Science, Faculty of
Chemistry, Department of
DSpace
UBCV
Vacariu, Condurache
Doctor of Philosophy - PhD
University of British Columbia
The periplasmic pore protein AmpG, normally responsible for the transport of peptidoglycan fragments from the periplasm to the cytosol of Gram-negative bacteria, has been implicated in the biochemical pathway for AmpC β-lactamase induction. Consequently, strains of pathogenic Gram-negative bacteria such as Pseudomonas aeruginosa have developed resistance to a wide swath of antibiotics bearing the β-lactam functional group (i.e. antibiotics of the penicillin family). One approach to combat this growing threat would be to prevent the induction of AmpC’s in the target organisms before treating them with antibiotics. AmpG is ubiquitous and highly conserved among Gram-negative bacteria; it therefore serves as a potential drug target. Unfortunately, relatively little is known about AmpG’s structure and substrate preference owing to the difficulty in both the expression and isolation of this integral membrane protein. Working in collaboration with the Strynadka lab at UBC, who are experts in membrane protein characterization, we sought to elucidate the structure of AmpG – in particular, its glycopeptide substrates’ recognition site – in the hopes of eventually designing an irreversible inhibitor specific to it. Towards this end, we chemically synthesized multimilligram quantities of various putative AmpG substrates – including a large glycopeptide bearing the unnatural meso-oxa-diaminopimelic acid – for our collaborators to test against recombinant AmpG that was purified and reconstituted either in detergent micelles or in peptide nanodiscs. The affinities of our putative substrates towards AmpG are currently being evaluated via isothermal calorimetry. In addition, we synthesized a dendron-based putative inhibitor of AmpG in which a bulky “plug” is attached to an AmpG substrate via a flexible linker: this putative inhibitor will be tested using a lipid planar bilayer or fluorescence assay experiment. Finally, we synthesized a putative AmpG substrate with a compact photoaffinity tag as a probe for the substrate recognition site. We undertook in vitro photolabeling experiments on the AmpG provided to us – unfortunately, we were unable to specifically tag AmpG’s substrate recognition site using our probe.
Synthesis of Tools for Studying the AmpG Pore Protein involved in Antibiotic Resistance by Condurache Vacariu A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Chemistry) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) July 2024 © Condurache Vacariu, 2024 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: Synthesis of Tools for Studying the AmpG Pore Protein involved in Antibiotic Resistance submitted by Condurache Vacariu in partial fulfilment of the requirements for the degree of Doctor of Philosophy in Chemistry Examining Committee: Martin Tanner, Professor, Chemistry, UBC Supervisor Marco Ciufolini, Professor, Chemistry, UBC Supervisory Committee Member Gregory Dake, Professor, Chemistry, UBC University Examiner Russ Algar, Professor, Chemistry, UBC University Examiner Additional Supervisory Committee Members: Raymond Andersen, Professor, Chemistry, UBC Supervisory Committee Member Hongbin Li, Professor, Chemistry, UBC Supervisory Committee Member iii Abstract The periplasmic pore protein AmpG, normally responsible for the transport of peptidoglycan fragments from the periplasm to the cytosol of Gram-negative bacteria, has been implicated in the biochemical pathway for AmpC β-lactamase induction. Consequently, strains of pathogenic Gram-negative bacteria such as Pseudomonas aeruginosa have developed resistance to a wide swath of antibiotics bearing the β-lactam functional group (i.e. antibiotics of the penicillin family). One approach to combat this growing threat would be to prevent the induction of AmpC’s in the target organisms before treating them with antibiotics. AmpG is ubiquitous and highly conserved among Gram-negative bacteria; it therefore serves as a potential drug target. Unfortunately, relatively little is known about AmpG’s structure and substrate preference owing to the difficulty in both the expression and isolation of this integral membrane protein. Working in collaboration with the Strynadka lab at UBC, who are experts in membrane protein characterization, we sought to elucidate the structure of AmpG – in particular, its glycopeptide substrates’ recognition site – in the hopes of eventually designing an irreversible inhibitor specific to it. Towards this end, we chemically synthesized multimilligram quantities of various putative AmpG substrates – including a large glycopeptide bearing the unnatural meso-oxa-diaminopimelic acid – for our collaborators to test against recombinant AmpG that was purified and reconstituted either in detergent micelles or in peptide nanodiscs. The affinities of our putative substrates towards AmpG are currently being evaluated via isothermal calorimetry. In addition, we synthesized a dendron-based putative inhibitor of AmpG in which a bulky “plug” is attached to an AmpG substrate via a flexible linker: this putative inhibitor will be tested using a lipid planar bilayer or fluorescence assay experiment. Finally, we synthesized a putative iv AmpG substrate with a compact photoaffinity tag as a probe for the substrate recognition site. We undertook in vitro photolabeling experiments on the AmpG provided to us – unfortunately, we were unable to specifically tag AmpG’s substrate recognition site using our probe. v Lay Summary To combat growing antibiotic resistance in certain strains of deadly bacteria, we hoped to target one of the bacterial protein mediators indirectly responsible for this troublesome acquired immunity. However, the target protein, called “AmpG,” is still mysterious since it is notoriously difficult to isolate and manipulate; without a thorough understanding of the structure of the target protein, we cannot adequately design drugs to suppress its function. We chemically synthesized molecules that should be recognized by AmpG so that, with help from our collaborators’ expertise in protein science, conclusive structural insights about the protein may be made. Studies are currently underway. We synthesized a preliminary inhibitor to “plug” AmpG and prevent it properly functioning. Finally, we synthesized and tested a molecular probe that should have successfully sought and irreversibly attached itself to AmpG; thereby stopping its function – unfortunately, it seems that our probe failed to specifically target AmpG. vi Preface A portion of Chapters 1 and 2 has been published and some figures are reproduced with permission from the following article: Condurache M. Vacariu and Martin E. Tanner. Chem. Eur. J. 2022, 28, e202200788 (© 2022 Wiley-VCH GmbH). A portion of Chapters 2 has been published with permission from the following article: Arvind S. Soni, * Condurache M. Vacariu, * Jeff Y. Chen, and Martin E. Tanner. Org. Lett. 2020, 22, 6, 2313–2317. (© 2020 American Chemical Society). * These authors contributed equally to the manuscript. All the synthetic experiments were completed by the author of this thesis under the supervision of Prof. Martin E. Tanner and all the microbiological studies – apart from the photoaffinity labeling experiments discussed herein, which were also completed by the author of this thesis – were completed by Helena Sverak under the supervision of Prof. Natalie C. J. Strynadka. vii Table of Contents Abstract ....................................................................................................................................................... iii Lay Summary .............................................................................................................................................. v Preface ......................................................................................................................................................... vi Table of Contents ...................................................................................................................................... vii List of Figures ............................................................................................................................................. xi List of Abbreviations ............................................................................................................................... xvi Acknowledgements ................................................................................................................................ xxiii Dedication ................................................................................................................................................ xxv Chapter 1: Introduction ............................................................................................................................. 1 1.1 Peptidoglycan ...................................................................................................................................... 1 1.1.1 Peptidoglycan Structure and Biosynthesis ................................................................................... 1 1.1.2 Peptidoglycan Recycling and Antibiotics .................................................................................... 4 1.1.3 Antibiotic Resistance and AmpC ................................................................................................. 6 1.2 AmpG .................................................................................................................................................. 9 1.2.1 History and Characterization ....................................................................................................... 9 1.2.2 Structural Modelling .................................................................................................................. 13 1.2.3 Contemporary Assays ................................................................................................................ 17 1.3 Thesis Goals ...................................................................................................................................... 22 Chapter 2: Synthesis and Testing of AmpG Substrates ........................................................................ 25 2.1 Anhydro-PG Fragments .................................................................................................................... 25 2.1.1 Uses of Anyhdro-PG Fragments in studying PG-Binding Enzymes and Proteins..................... 25 2.1.2 Meso-Dap ................................................................................................................................... 27 2.1.3 Meso-oxa-Dap ............................................................................................................................ 33 2.1.4 Chapter 2 Project Goals ............................................................................................................. 