4 SSNMR spectroscopy of βUT5600-PR

First, the 1D 15N spectrum was measured for βUT5600-PR. The spectrum was compared to a previous PR sample prepared in BL21 cells grown on 2-13C-glycerol with exogenous retinal

13C-labelled at C13 and C20 (henceforth referred to as BL21-PR) (shown in red in Figure 55).

Overall, the spectral resolution and secondary structure between the two samples was comparable.

Additionally, the Schiff base peak is located at the same position. However, there are some differences which most likely originated from the addition of unlabelled Pro, Trp, and Leu. Firstly, the Pro peak visible in BL21-PR is not present in the βUT5600-PR (Figure 55). Additionally, the spectral bands corresponding to Arg and Lys are different between the samples, with the relative amplitude of Arg bands (if normalized to the backbone peak) being much larger in the βUT5600-

PR. While this is most likely to reflect the removal of many P, W, L backbone nitrogens, it may be also indicative of different scrambling in the samples, which is not unexpected in 2-13C-glycerol growths. This may arise in one of two different ways. Firstly, the addition of the large quantity of unlabelled Trp could possibly result in the incorporation of its natural abundance atoms. Secondly,

UT5600 is a K-12 derivative. As such, previous studies have found that while K-12 E. coli do not have the MVA pathway, they do contain an efficient citric acid cycle [194]. Thus, any amino acids produced through TCA derived precursors in UT5600, will exhibit scrambling which may differ from the scrambling observed in BL21.

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15N chemical Shift (ppm)

Figure 55 – Normalised 15N 1D SSNMR spectra of βUT5600-PR (red) and BL21-PR (black). Both samples were produced using 2-13C-glycerol.

In comparing the βUT5600-PR and BL21-PR 1D 13C SSNMR spectra, there are several peaks which can be assigned to the labelled carbons in the retinal molecule. Peaks at 165.9, 147,

139.7, 137, 132 and 121.8 ppm can be assigned to C13, C9, C6, C5, C10, and C14 respectively

(Figure 56). These tentative assignments can be done using known chemical shifts for retinal in other proteins, as well as previously determined shifts for C14, C13, and C10 for PR (see Table 4 above), but must be verified by carbon-carbon correlations in 2D spectra (see below). C1 and C2, located on the retinal ring, should be seen around 30-40 ppm, but cannot be resolved in 1D.

Additionally, the reduced signal as a result of the addition of Leu, around 40 ppm for Cβ and 25 ppm for Cγ (26 residues in total), and Trp, around 110 and 120 ppm (10 residues in total), can be seen through comparison of these samples. Further evidence of different scrambling as a result of

105 using UT5600 instead of BL21 can be seen in the Cb region around 30 ppm, and CO region around

180 ppm, even though a significant part of the difference in the latter comes from the removal of

Leu resonances. However, good agreement between the samples can be seen in the CA region, even though the absence of Trp peak around 60 ppm is noticeable (Figure 56).

Figure 56 – 13C 1D SSNMR spectra of βUT5600-PR (black) and BL21-PR (red). Both samples were produced using 2-13C-glycerol.

In order to unambiguously assign retinal carbon chemical shifts, 2D DARR [195, 196] experiments were conducted. Low overall number of crosspeaks for protein atoms argues for incomplete labeling as suggested by other results above. Predicted cross-peaks within the labelled retinal molecule include C1-C2, C5-C6, C6-C1, C9-C10, and C13-C14. Previously, the chemical shift of C13 was assigned in BL21-PR as 166.5 ppm (Table 4). A strong cross-peak in βUT5600-

PR at 165.9/121.8 ppm identifiable as C13/C14 correlation was found, showing no correspondence

106 with any peaks in BL21-PR (Figure 57). Strong crosspeaks were also found at 139.7/37 ppm

(Figure 58) and 147/132 ppm (Figure 59). The lack of spectral overlap with BL21-PR allows for unambiguous assignment in this case. Therefore, these peaks can be assigned to the C6-C1 and the

C9-C10 cross peaks respectively. Additional weak crosspeak was identified in βUT5600-PR at

140 and 137 ppm, which shows some spectral overlap with BL21-PR (Figure 60). However, considering the unambiguous assignments for C6 atom at 139.7 ppm above, this peak can be assigned to the interaction of C6 and C5. The low strength of this crosspeak may be correlated to the large number of adjacent labeled carbons on the retinal ring which could result in dipolar truncation of the signal.