35 2.2 The Synthesis of Potential AmpG Substrates ................................................................................... 36 2.2.1 Synthesis of the Glycosyl Acceptor ........................................................................................... 36 2.2.2 Glycosylation of 1,6 Anhydro-MurNAc Methyl Ester .............................................................. 38 2.2.3 Peptide Coupling ........................................................................................................................ 44 2.3 Our Collaborators’ Work .................................................................................................................. 57 2.3.1 Expressing and Isolating AmpG ................................................................................................ 57 2.3.2 AmpG Crystallography .............................................................................................................. 59 viii 2.3.3 AmpG Electron Microscopy ...................................................................................................... 61 2.3.4 Planar Lipid Bilayer Studies with AmpG .................................................................................. 62 2.3.5 Calorimetric Studies on AmpG .................................................................................................. 65 Chapter 3: Synthesis of Small Molecule AmpG Plug ............................................................................ 73 3.1 Obstructing a Permease ..................................................................................................................... 73 3.1.1 Proposed Small Molecule Plug .................................................................................................. 73 3.1.2 Protein Plug Domains ................................................................................................................ 74 3.1.3 Planar Lipid Bilayer Inhibition Experiments ............................................................................. 76 3.2 Synthesis of Compound 6 ................................................................................................................. 79 3.2.1 Dendron Framework .................................................................................................................. 79 3.2.2 Linking Sugar and Dendron ....................................................................................................... 81 3.3 Assays for Transport Inhibition by Small Molecule Plug ................................................................. 83 Chapter 4: Synthesis and Evaluation of An AmpG Photoaffinity Probe............................................. 86 4.1 Background ....................................................................................................................................... 86 4.1.1 Photoaffinity Theory .................................................................................................................. 86 4.1.2 “Big Three” Photoaffinity Groups ............................................................................................. 88 4.2 The Design of an AmpG Photoaffinity Probe ................................................................................... 93 4.2.1 Choice of Photofunctional Group .............................................................................................. 93 4.2.2 Choosing an AmpG Substrate Group ......................................................................................... 98 4.2.3 Choosing a Fluorescent Reporter ............................................................................................... 99 4.2.4 Synopsis of AmpG Photolabeling Experiment ........................................................................ 100 4.3 Chemical Synthesis of the AmpG Photoaffinity Probe................................................................... 102 4.3.1 Attempt to Follow Literature Synthesis of Diazirine Tag ........................................................ 102 4.3.2 Alternative Synthetic Route to Diazirine Tag .......................................................................... 104 4.4 Reacting Photoaffinity Probe 7 with AmpG ................................................................................... 110 4.4.1 Covalent Binding Between AmpG and Probe 7 ...................................................................... 110 4.4.2 Selectivity of Probe Towards AmpG Active Site .................................................................... 113 Chapter 5: Overview and Future Work ............................................................................................... 121 5.1 Thesis Summary .............................................................................................................................. 121 5.1.1 Synthetic Contribution ............................................................................................................. 121 5.1.2 Biological Contribution............................................................................................................ 123 5.2 Future Work .................................................................................................................................... 124 5.2.1 AmpG Substrates ..................................................................................................................... 124 5.2.2 AmpG Plug Inhibitors .............................................................................................................. 128 5.2.3 AmpG Photoaffinity Probes ..................................................................................................... 129 ix 5.2.4 Establishing a Proton Gradient ................................................................................................ 131 Experimental ........................................................................................................................................... 134 General Information .............................................................................................................................. 134 Photoaffinity Labeling Experiments ..................................................................................................... 135 Synthetic Procedures ............................................................................................................................. 137 Compound 2 ...................................................................................................................................... 137 Compound 3 ...................................................................................................................................... 138 Compound 4 ...................................................................................................................................... 139 Compound 5 ...................................................................................................................................... 140 Compound 6 ...................................................................................................................................... 142 Compound 7 ...................................................................................................................................... 143 Compound 9 ...................................................................................................................................... 144 Compound 14 .................................................................................................................................... 145 Compound 15 .................................................................................................................................... 146 Compound 16 .................................................................................................................................... 147 Compound 18 .................................................................................................................................... 148 Compound 21 .................................................................................................................................... 150 Compound 22 .................................................................................................................................... 151 Compound 23 .................................................................................................................................... 152 Compound 24 .................................................................................................................................... 154 Compound 25 .................................................................................................................................... 155 Compound 26 .................................................................................................................................... 156 Compound 27 .................................................................................................................................... 158 Compound 28 .................................................................................................................................... 160 Compound 29 .................................................................................................................................... 161 Compound 30 .................................................................................................................................... 162 Compound 31 .................................................................................................................................... 164 Compound 36 .................................................................................................................................... 166 Compound 37 .................................................................................................................................... 167 Compound 38 .................................................................................................................................... 168 Compound 39 .................................................................................................................................... 170 Compound 45 .................................................................................................................................... 172 Compound 46 .................................................................................................................................... 173 Compound 47 .................................................................................................................................... 174 Compound 48 .................................................................................................................................... 175 x Compound 49 .................................................................................................................................... 176 Compound 50 .................................................................................................................................... 177 Compound 51 .................................................................................................................................... 178 Compound 52 .................................................................................................................................... 179 Compound 53 .................................................................................................................................... 181 Compound 54 .................................................................................................................................... 182 Compound 55 .................................................................................................................................... 183 Compound 56 .................................................................................................................................... 184 Compound 57 .................................................................................................................................... 185 Compound 58 .................................................................................................................................... 187 Bibliography ............................................................................................................................................ 189 Appendix: NMR Data ............................................................................................................................. 204 xi List of Figures Figure 1.1 The structure and biosynthesis of peptidoglycan in Gram-negative bacteria ....................... 3 Figure 1.2 Enzymes responsible for the cleavage of the peptidoglycan glycosidic bonds .................... 4 Figure 1.3 Various β-lactam antibiotics ................................................................................................. 5 Figure 1.4 PG recycling and induction pathway of AmpC by β-lactam antibiotics .............................. 7 Figure 1.5 The “rocker-switch” mechanism of passive transport ........................................................ 10 Figure 1.6 Membrane topology diagram of E. coli AmpG .................................................................. 11 Figure 1.7 Preference of E. coli AmpG uptake of PG fragments into bacteria .................................... 12 Figure 1.8 H+-symport “rocker-switch” mechanism for a major facilitator ......................................... 13 Figure 1.9 Modelled P. aeruginosa AmpG membrane topology ......................................................... 14 Figure 1.10 Modelled P. aeruginosa AmpG tertiary structure ............................................................ 14 Figure 1.11 3-D model of N. gonorrhoeae AmpG ............................................................................... 15 Figure 1.12 3-D model of K. aerogenes AmpG ................................................................................... 15 Figure 1.13 Summary of Vocadlo’s fluorescence assay ...................................................................... 18 Figure 1.14 The results obtained with Vocadlo’s various AmpG probes ............................................ 19 Figure 1.15 Summary of the HTS assay methodology for finding AmpG inhibitors .......................... 21 Figure 1.16 AmpG substrates, inhibitors, and photoaffinity probes prepared in this thesis ................ 24 Figure 2.1 Examples of previously synthesized 1,6-anhydro-PG fragments ....................................... 26 Figure 2.2 The lytic transglycosylase, SltB3, holoenzyme crystal structure ....................................... 27 Figure 2.3 Meso-diaminopimelic acid (meso-Dap) in the stem peptide of Gram-negative bacteria .... 28 Figure 2.4 Trans-peptidation (cross-linking) reaction of PG layers via penicillin-binding proteins (PBP’s) .................................................................................................................................... 29 Figure 2.5 Chemical syntheses of meso-Dap from literature ............................................................... 31 xii Figure 2.6 Previous chemical syntheses of meso-Dap glycopeptides .................................................. 32 Figure 2.7 Vederas’ chemical synthesis of meso-oxa-Dap .................................................................. 33 Figure 2.8 Dr. Soni’s meso-oxa-Dap tripeptide synthesis .................................................................... 34 Figure 2.9 AmpG substrates as synthetic targets ................................................................................. 36 Figure 2.10 Chemical synthesis of the 1,6-anhydro-MurNAc methyl ester 13. .................................. 38 Figure 2.11 Preliminary glycosyl donor and glycosylation attempts ................................................... 39 Figure 2.12 Glycosyl bromide synthesis and subsequent Koenigs-Knorr glycosylation ..................... 40 Figure 2.13 Attempted deprotection of the phthalimide group of compound 16 ................................. 41 Figure 2.14 Dr. Hadi’s use of anchimeric assistance from a trichloromethyl group ........................... 42 Figure 2.15 The final glycosylation method used in the disaccharide synthesis ................................. 43 Figure 2.16 Dehalogenation and synthesis of GlcNAc-1,6-anhydro-MurNAc 1 ................................. 44 Figure 2.17 Synthesis of GlcNAc-1,6-anhydro-MurNAc-L-Ala 2 ...................................................... 45 Figure 2.18 Elaboration of 1 into GlcNAc-1,6-anhydro-MurNAc-L-Ala-NH2 3 ................................ 46 Figure 2.19 Elaboration of 1 into GlcNAc-1,6-anhydro-MurNAc-L-Ala-iso-D-Glu 4........................ 47 Figure 2.20 Dr. Soni’s initial meso-oxa-Dap pentapeptide synthesis .................................................. 48 Figure 2.21 Dr. Soni’s final meso-oxa-Dap pentapeptide synthesis .................................................... 49 Figure 2.22 Comparison of yields between our meso-oxa-Dap pentapeptide synthesis and other meso-Dap peptide syntheses ........................................................................................................ 50 Figure 2.23 Initial coupling of disaccharide 1 to meso-oxa-Dap pentapeptide P5 .............................. 51 Figure 2.24 Nucleophilic demethylation of disaccharide 18 and subsequent peptide coupling .......... 52 Figure 2.25 Protected disaccharide 28 coupling to P5 ......................................................................... 53 Figure 2.26 Protected disaccharide-L-Ala coupling to meso-oxa-Dap tetrapeptide P4 ....................... 54 Figure 2.27 Deprotection of disaccharide pentapeptide 31 .................................................................. 55 Figure 2.28 Summary of disaccharide meso-oxa-Dap pentapeptide chemical synthesis ..................... 56 xiii Figure 2.29 Isolating, purifying, and reconstituting integral membrane proteins ................................ 58 Figure 2.30 Verifying the identity of AmpG in micelles and in nanodiscs by SDS-PAGE................. 59 Figure 2.31 Simplified illustration of integral membrane protein crystallization ................................ 60 Figure 2.32 Current cryo-EM micrograph of AmpG in nanodiscs ...................................................... 62 Figure 2.33 Illustration of proposed AmpG reconstitution into a lipid planar bilayer and detection of substrate transport ............................................................................................................. 64 Figure 2.34 Simple illustration of an isothermal titration calorimetry (ITC) setup ............................. 65 Figure 2.35 Heat change upon addition of sample to protein solution................................................. 66 Figure 2.36 Calorimetry titration curves at different c-values ............................................................. 67 Figure 2.37 Our collaborators’ ITC results .......................................................................................... 70 Figure 2.38 ITC curve of D-xylose binding to XylE ............................................................................ 71 Figure 2.39 ITC curves of Co2+ binding to BbFPNΔC ........................................................................ 72 Figure 2.40 ITC curves of arginine and agmatine binding to AdiC ..................................................... 72 Figure 3.1 Structure of our proposed AmpG “plug,” 6 ........................................................................ 74 Figure 3.2 Literature models of β-barrel transmembrane proteins with plug domains ........................ 75 Figure 3.3 Structure of WT SecY from Escherichia coli with plug domain ........................................ 76 Figure 3.4 Literature examples of permease inhibition measured by planar lipid bilayer ................... 78 Figure 3.5 Synthesis of various 2,2-dihydroxymethylpropionate dendrons ........................................ 80 Figure 3.6 Synthesis of disaccharide-alanine protected with an acid-labile acetonide ........................ 81 Figure 3.7 Convergent synthesis of disaccharide-alanine linked to a G3 dendron .............................. 82 Figure 3.8 Competitive inhibition of AmpG by dendron “plug” prevents transport............................ 83 Figure 3.9 Proposed detection of AmpG inhibition by the dendron plug ............................................ 85 Figure 4.1 General procedure for the use of photoaffinity probes ....................................................... 87 Figure 4.2 The “big three” functional groups in photoaffinity studies ................................................ 89 xiv Figure 4.3 Diazirine reactivity ............................................................................................................. 90 Figure 4.4 General diazirine synthesis ................................................................................................. 90 Figure 4.5 Benzophenone reactivity and general synthesis ................................................................. 91 Figure 4.6 Azide reactivity ................................................................................................................... 92 Figure 4.7 General azide synthesis ....................................................................................................... 92 Figure 4.8 Alternative photofunctional moieties and their reactive intermediates............................... 93 Figure 4.9 Two literature-known small molecule diazirines ................................................................ 94 Figure 4.10 Examples of both the C-diaz tag and the N-diaz tag used in the literature....................... 