Figure 57 - 13C-13C DARR spectra of βUT5600-PR (black) and BL21-PR (red) showing the region of 165.9 and 121.8 ppm correlation identified as C13-C14 crosspeak. Both samples were produced using 2-13C-glycerol. Spectra were measured at 800 MHz with a mixing time of 50 ms used for both samples.

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Figure 58 – 13C-13C DARR spectra of βUT5600-PR (black) and BL21-PR (red), showing the region of 139.7 and 37 ppm correlation identified as C6-C1 crosspeak. Both samples were produced using 2-13C-glycerol. Spectra were measured at 800 MHz with a mixing time of 50 ms used for both samples.

Figure 59 - 13C-13C DARR spectra of βUT5600-PR (black) and BL21-PR (red), showing the region of 147 and 132 ppm correlation identified as C9-C10 crosspeak. Spectra were measured at 800 MHz with a mixing time of 50 ms used for both samples.

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Figure 60 - 13C-13C DARR spectra of βUT5600-PR (black) and BL21-PR (red), showing the region of 140 and 137 ppm correlation identified as C6-C5 crosspeak. Spectra were measured at 800 MHz with a mixing time of 50 ms used for both samples.

The only predicted cross-peak which could not be identified was that of C1-C2 which is most likely subject to the same dipolar truncation which affects the C5-C6 cross peak signal. C1 was assigned to have a chemical shift of 37 ppm, from the C6-C1 cross-peak. Previous studies have found C2 to have a chemical shift between 39 and 44 ppm depending on the isomerization state (Table 4). However, this region is located close to the main diagonal where the majority of protein peaks are clustered (Figure 61). Also, a longer mixing time of 200 ms did not reveal any additional peaks in this region. As such, one can only unambiguously assign a chemical shift of

37 ppm to C1. This cross-peak could potentially be isolated in future experiments with a different labeling pattern.

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Figure 61 - 13C-13C DARR spectra of βUT5600-PR (black) and BL21-PR (red), shown in the region of 100 to 0 ppm. Putative location of the C1-C2 cross peak is shown by box. Spectra were measured at 800 MHz with a mixing time of 200 ms for βUT5600-PR and 50 ms for BL21-PR.

In summary, the biosynthetic production of retinal within PR allowed for the unambiguous assignment of chemical shifts to seven carbons in the retinal molecule. Chemical shifts of 165.9,

147, 139.7, 137, 132, 121.8, and 37 ppm can be assigned to C13, C9, C6, C5, C10, C14, and C1 respectively (Table 8). Four of these assignments (C1, C5, C6 and C9) were previously unknown for PR (Table 8). Furthermore, the assignment of carbons on the retinal ring (C1, C5 and C6) was the first instance in which they could be assigned in either PR or BR (Table 8). There were differences of 2 ppm between previously determined retinal chemical shifts in PR and that of

βUT5600-PR (Table 8). However, this is most like due to differences in referencing which has been shown to result in a chemical shift of that magnitude. We demonstrated that simultaneous sparse isotope labeling of both 7TM helical protein and its ligand is possible, laying a solid foundation for measuring protein-chromophore distances for structure determination. As such, this

110 expression system shows great promise in investigating the chromophore binding pocket in retinal proteins.

Table 8 – Chemical shifts of all-trans-retinal and 13-cis-retinal in solution and in BR for all 13C-labelled carbon atoms. Retinal and BR shifts adapted from Harbison et al. [189] and [190] respectively. PR peaks were adapted from Pfleger et al. [179], Mehler et al. [191], Reckel et al. [174], and Shi et al. (Unpublished data). βUT5600-PR retinal chemical shifts added for comparison. Position All-trans-retinal 13-cis-retinal BR568 BR548 PR βUT5600-PR 1 33.7 34.3/ 34.9 37.0 2 40.2 39.5/ 43.5 3 20.4 4 32.4 34.4/ 35.6 5 128.5 126.7/ 136.8 144.8 144.8 137.0 6 134.2 136.8/ 139.5 135.4 134.9 139.7 7 130.2 129.5 130.7 8 135.15 138.9 132.7 131.6 9 141.4 142.2/ 141.1 146.4 148.4 147.0 10 130.2 133.0 129.7 130.0 132.0 11 133.9 139.1 135.4 138.5 12 134.0 134.3 124.2 13 155.5 155.2/ 153.7 169.0 165.3 166.5 165.9 14 129.8 122.0 110.5 120.2 121.8 15 189.4 191.85/ 190.1 163.2 165.3 161.7 16 30.9 29.7/ 30.5 17 29.7 28.4/ 29.2 18 21.7 23.6/ 20.8 22.0 22.0 19 13.1 13.25/ 12.6 11.3 11.3 20 13.1 21.60 13.3 13.3 14.5

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Chapter Six

Final conclusions and future directions

The seven transmembrane alpha-helical architecture of A2aR, ASR and PR is ubiquitous in the natural world. This elegant system consisting of a receptor and a ligand contains many different permutations in structure which lead to differences in ligand specificity and protein function. As such, the study of this system of molecules using biophysical methodology is necessary for elucidating the key differences between proteins of this type which dictate their function.