95 Figure 4.11 Literature example of muramyl dipeptide (MDP) photoaffinity probes ........................... 97 Figure 4.12 Our proposed AmpG photoaffinity probe, 7 ..................................................................... 98 Figure 4.13 Our proposed azide-bearing fluorescent tag ..................................................................... 99 Figure 4.14 Photoaffinity experimental design .................................................................................. 101 Figure 4.15 Initial attempt to make C-diaz ........................................................................................ 102 Figure 4.16 Literature synthetic route for N-diaz .............................................................................. 103 Figure 4.17 Our first successful synthesis of a test diazirine ............................................................. 103 Figure 4.18 Successful synthesis of a test diazirine via the t-butyl hypochlorite reagent .................. 104 Figure 4.19 Synthesis of ketones poised for diazirine transformation ............................................... 105 Figure 4.20 Protecting pro-diazirine ketones as ethylene ketals ........................................................ 105 Figure 4.21 Boc groups are removed preferentially to ethylene ketal ............................................... 106 Figure 4.22 Final successful syntheses of pro-diazirine ketones ....................................................... 107 Figure 4.23 Synthesis of final diazirine tags via the t-butyl hypochlorite reagent ............................. 108 Figure 4.24 Final coupling reaction to convergently assemble probe 7 ............................................. 109 Figure 4.25 SDS-PAGE of initial AmpG photolabeling with Rayonet photochemical reactor ......... 112 Figure 4.26 SDS-PAGE of “frozen and thawed” micelle-AmpG photolabeling ............................... 115 xv Figure 4.27 SDS-PAGE of micelle-AmpG photolabeling via 350 nm Tungsten lamp and 10 µM of 7 .................................................................................................................................... 117 Figure 4.28 SDS-PAGE of micelle-AmpG photolabeling via 350 nm Tungsten lamp and 2 µM of 7 ...................................................................................................................................... 118 Figure 4.29 SDS-PAGE of nanodisc-AmpG photolabeling ............................................................... 119 Figure 5.1 Summary of key compounds 1-7 chemically synthesized in this work ............................ 122 Figure 5.2 Proposed improved disaccharide pentapeptide chemical synthesis .................................. 125 Figure 5.3 Synthesis and possible utility of butyryl-protected disaccharide-L-Ala 58 ...................... 126 Figure 5.4 Disaccharide tripeptides and disaccharide tetrapeptides as putative AmpG substrates .... 127 Figure 5.5 Structures of possible AmpG “plugs” with different dendron sizes ................................. 128 Figure 5.6 Alternative AmpG photoaffinity probes bearing at least meso-oxa-Dap tripeptides ........ 130 Figure 5.7 Establishing an in vitro proton gradient before photolabeling AmpG .............................. 132 Figure 5.8 Leveraging Vocadlo’s spheroplast assay to establish a proton gradient ........................... 133 Figure E1 Illustration of photoaffinity experimental setup ................................................................ 136 xvi List of Abbreviations Å Angstrom Aβ amyloid beta Abs absorption Ac acetyl AcChoR acetylcholine receptor ACT 2-aryl-5-carboxytetrazole AIBN azobisisobutyronitrile Ala alanine aq aqueous Arf6 ADP-ribosylation factor 6 ATP adenosine triphosphate BLM black lipid membrane Bn benzyl Boc tert-butoxycarbonyl BSA bovine serum albumin Bu butyl oC degrees Celcius C55-P undecaprenyl phosphate cal calories CbmCho carbamoylcholine xvii Cbz carboxybenzoyl CCCP carbonyl cyanide m-chlorophenylhydrazone CHAPS 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate Chiraphos bis(diphenylphosphino)butane CMC critical micelle concentration CRBN cereblon cryo-EM cryogenic electron microscopy CuAAC copper-catalysed azide–alkyne cycloaddition D deuterium Dap diaminopimelic acid DCC N,N'-dicyclohexylcarbodiimide DCM dichloromethane DCU N,N'-dicyclohexylurea DDM n-dodecyl-β-maltoside DET diethyltartrate DIBALH diisobutylaluminium hydride DIC N,N'-diisopropylcarbodiimide DIPEA diisopropylethylamine DMA N,N-dimethylacetamide DMAP 4-N,N-dimethylaminopyridine DPPC dipalmitoylphosphatidylcholine DMF N,N-dimethylformamide E. coli Escherichia coli xviii EDCI 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide EEDQ N-ethoxycarbonyl-2-ethoxy-1,2-dihydroquinoline eIF3i eukaryotic translation initiation factor 3 subunit i Em emission EM electron microscopy eq equivalent Eqn. equation ESI electrospray ionization Et ethyl Fmoc fluorenylmethyloxycarbonyl g grams GlcN glucosamine GlcNAc N-acetyl glucosamine Glu glutamic acid h hours hv light HATU hexafluorophosphate azabenzotriazole tetramethyl uronium hBK channel human large conductance voltage- and calcium-activated potassium channel HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) HOBt hydroxybenzotriazole HPLC high-performance liquid chromatography HRMS high resolution mass spectrometry xix HTS high throughput screening IGF-1R insulin-like growth factor 1 receptor ILS illudin S Im imidazole IMAC immobilized metal affinity chromatography ITC isothermal calorimetry J NMR coupling constant K. aerogenes Klebsiella aerogenes KD dissociation constant kDa kilo-Dalton KI inhibition constant KM Michaelis-Menten constant L litres LAH lithium aluminium hydride LC liquid chromatography LDA lithium diisipropylamide Len lenalidomide LT lytic transglycosylase Lys lysine M molar (moles per litre) MAG monoacylglycerol Me methyl min minutes xx Ms methanesulfonyl MS mass spectrometry -or- molecular sieves MSP membrane scaffold protein MST microscale thermophoresis MurNAc N-acetyl muramic acid MHz megahertz NADPH nicotinamide adenine dinucleotide phosphate NaHMDS sodium bis(trimethylsilyl)amide N. gonorrhoeae Neisseria gonorrhoeae NHS N-hydroxylsuccinimide nm nanometer NMM N-methylmorpholine NMR nuclear magnetic resonance spectroscopy Nod nucleotide-binding oligomerization domain NSAID nonsteroidal anti-inflammatory drug OPAL 1,3,4-oxadiazin-5(6H)-one [Ox] oxidation P. aeruginosa Pseudomonas aeruginosa pA picoamps PA phosphatidic acid PAB pseudolaric acid B PBP penicillin binding protein PCC pyridinium chlorochromate xxi Pd/C palladium on carbon PDC peptide-detergent complex -or- pyridinium dichromate PEG poly(ethylene glycol) PEP phosphoenolpyruvate PG peptidoglycan Ph phenyl Phth phthalimido p-NZ p‐nitrobenzyloxycarbonyl PPTS pyridinium hydro-p-toluenesulfonate PUFAs poly-unsaturated fatty acids PyBOP benzotriazol-1-yloxytripyrrolidinophosphonium hexafluorophosphate Pyr pyridine quant quantitative rt room temperature s seconds sat saturated SDS sodium n-dodecylsulfate SDS-PAGE sodium dodecyl-sulfate polyacrylamide gel electrophoresis SMALPs styrene-maleic acid lipid particles SulRho sulforhodamine TBAF tetrabutylammonium fluoride TBHP tert-butyl hydroperoxide xxii TBS tert-butyldimethylsilyl TCT tracheal cytotoxin TDS dimethylthexylsilyl Tf trifluoromethylsulfonyl TFA trifluoroacetic acid TLC thin layer chromatography TMS trimethylsilyl TMSE trimethylsilylethyl Troc 2,2,2-trichloroethoxycarbonyl Trt triphenylmethyl (trityl) Ts p-toluenesulfonyl THF tetrahydrofuran TOF time of flight UDP uridine diphosphate UV ultra-violet W Watts wt weight WT wild-type w/u workup w/v weight to volume w/w weight per weight xxiii Acknowledgements I thank my supervisor, Martin Tanner for the opportunity to work with challenging glycopeptides to develop my skills and intuition as a synthetic chemist; for his guidance as a chemist, enzymologist, and an editor; for tolerating my often lofty promises of high yields in little time as well as my numerous “generous” deadlines. I thank my collaborator, Helena Sverak, and her supervisor, Natalie Strynadka, for their tireless efforts and bracing optimism with an often-frustrating project. Perhaps we will all be famous for “solving” AmpG one day, but for now it seems that the protein is content to keep its secrets. I thank Marco Ciufolini, Raymond Anderson, and Hongbin Li for their feedback and interest during my comprehensive exam. Marco Ciufolini, in particular, I admire as a chemist, a professional, and a Renaissance man – he is the primary person responsible for my becoming a synthetic chemist. Despite my initial failings as an undergrad student under his tutelage, he evidently saw enough potential in my abilities to offer an undergrad research position. If he can have faith in my potential as a professional, then maybe I can too. I thank Maria Ezhova for her efforts and intervention in deconvoluting my project’s many difficult NMR problems. I thank the machinists in the Chem building mechanical shop for their generous services and interesting conversations. I thank Jose Nuñez for always finding me a way to get paid, despite my occasional lack of foresight. I thank my mother, father, and grandmother for their mentorship and support – if they didn’t believe in my potential as a person, I certainly wouldn’t. Indeed, they’ll make sure I do, or else… My mother, in particular, has endured me at my absolute lowest and insists that even the xxiv darkest night can fade into a better day. I suppose she’ll need to have enough conviction for the both of us. To my partial chagrin, I also thank her for the wonderful food, housing, and general familial environment: if I do ever dare hope for something, it is that I can one day compensate her and the rest of my family for their herculean investment of time, money, and effort into my upbringing. I do not think I could ever hope for better friends and colleagues. Whenever I needed help or advice, they offered without hesitation. I thank Walid Abdelmagid for his initiative, gregarious charm, and life advice. Su Wang for his honesty, persistence, and loyalty to his friends. I thank Ziwang Wei for his curiosity, willingness to learn, and cheerful sense of duty. I thank Arvind Soni for his trust, humour, and grounded perspective – as well as for his synthetic contribution to our shared projects. I thank Tanya Adak for her conscientiousness, warmth, and altruism. I thank Дмитро Воскобойник (Dmytro Voskoboinyk) for his courage, patience, and integrity. These are just some of the myriad of virtues my friends possess and I aspire to emulate them all as far as I am able. Furthermore, I yearn to keep them and to repay them in equal virtue. Meeting them and working with them has been an honour. I thank God for reason, language, music, and dogs – may He never resolve to withdraw His magnanimity. xxv Dedication I dedicate this thesis to my grandmother, who by the grace of God is still alive to see her grandson succeed her and become a Doctor (of Chemistry). Lord willing, may she yet partake in many more years of vicarious professional satisfaction. I further dedicate this document in memory of my beloved Kourou, a loyal companion and joy of my home for almost 13 long blissful years of love, care, and attention. We’ve shared our journey through life since before the beginning of my undergraduate studies and I wish I could have at least traversed through one last Christmas together. Alas, before His name-day or ours, God beckoned to him that our paths must separate – and it is ever in a dog’s nature to heed his Master’s call. He was the best friend I’ve yet had: “The faithful dog – why should I strive To speak his merits, while they live In every breast, and man's best friend Does often at his heels attend.” - C.S. Winkle It is said that the souls of righteous men are rewarded eternally in Elysium. But what of the souls of dogs? While they can certainly be described as loyal, dependable, and stalwart, it cannot truly be said that creatures as innocent as they are have even the vaguest concept of “vice”; much less of its remedy in “righteousness.” When the Almighty retrieves the gift of life they borrowed, where do the dogs go? In faith, I do not know where dogs go when they die – but wherever they go; that’s where I want to end up, too. xxvi Kourou 2010 - 20231 Chapter 1: Introduction 1.1 Peptidoglycan 1.1.1 Peptidoglycan Structure and Biosynthesis Peptidoglycan (PG) is a major component of bacterial cell walls and is a rigid polymer responsible for maintaining the shape and integrity of the organism. 1 Its molecular structure contains a vast interconnected array of polysaccharides comprised of the repeating N-acetylglucosamine-β-1,4’-N-acetylmuramic acid (GlcNAc-MurNAc) disaccharide unit (Figure 1.1). Attached to this polysaccharide “backbone” are oligopeptides known as “stem peptides” that are covalently bound through their N-termini to the carboxylic acid of each MurNAc residue. Prior to crosslinking, each of these stem peptides is comprised of either L-Ala-iso-D-Glu-L-Lys-D-Ala-D-Ala (common in Gram-positive bacteria) or L-Ala-iso-D-Glu-meso-Dap-D-Ala-D-Ala (common in Gram-negative bacteria), where “Dap” corresponds to diaminopimelic acid. Crosslinking of the otherwise linear PG polymers occurs via the stem peptides by the action of transpeptidases: the ε-amino group of a stem peptide L-Lys (sometimes with intervening L-Ala residues) or meso-Dap is attached to the internal D-Ala of another stem peptide with the displacement the terminal D-Ala. This crosslinking is vital to survival of bacteria as it provides the required rigidity to protect them from lysis induced by osmotic pressure. 2 Assembly of PG’s monomeric subunit begins in the bacterial cytosol (Figure 1.1). 3 4 UDP-GlcNAc is converted into UDP-MurNAc via the action of MurA, which condenses phosphoenolpyruvate (PEP) with the 3-OH position of GlcNAc, and MurB, which uses NADPH to stereospecifically reduce the resultant vinyl ether. The ATP-dependent ligases MurC-MurF then sequentially construct the full stem pentapeptide to give the key UDP-MurNAc pentapeptide 2 intermediate – also known as “Park’s Nucleotide”. Thereafter the glycopeptide loses its cytosolic solubility as it undergoes pyrophosphate exchange with inner membrane-incorporated undecaprenyl phosphate (C55-P or “bactoprenol”) and becomes the membrane-bound intermediate “Lipid I”. 5 The membrane-associated glycosyl transferase MurG then transfers GlcNAc from UDP-GlcNAc onto the 4-OH of Lipid I to give the key intermediate “Lipid II,” which carries the disaccharide component of PG. Flippases transport this phospholipid sugar to the exterior of the inner membrane (to the extracellular surface in the case of Gram-positive bacteria). Finally, periplasmic enzymes called penicillin binding proteins (PBPs) catalyse the polymerization (glycosyltransferase activity) and crosslinking (transpeptidation activity) that generates the final mesh-like structure of PG. 6 3 Figure 1.1 The structure and biosynthesis of peptidoglycan in Gram-negative bacteria. 4 1.1.2 Peptidoglycan Recycling and Antibiotics As a bacterium grows and reproduces, this same rigid PG polymer must necessarily be broken down to accommodate the organism’s changing shape without unnecessarily wasting material. Therefore, bacteria have developed a series of enzymes specializing in the breakdown and recycling of their own cell walls. The sugar framework can be cleaved either by lysozymes (almost always from phages or host eukaryotes) and N-acetyl-D-glucosaminidases, which hydrolyze glycosidic bonds to give free sugars, or by lytic transglycosidases, which cleave glycosidic bonds by employing the internal 6-OH groups of MurNAc sugars as nucleophiles and form 1,6-anhydro-MurNAc (Figure 1.2). 7 Various bacterial amidases and peptidases cleave the amide bonds of stem peptides – including the cross-linked sections that lend the cell wall its characteristic rigidity. Thereafter, fragments of disintegrated PG – usually monomeric disaccharides with stem peptides of varying lengths – are transported back into the cytosol where most of the material (an impressive 90% in Escherichia coli 8) is retained and recycled. Figure 1.2 Enzymes responsible for the cleavage of the peptidoglycan glycosidic bonds. 5 Medicinal antibiotics often target one or more of the enzymes responsible for the formation of the bacterial cell wall . 9 β-Lactam antibiotics 10 (Figure 1.3), for example, induce bacterial lysis by irreversibly binding to the various transpeptidases responsible for cross linking the PG layers. Likewise, glycopeptide antibiotics, 11 such as vancomycin (Figure 1.3), target these same transpeptidases – albeit indirectly by binding to the stem peptides and preventing their adherence to the transpeptidase active site. Antibiotic resistance acquired from decades of overprescription and low-dosage exposure has rendered strains of pathogenic bacteria insensitive to these same previously potent drugs. 12 As resistance to conventional drugs grows, much of the research on pathogenic bacteria is done with the aim of finding new antibiotic agents against either existing drug targets, or against entirely new targets that are essential to bacterial survival and/or pathogenicity. Figure 1.3 Various β-lactam antibiotics (β-lactam moieties highlighted) and a glycopeptide antibiotic. 6 1.1.3 Antibiotic Resistance and AmpC As β-lactams accumulate in the bacterial periplasm and irreversibly inhibit PBP’s and cell wall growth, the enzymes responsible for cell wall breakdown nevertheless remain unimpeded: the aforementioned lysozymes, N-acetyl-D-glucosaminidases, and lytic transglycosylases continue to produce their respective PG fragments. The net effect of the inhibition of PG formation without the inhibition of PG recycling is the accumulation of a vast pool of PG fragments within the bacterium. Of particular interest to our research is antibiotic resistance induced via the processing of the cell wall by lytic transglycosylases and the ensuing accumulation of GlcNAc-1,6-anhydro-MurNAc PG fragments (Figure 1.4). These fragments are transported via the permease AmpG from the periplasm to the cytoplasm. 13 Once in the cytosol, the glycopeptides are processed by the glycosidase NagZ, which splits GlcNAc from 1,6-anhydroMurNAc while leaving the stem peptides intact, 14 and subsequently by the amidase AmpD, which cleaves the stem peptide as well. The resultant 1,6-anhydro-MurNAc is eventually processed back into GlcNAc 15 (Figure 1.4). In certain β-lactam-resistant organisms, however, the products of NagZ play a key role in antibiotic resistance. These partially processed PG fragments act as inducers and bind to the repressor AmpR which results in the expression of the ampC gene. 16 17 18 AmpC's (ampicillin C β-lactamases) are broad spectrum -lactamases that recognize both cephalosporins and penicillins, and are responsible for the acquired antibiotic resistance of many pathogenic organisms 19 20 such as Pseudomonas aeruginosa 21 and Mycobacterium tuberculosis. 22 Once expressed and subsequently transported to the cytoplasm, AmpC’s primary role is the hydrolysis of β-lactam antibiotics; thereby re-establishing the equilibrium between cell-wall breakdown and cell-wall formation. Once the β-lactams are inactivated, the pool of backed-up peptidoglycan fragments can be elaborated into PG 7 subunits such as Lipid II and finally polymerized into new PG layers via the action of the PBP’s (Figure 1.4). Finally, AmpC expression is down-regulated as its repressor returns and the pathogenic organism resumes proper growth and reproduction. 16 Figure 1.4 PG recycling and induction pathway of AmpC by β-lactam antibiotics. 8 As AmpC renders many valuable -lactam antibiotics ineffective, molecules that either inhibit AmpC, or its induction pathway, could serve as effective antibiotics when administered in conjunction with the appropriate -lactam. Targeting AmpC directly for inhibition is impractical due to its demonstrated propensity towards acquiring mutational resistance. 23 24 Targeting cytosolic enzymes implicated in the biochemical signaling pathway for AmpC expression (i.e. NagZ) is also problematic due to the relative impermeability of the bacterial inner membrane. In contrast, the periplasm is more readily accessible to small molecules. Thus, the possibility of “clogging” and/or “disrupting” the function of the aforementioned membrane-spanning permease AmpG, represents an attractive target for the development of novel antibiotics. In this thesis we aim to study the structure, specificity, and inhibition of AmpG. This is a highly collaborative project undertaken with the Strynadka laboratory in the Department of Biochemistry at UBC. Our role in this project is to provide substrates and chemical tools to study AmpG, and to carry out photoaffinity labeling studies. The Strynadka lab focuses on the purification, structural elucidation, and biophysical characterization of AmpG. 9 1.2 AmpG 1.2.1 History and Characterization AmpG is a membrane-spanning integral membrane protein known since the 90’s to be implicated in, and necessary for, the AmpC induction pathway. 25 Upon sequencing the ampG of E. coli, the prevalence of long stretches of hydrophobic residues indicated that the protein was likely a ~53 kDa, 491 amino acid, membrane-spanning permease with 10 membrane-spanning regions; 26 this same study undertaken by the Lindquist group proposed AmpG transported a signal molecule responsible for AmpC induction from the periplasm to the cytosol. Indeed, shortly thereafter, AmpG was identified as a member of the major facilitator superfamily, 27 28 a broad category of membrane-spanning integral membrane proteins – ubiquitous to both prokaryotes and eukaryotes – composed of ~12 parallel hydrophobic α-helices which passively transport small molecules through a lipid bilayer via a “rocker-switch” mechanism 28 (Figure 1.5). When inactive mutants of ampG are transformed into bacteria, the organism begins to “leak” significant quantities of GlcNAc-1,6-anhydro-MurNAc-peptide into extracellular space. 