There are many challenges associated with structural investigations of membrane proteins.

Not the least of which is the possibility of contamination within the sample itself. As such, it is necessary to use a variety of biophysical and biochemical methods to analyse sample quality prior to large scale isotopic labelling. Sample contamination can occur as a result of poor expression in the case of A2aR or interactions with the host organism as seen in FL-ASR. FTIR and SSNMR are extremely sensitive to minute changes in secondary structure which can occur as a result of this contamination. In terms of A2aR, any future studies require re-transformation of the DNA into

P. pastoris. By increasing the copy number in the cells, expression will increase accordingly. Only then can its future potential for SSNMR structural studies be decided upon. Furthermore, the ability to monitor differences between the truncated and full-length forms of A2aR through

SSNMR could lead to some interesting results. This is especially relevant in light of the effect truncation has on recombinant ASR.

The effect of the full-length construct of ASR on recombinant expression is significant.

The C-terminal tail of FL-ASR results in accelerated cell death of E. coli during expression. This can be theorized to be a result of interactions of ASR with its co-purification partner, the outer

112 membrane protein OmpF. There exists a precedent for this interaction in the mechanism of the toxin Colicin N. A binding cleft between OmpF monomers could potentially interact with the C- terminal tail of FL-ASR, thereby leading to cell death once FL-ASR production is high enough.

The hydrophilic nature of this tail, and the possibility of in-membrane inversion of FL-ASR allows for this interaction. Any future studies on this protein may necessitate the use of a different expression host which is deficient in OmpF, or late induction labelling, which has been shown to reduce labelling in contaminating proteins for in vivo measurements. Additionally, biochemical analyses of interactions of the protein and ASRT may be undertaken.

Finally, the possibility of concurrently producing a ligand during the expression of a protein in isotopically labelled media is quite attractive for structural studies. The βUT5600-PR system has proven capable of producing isotopically labelled retinal which led to the unambiguous assignment of seven retinal carbons in PR. Three of which were located on the retinal ring, which has previously been unassigned in either BR or PR. Further multidimensional experiments, such as NCOCA could prove useful in further investigation of the scrambling seen in the βUT5600-PR spectra. Additionally, growth with lower Trp concentration should be attempted to reduce the possibility of secondary metabolism induced signal dilution, as the cost of adding labeled Trp would be economically unfeasible. The removal of spin dilution is needed to increase the extent of labeling so that the chromophore-protein distances can be measured in the near future. However, the possibility of using a different K-12 derivative which is not auxotrophic for proline, leucine, and tryptophan could be beneficial given the number of tryptophan residues in the retinal binding pocket. Thus, spin dilution would be decreased while also ensuring these residues would be labelled in the sample.

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Cost is always a prohibitive consideration when undertaking any NMR-based experiments.

Retinal is a ubiquitous ligand in that it is found in signalling pathways across mammalian, archaeal, and microbial organisms. Thusly, the ability to produce this molecule in an efficient manner within labelled media has numerous applications. Extraction of the labelled-retinal after expression in PR would allow this ligand to be used during the expression of other recombinant rhodopsins such as

ASR without requiring additional transformation or growth optimization. Retinal is highly prone to degradation in the cell, therefore one must find a method of extraction which prevents this.

There have been studies which have concurrently produced and extracted retinoids from expression hosts which could be adapted to the βUT5600-PR system. Alternatively, retinal can be extracted from the regenerated βUT5600-PR and isolated in organic solvents. This could be further extended to the production of fully deuterated retinal, the use of which could diminish the proton cloud seen in deuterated rhodopsins wherein the protonated retinal was added as the cost of purchasing fully deuterated retinal is prohibitive.

In conclusion, there exist numerous biophysical methods which can be used to investigate integral membrane proteins. Each heterologous expression system is accompanied by its own unique challenges which must be overcome. As such, one must optimize every step of sample preparation as the quality of the sample is inextricably contingent upon this process. Furthermore, in cases wherein the cost of sample production is higher such as isotope labelling, every effort possible must be used to eliminate contamination and ensure homogeneous samples which will lead to higher quality results with minimal economic impact.

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