29 30 31 By this observation, the identity of the transported signal molecules was deduced. 30 10 Figure 1.5 The “rocker-switch” mechanism of passive transport for a major facilitator superfamily protein. The tertiary structure of E. coli’s AmpG was further refined in the early 2000’s through a series of β-lactamase fusion experiments (Figure 1.6). 27 Computational analysis of sequence hydrophobicity identified 14 (not only 10) hydrophobic segments as tentative membrane-spanning α-helixes. Interposing a β-lactamase gene along various sites in ampG – most fusion sites were placed near proline turns, although some were inserted directly within particular hydrophobic α-helixes – allowed the inference of AmpG’s membrane-spanning topology: the chimeric AmpG would only grant ampicillin resistance to the organism if its β-lactamase domain were found on the periplasmic side of the inner membrane. Through many of these insertion mutations, 10 of the 14 hydrophobic helixes were verified as membrane-spanning (8 of these 10 concurred with Lindquist’s preliminary model); the remaining four segments were situated on the cytosolic side of the permease and likely contributed in some way to the “rocker-switch” passage mechanism (Figure 1.5) – as seen in other major facilitator superfamily proteins. 32 33 34 11 Figure 1.6 Membrane topology diagram of E. coli AmpG determined by β-lactamase fusion experiments (recreated from literature). 27 An analysis of the possible substrates of AmpG was undertaken by the Park group wherein the passage of various tritium-labeled sugars, peptides, and glycopeptides derived from peptidoglycan was observed. 13 The permeability of the bacterial outer membrane was increased by a freeze-pump-thaw method to encourage radio-analyte uptake – moreover, a bacterial strain lacking AmpG function was used as a negative control. The results of Park’s radiolabel uptake experiment would conclusively demonstrate AmpG’s preference towards anhydro-PG fragments: GlcNAc-1,6-anhydro-MurNAc fragments with peptides of lengths ranging from 3-5 amino acids were transported in seemingly equal measure (Figure 1.7). Surprisingly, the simple disaccharide Glc-NAc-1,6-anhydro-MurNac (no stem peptide) was most readily accepted by the membrane protein. This seminal work also provided a tentative apparent Michaelis-Menten constant (KM,app) of 100µM for the GlcNAc-1,6-anhydro-MurNAc-tripeptide. The addition of the oxidative phosphorylation inhibitor and protonophore, carbonyl cyanide m-chlorophenylhydrazone (CCCP), seemingly reduced AmpG’s activity; supporting a model of AmpG functionality dependent on proton motive force, whereby a proton gradient across a given membrane drives the shuttling of both substrates and protons from compartments of a higher proton concentration to compartments 12 of a lower proton concentration – a H+-coupled “symport” mechanism reminiscent of another bacterial sugar permease, LacY (Figure 1.8). 33 35 Conversely, the AmpG knockout mutants did not readily accumulate radioactive anhydro-PG fragments within their cytosols in all circumstances; and were therefore far less affected by CCCP. Accordingly, Park suggested that AmpG’s susceptibility to disruption by CCCP could be leveraged to re-establish β-lactam sensitivity; if anhydro-PG fragments cannot accumulate in the cytosol, AmpC production cannot be induced. 36 Figure 1.7 Preference of E. coli AmpG uptake of PG fragments into bacteria: the TP74 ampG strain lacks AmpG function, the TP73 ampG+ strain has its AmpG restored via a plasmid, and carbonyl cyanide m-chlorophenylhydrazone is CCCP (a protonophore). Substrates denoted with “*” are labelled with tritiated DAP instead of tritiated GlcN. 13 13 Figure 1.8 H+-symport “rocker-switch” mechanism for a major facilitator. 1.2.2 Structural Modelling It should be noted that AmpG shows sequence and structural diversity between bacterial genera (differences in size and transmembrane α-helix numbers) and that structural models of E. coli AmpG are not cleanly superimposable on those of P. aeruginosa AmpG. In 2016, the membrane topology of P. aeruginosa AmpG was computationally modelled, and in addition, a 3-dimensional structure was constructed using the known crystal structures of other members of the major facilitator superfamily as basis sets. 37 The two models were not in accordance: the topology model predicted 14 transmembrane α-helixes (Figure 1.9), whereas the tertiary structure model predicted only 12 (Figure 1.10). Furthermore, a comparison of P. aeruginosa AmpG to that of 133 other strains of bacteria yielded certain clear sequence homologies, and four conserved amino acids were selected for site-directed mutagenesis. In particular, one of these four amino acids, A129 (thought to be embedded within the inner membrane), cannot be mutated into a more polar 14 residue (glutamate or threonine) without disrupting the function of AmpG (Figure 1.9), thereby making the bacterial mutant sensitive to ampicillin. This loss of function from the change of a single amino acid is hypothesized to proceed by interfering either with the binding of substrate to a currently unknown recognition site or with the conformational change resultant from that same recognition site receiving substrate. Figure 1.9 Modelled P. aeruginosa AmpG membrane topology recreated from literature 37 with key residue A129 highlighted. Figure 1.10 Modelled P. aeruginosa AmpG tertiary structure taken from literature. 37 (A) is the view of AmpG from the side; (B) is the view of AmpG from the periplasm. 15 Soon thereafter, other 3-D structural models were devised for AmpG’s of different bacterial strains (Figures 1.11 and 1.12); 38 39 likewise using crystallographic data of similar integral membrane proteins from databanks as basis sets. These contemporary models provide reasonable approximations of the AmpG structure; nevertheless, the substrate recognition domains/residues and their mechanism of interaction remain elusive. Subsequent attempts at targeting key conserved amino acids in the AmpG of Neisseria gonorrhoeae with point mutations succeeded in finding candidates for substrate recognition and/or protein rearrangement. 38 Without an actual crystal structure with bound substrate, however, it is impossible to conclusively assign roles to any of these candidates. Figure 1.11 3-D model of N. gonorrhoeae AmpG taken from literature. 38 Figure 1.12 3-D model of K. aerogenes AmpG taken from literature. 39 16 In summary, AmpG is an integral membrane protein and member of the major facilitator superfamily. Its role is the passive transport of GlcNAc-1,6-anhydro-MurNAc PG fragments from the periplasm into the cytosol via a “rocker switch” mechanism (Figure 1.5), possibly with the symport of H+ (Figure 1.8). It is an essential intermediary for both PG recycling and AmpC induction (Figure 1.4). Indeed, AmpG is produced in most strains of Gram-negative bacteria 37 and organisms subject to AmpG knockout release a surfeit of PG fragments to extracellular medium 29 31 40 (ablating approximately 40% of their cell wall each generation) 29 and lose prior β-lactam resistance. 29 37 41 These characteristics coupled with its ease of access by small molecules from the periplasm make AmpG an attractive possible target for drug development. 37 39 42 However, despite the interest in AmpG, almost nothing is confirmed about its structure – much less its “lock-and-key” recognition domain responsible for permitting the passage of PG fragments from the periplasm to the cytoplasm. Transmembrane peptide domains from different bacterial genera have been identified 36 or modeled 37 38 39 and compiled into reasonable tertiary structures. Nevertheless, a crystal structure of AmpG, either with or without bound substrate, has never been produced. Experiments have demonstrated AmpG’s “promiscuity” towards transporting PG fragments with stem peptide lengths ranging from 3-5 amino acid units; evidently, only the GlcNAc-1,6-anhydro-MurNAc subunit is strictly necessary for recognition by the permease. 13 In theory, AmpG might recognize and transport PG fragments with stem peptides of 1-2 amino acids or even with stem peptides bearing “unorthodox” amino acids. Without first understanding AmpG’s true structure and recognition domain, attempts to rationally design inhibitors – or even to construct a pro-drug that might infiltrate the cytosol by commandeering AmpG – are limited. 17 1.2.3 Contemporary Assays Prior to the work described in this thesis, AmpG has never been successfully purified and therefore no assays with purified protein have ever been performed. Instead, all studies to date have involved either intact bacteria or spheroplasts which lack the outer membrane. One such assay has been developed by the Vocadlo group and involves the preparation of E. coli spheroplasts both for ease of access to select fluorescent AmpG “probes” and for a clearer corresponding fluorescent signal. (Figure 1.13). 43 These probes are comprised of GlcNAc-1,6-anhydro-MurNAc with an ethylene diamine linker appended to a small fluorescent tag in the place of a stem peptide. The choice of fluorescent tags is apparently critical to the success of the fluorescent assay: the wavelength of light for the tag’s absorption maximum must freely penetrate the spheroplasts without damaging any proteins and the tag’s fluorescence emission maximum must be convenient to observe, with little or no signal lost to the cytoplasmic environment. Furthermore, for reasons still unclear, only certain fluorescent molecules are accommodated by AmpG, despite the apparent promiscuity of the permease towards various stem peptides. Disaccharides with the (notably expensive) Alexa Fluor 350 or BODIPYTM tags successfully infiltrate the spheroplasts, whereas disaccharides with the (comparatively cheap) 7-hydroxycoumarin tags do not 44 (Figure 1.14). This fluorescent assay was tested against simple GlcNAc-1,6-anhydro-MurNAc, GlcNAc, and 1,6-anhydro-MurNAc as “competitive inhibitors” to the Alexa Fluor probe. As expected, only the disaccharide competed with uptake of the probe by E. coli spheroplasts and apparent kinetic constants were established: KM,app = 340 µM for the Alexa Fluor probe and Ki,app = 70 µM for GlcNAc-1,6-anhydro-MurNAc – similar values to those found in Park’s radioassay.13 18 Figure 1.13 Summary of Vocadlo’s fluorescence assay. 43 19 Attempts by the Vocadlo group to devise “masked” fluorescent probes that only fluoresce when exposed to the cellular environment – i.e. 7-hydroxy/7-aminocoumarin with its phenolate/aniline moiety covered by either a fragmenting ester to be processed by a generic esterase or a small peptide to be processed by a generic peptidase – were unsuccessful; the substrates themselves seemed to be rejected by AmpG (Figure 1.14). 44 The tentative working hypothesis proposed by Vocadlo is that AmpG probes of the sort devised by his group must remain zwitterionic for acceptance by AmpG: a hypothesis supported by AmpG’s passage of disaccharides with larger stem peptides (i.e. L-Ala-iso-D-Glu-meso-Dap-D-Ala-D-Ala) but nonetheless challenged by the preferential passage of simple GlcNAc-1,6-anhydro-MurNAc. Figure 1.14 The results obtained with Vocadlo’s various AmpG probes – some of which fluoresce only after enzymatic processing. A red “X” indicates a probe that is not accepted by AmpG. 44 20 More recently, a high-throughput screening assay for AmpG inhibitors has been published. 45 The highly-drug resistant P. aeruginosa strain PAO1, wherein AmpC is overexpressed in the presence of β-lactam antibiotics, was tested against the ~645,000 compounds from the Scripps Diverse Drug Library for cell viability via optical density measurements (Figure 1.15, A). Hits on the screening assay were then redone in triplicate and counter-screened in the absence of the antibiotic cefoxitin (Figure 1.3); hits on the counter-screen were eliminated as general cytotoxins with no dependence on the AmpC induction pathway. True potential AmpG inhibitors were further refined by a concentration-response assay meant to elicit an IC50 with a sufficiently steep “drug-like” response curve. Initially, an optical density assay was attempted, but it yielded too few hits. Therefore, the authors developed a luciferase reporter assay: a PAO1 strain was developed with a lux (luciferase) fused to ampC (PAO1ampC-lux) for these assays and each instance of AmpC expression would be accompanied by luciferase expression – the amount of AmpG activity was coupled to the fluorescent signal of a luciferase assay. This increasingly sensitive series of screening procedures produced 8 total hits (Figure 1.15, B), although none of the compounds were particularly potent; nor did the hits tested meet the threshold to be considered “synergistic” with cefoxitin. Their work nonetheless produces a critical foundation for AmpG drug screening. HTS assays may be refined, and promising drug candidates may be derivatized for more conclusive results in the future. 21 Figure 1.15 A) Summary of the HTS assay methodology for finding AmpG inhibitors. B) The selected best putative AmpG inhibitors. 45 22 1.3 Thesis Goals Vocadlo’s continuous fluorescent spheroplast assay has demonstrated sensitivity towards competitive inhibition; probe uptake is significantly impeded by the presence of GlcNAc-1,6-anhydro-MurNAc. In future studies, this same assay can be used to gauge AmpG’s affinity for any number of small molecules: PG anhydro-MurNAc-fragments with stem peptides of varying lengths and sequences, other fluorescent probes or GlcNAc-1,6-anhydro-MurNAc derivatives, or even serendipitous inhibitors of AmpG discovered by high throughput screening. While tremendously useful, its application necessitates the expertise and materials for turning intact bacteria into spheroplasts since the assay fluorophores will not pass through the outer membrane of the Gram-negative organisms studied. Moreover, the spheroplasts themselves are extremely fragile; thereby leading to falsely deflated signal if the assay is undertaken with insufficient care. Therefore, there is still a great opportunity to improve upon current knowledge of AmpG structure and functional assessment. In this thesis we report the syntheses of anhydro-PG fragments to yield substrates, inhibitors, and photoaffinity probes for AmpG. This work was done in collaboration with the UBC Biochemistry department’s Strynadka lab – experts in protein expression, protein crystallization, and cryo-electron microscopy – to test these compounds on recombinant purified AmpG for the first time. In chapter 2 we outline our syntheses of substrates 1 - 5 which consist of the GlcNAc-1,6-anhydro-MurNAc motif with or without stem peptides of varying lengths (Figure 1.16). Compound 1 is the disaccharide alone whereas compounds 2 and 3 bear alanine and C-amido alanine, respectively. Compound 4 further adds D-glutamate to the stem peptide. Compound 5 mimics the full disaccharide pentapeptide in which, for ease of synthesis, an isosteric meso-oxa-23 Dap replaces the meso-Dap found in natural PG fragments. These compounds were used to determine the binding specificity of AmpG via isothermal calorimetry experiments (ITC) (vide infra). They could also be used to analyze the relative transport kinetics by utilizing AmpG that has been reconstituted in a planar lipid bilayer (vide infra) and/or to identify AmpG’s binding pocket via co-crystallization experiments. In Chapter 3 we describe the synthesis of a putative inhibitor to the permease, compound 6 (Figure 1.16). This compound bears the disaccharide crucial for recognition attached to a bulky dendron via a flexible, sterically non-bulky linker. We envision that AmpG could still bind to the disaccharide and that the dendron portion could act as a “plug” that would prevent passage through the protein channel. The efficacy of inhibitor 6 could be measured either in a lipid bilayer experiment or by using Vocadlo’s spheroplast assay. In Chapter 4, the synthesis and testing of the photoaffinity label, compound 7, is described. This compound contains the disaccharide-alanine unit that is important for recognition by AmpG. A diazirine warhead is attached to the alanine moiety that should generate a reactive carbene when exposed to light. This carbene could react with residues in the active site of the protein to covalently attach the molecule to the protein. Finally, compound 7 bears a terminal alkyne that will allow for fluorescent labeling of the modified protein using click chemistry. The goal is to use compound 7 to label residues in the AmpG recognition site and identify them using a subsequent trypsin digest and mass spectrometry analysis of the resulting peptides. In Chapter 5, an overview of the thesis is presented along future research directions. 24 Figure 1.16 AmpG substrates, inhibitors, and photoaffinity probes prepared in this thesis. 25 Chapter 2: Synthesis and Testing of AmpG Substrates 2.1 Anhydro-PG Fragments 2.1.1 Uses of Anyhdro-PG Fragments in studying PG-Binding Enzymes and Proteins Peptidoglycan fragments and their derivatives are used to optimize assays, attain crystal structures, and design potential enzyme inhibitors. Classic methods of attaining various PG fragments and intermediates involve the breakdown and processing of bacterial sacculi and lysates. 46 Although possible in labs with sufficient expertise, this method of attaining substrates is unideal for various reasons: the PG fragments are difficult to purify by HPLC as they are necessarily formed as a complex mixture, 47 48 post-purification derivatization is limited – especially via synthetic chemical routes – and harvesting larger quantities of a particular PG fragment or biosynthetic intermediate sufficient for a desired experiment is often not possible from bacterial lysates. Therefore, to meet the demand for various intermediates and substructures involved in PG assembly and breakdown, chemical synthesis is a useful alternative to bacterial processing. As mentioned previously, peptidoglycan fragments possessing a 1,6-anhydro-MurNAc (Figure 2.1) at their termini are the result of the cell wall being processed by non-hydrolytic enzymes called lytic transglycosylases (abbreviated as “LT’s”). In Gram-negative bacteria, they are found within the periplasm, and serve to break down the cell wall as the growing and reproducing organism requires. 7 26 Figure 2.1 Examples of previously synthesized 1,6-anhydro-PG fragments. 49 Research by the Mobashery group has identified up to 11 different LT’s in a Gram-negative Pseudomonas aeruginosa strain, 49 and shown that the function of at least some LT genes are essential for bacterial survival. 50 Turnover efficiency, substrate specificity, 49 51 and crystal structures of some of these LT’s were attained by employing chemically synthesized PG and anhydro-PG fragments (Figure 2.2). 52 53 54 Likewise, Mobashery (and his collaborators) used similar anhydro-PG fragments to determine the crystal structure (Figure 2.2) 55 15 and later the substrate specificity of AmpD in both E. coli 56 57 and P. aeruginosa. 58 27 Figure 2.2 The lytic transglycosylase, SltB3, holoenzyme crystal structure complex with GlcNAc-1,6-anhydro-MurNAc pentapeptide (left); 53 the structure of the amidase, AmpD, in complex with 1,6-anhydro-MurNAc and a tripeptide (right). 55 Besides being ubiquitous to the study of proteins implicated in the AmpC induction pathway, anhydro-PG fragments and their derivatives are also useful as substrates for certain PBP’s 59 or as potentiators of the immunological receptors Nod1 and Nod2 48 – and the various examples of anhydro-PG utility given thus far certainly do not encompass a comprehensive list. 2.1.2 Meso-Dap In this Chapter, the synthesis of potential AmpG substrates are described. For any Gram-negative PG fragment bearing a peptide of greater than three amino acids it is necessary to introduce a meso-diaminopimelic acid (meso-Dap) residue. As one of our targets bears a pentapeptide chain, we needed a synthetic strategy to introduce meso-Dap (or an isosteroic homolog). In this section we will summarize the literature methods to introduce meso-Dap into 28 peptides, and in the subsequent section we will review work from our lab that employs the structural homolog meso-oxa-Dap. Meso-Dap is an unusual amino acid in that it bears a side chain containing both a carboxylate and a primary amino group (Figure 2.3). In bacteria, it serves as both a precursor to lysine, 60 and a nucleophile responsible for cross-linking one “sheet” of PG with another via the action of PBP’s (Figure 2.4). 2 6 The synthesis and incorporation of meso-Dap into peptides presents significant challenges to synthetic chemists: its specific stereoconfiguration must be controlled and each of the four carboxylate and amino groups must be orthogonally protected to ensure the (S)-stereocenter is linked to the PG stem peptide. Figure 2.3 Meso-diaminopimelic acid (meso-Dap) in the stem peptide of Gram-negative bacteria. 29 Figure 2.4 Trans-peptidation (cross-linking) reaction of PG layers via penicillin-binding proteins (PBP’s). The majority of anhydro-PG fragments that have been chemically synthesized possess meso-Dap stem peptides, likely because many of the Gram-negative bacteria being investigated – including the ubiquitous model organism Escherichia coli and the aforementioned Pseudomonas aeruginosa and Mycobacterium tuberculosis – utilize meso-Dap as their nucleophile in cell wall cross-linking. The high demand and deceptively simple structure of meso-Dap compelled many different groups to attempt their own rendition of a synthetic route. However, only some of these routes produce orthogonally functionalized meso-Dap such that it can be incorporated into a stem peptide spanning more than 3 amino acids. Any proposed synthetic route wherein meso-Dap would thereafter be incorporated in an oligopeptide chain would need its various functional groups to be “orthogonally protected”: the (R)-carboxylate/amine and (S)-carboxylate/amine would all need to possess protecting groups rendered labile under completely different reaction conditions. 30 Some notable meso-Dap chemical syntheses of meso-Dap are shown in Figure 2.5. Holcomb’s work produced orthogonally-protected meso-Dap I via a Horner-Wadsworth-Evans reaction and subsequent enantiospecific hydrogenation, 61 Boons’ synthesis of meso-Dap II relied on the Grubbs cross-metathesis between (S)-vinyl glycine and (R)-allyl glycine, 62 Fujimoto and Fukase’s meso-Dap III work established the carbon framework via the Julia-Kocienski olefination of homologated Garner’s aldehyde, 48 and Hernandez and Martin’s meso-Dap IV route opted for a duo of Wittig olefinations of aspartic semialdehyde V and a subsequent generation of a chiral azide VI via the ring opening of a Sharpless epoxidation intermediate with sodium azide (Figure 2.5). 63 Kubash and Schmidt were the first to incorporate a meso-Dap stem peptide – the synthesis of which closely followed Holcomb’s methodology – onto MurNAc VII (Figure 2.6). 64 Fujimoto and Fukase were the first to incorporate a meso-Dap peptide into a “true” anhydro-PG fragment – more specifically, the Tracheal CytoToxin (TCT) 65 – with both GlcNAc and 1,6-anhydro-MurNAc (Figure 2.6). 48 Hernandez and Martin’s route would be adapted and modified by the Mobashery group to form the basis of their meso-Dap stem peptide syntheses (Figure 2.6) 66 – this route was employed in the only reported synthesis of the GlcNAc-1,6-anhydro-MurNAc pentapeptide VIII to date. 31 Figure 2.5 Chemical syntheses of meso-Dap from literature. 48 61 62 63 32 Figures 2.6 Previous chemical syntheses of meso-Dap glycopeptides. 48 64 66 33 2.1.3 Meso-oxa-Dap Despite the decades of innovation, the synthesis of this simple molecule still necessitates multiple steps with stereochemical control and expensive reagents. Circumventing these issues would greatly improve the accessibility of meso-Dap to interested researchers. To this end, the Vederas group developed an isosteric meso-Dap analogue, meso-oxa-Dap IX, derived from a formal substitution reaction between D-serine and L-serine (Figure 2.7). 67 O-Mesylation of N-trityl-D-serine results in formation of an aziridine. Exchange of the trityl group with a carbamate primes the aziridine X for nucleophilic opening with the hydroxyl group of an appropriately protected L-serine XI in the presence of a Lewis acid. This methodology is a succinct and potentially promising new way to prepare “meso-Dap” for various oligopeptides; especially those of PG fragments. We believe that the syntheses of these fragments can be shortened – and their overall yields increased – via the substitution of bone fide meso-Dap with the meso-oxa-Dap analogue. Figure 2.7 Vederas’ chemical synthesis of meso-oxa-Dap. 67 34 The Tanner group’s first publication in this area centered around the production of PG fragments containing meso-oxa-Dap. 68 These compounds were used as small molecule substrates for PG-modifying proteases. Dr. Soni in our group successfully employed a modification of Vederas’ methodology in which the aziridine was embedded within a tripeptide XII instead of a simple serine derivative (Figure 2.8). This ring opening reaction was accomplished with BF3 catalyst, which gave a modest yield (30%) of the desired meso-oxa-Dap containing tripeptide XIII. This modest yield was primarily due to the formation of an oxazoline by-product that is formed from an intramolecular Heine rearrangement 69 70 Despite the modest yield of the aziridine ring opening reaction, this methodology provided rapid access to PG fragments bearing meso-oxa-Dap in place of meso-Dap, and will be applied to the preparation of potential AmpG substrates in this Chapter. Figure 2.8 Dr. Soni’s meso-oxa-Dap tripeptide synthesis. 68 35 2.1.4 Chapter 2 Project Goals The goals in this chapter were to examine the specificity of purified AmpG with a variety of potential substrates. We would undertake the chemical synthesis of a panel of potential substrates that either lacked a peptide side chain or bore side chains of varying length. Our collaborators in the Strynadka lab would undertake the overproduction and purification of recombinant AmpG (this protein had never been purified at that time), as well as the biophysical characterization of the binding and transport events. As initial substrate targets, we chose the simple disaccharide, GlcNAc-1,6-anhydro-MurNAc 1, as the precursor to the rest of our four glycopeptides: GlcNAc-1,6-anhydro-MurNAc-L-Ala 2, GlcNAc-1,6-anhydro-MurNAc-L-Ala-NH2 3, GlcNAc-1,6-anhydro-MurNAc-L-Ala-iso-D-Glu 4, and GlcNAc-1,6-anhydro-MurNAc-L-Ala-D-iso-Glu-meso-oxa-Dap-D-Ala-D-Ala 5 (Figure 2.9). In vivo, AmpG transports disaccharides with stem peptides ranging from three to five amino acids, 38 41 but Park’s work demonstrated not only a willingness but an apparent preference for AmpG transporting the simple disaccharide substrate. 13 Substrate 1 also had demonstrable utility in Vocadlo’s assay; 43 therefore, it is an obvious synthetic target for us that is also the easiest to produce. Although, to the best of our knowledge, no group has attempted to measure AmpG’s capacity to accommodate disaccharides with smaller peptides, we suspect that it should recognize substrates 2, 3, and 4 without issues due to the membrane protein’s inherent promiscuity. This trio of synthetic targets which seem trivial to derive from 1. In contrast, the final compound 5 with its meso-oxa-Dap pentapeptide presents a challenging synthetic target. We possesses both the material and the expertise to incorporate such unnatural amino acids into a larger peptide framework for a convergent glycopeptide synthesis. 68 71 72 We surmise that this meso-oxa-Dap glycopeptide should be an excellent mimic of the disaccharide pentapeptide routinely recognized 36 by AmpG in vivo – a natural substrate analogue with a less cumbersome synthetic route owing to it lacking a true meso-Dap. Figure 2.9 AmpG substrates as synthetic targets. 2.2 The Synthesis of Potential AmpG Substrates 2.2.1 Synthesis of the Glycosyl Acceptor The synthesis of the GlcNAc-1,6-anhydro-MurNAc disaccharide initially followed the route published by the Vocadlo group, 43 which in turn followed the procedure established by the Paulsen group. 73 Specifically, the synthesis of a 1,6-anhydromuramate molecule (Figure 2.10) directly followed literature. Starting from D-glucal, iodolevoglucosan 8 was synthesized via the action of bis(tributyltin)oxide and iodine. We found this reaction to be more tolerant of undistilled “wet” solvent than literature would suggest. 74 75 This anhydrosugar was then elaborated to azidolevoglucosan 9 with sodium azide in refluxing H2O/DMF containing suspended NaHCO3. 37 The retention of configuration at C-2 suggests that this reaction proceeds via an epoxide intermediate (double displacement) with nucleophilic attack preferentially occurring at C-2 to give a 1,2-diaxial product due to the Platt-Fürstner rule. 76 77 Following treatment with stoichiometric amounts of TBS-Cl, the resultant mixture of constitutional isomers was treated with BaO/Ba(OH)2•8H2O and simultaneously alkylated with benzyl bromide to coalesce into a single product, 10. This can be explained by the ability of the TBS group to migrate between the C-3 and C-4 hydroxyl groups with a less sterically hindered alkylation at C-4. 78 After deprotection of the 3-O-TBS group with TBAF to give 11, alkylation with freshly made methyl 2-(S)-trifluoromethanesulfonylpropionate 79 was undertaken as described in the literature. 48 73 The ensuing anhydrosugar, 12, was hydrogenated and acetylated in one pot to yield methyl 1,6-anhydromuramate 13 – the more complex half of the glycan structure essential to our research. The desired disaccharide framework of our AmpG substrates would be made via a glycosylation reaction and 13 would be a reliable and ubiquitous glycosyl acceptor for our research. 38 Figure 2.10 Chemical synthesis of the 1,6-anhydro-MurNAc methyl ester 13. 2.2.2 Glycosylation of 1,6 Anhydro-MurNAc Methyl Ester We initially aspired to leverage Schmidt methodology for glycosylation. 80 We retained the N-phthalimido protecting group as per Paulsen’s paper 73 but sought to activate the anomeric position with a trichloroacetimidate (Figure 2.11). First attempts to convert simple glucosamine hydrochloride into the pre-glycosyl donor N-phthalimido-per-acetyl-glucosamine 14 were marred 39 by low yields. Much better results were obtained by reacting commercial tetra-acetyl glucosamine hydrochloride with phthalic anhydride in the presence of DMAP (Figure 2.11). With the phthalimido group in place, we then selectively deprotected the anomeric position with hydrazine acetate. Careful monitoring of the reaction progress was necessary to prevent the release of other acetyl groups. Overexposure of the sugar to this reagent results in global deprotection. Finally, the glycosyl trichloroacetimidate 15 was made at 0 ºC in as a ~9:1 ratio of β:α anomers via the action of powdered sodium hydride and trichloroacetonitrile electrophile. We attempted Schmidt glycosylation with either TMS-OTf or BF3. In both cases, the reaction was successful but low yielding. Eventually, we were able to optimize Schmidt glycosylation with BF3 to give yields of ~40% by increasing mole equivalents of the glycosyl donor to give disaccharide 16. Nevertheless, we were dissatisfied with these results and abandoned Schmidt methodology for our project. Figure 2.11 Preliminary glycosyl donor and glycosylation attempts. Methyl 1,6-anhydro-MurNAc would instead be glycosylated by an appropriate glycosyl bromide under Koenigs-Knorr conditions. 80 As Paulsen 73 and Vocadlo 43 had done, we treated peracetylated GlcNPhth with HBr in HOAc and acetic anhydride to produce the corresponding glycosyl bromide 17 as a ~3:1 ratio of α:β anomers (Figure 2.12). This glycosyl donor could be produced on a larger scale in fewer steps and with higher yields than its trichloroacetimidate 40 counterpart. With AgOTf and crushed dry 4Å molecular sieves, we successfully condensed 13 with the glycosyl bromide 17 to give the disaccharide 16 in good yields. In both glycosylation reactions, only a -configured product was obtained, presumably due to neighbouring group participation of the phthalimido group at C-2. Figure 2.12 Glycosyl bromide synthesis and subsequent Koenigs-Knorr glycosylation. However, subsequent attempts to deprotect the disaccharide phthalimide 16 with hydrazine and to then re-acetylate the resultant free amine produced a new diacetyl acyl hydrazide side-product, which was tremendously difficult to separate from the desired product (Figure 2.13). Occasionally batches of sugar treated with hydrazine hydrate would give good yields of 18 with very little or no side-product; however, we could not reliably reproduce this favorable outcome. Preliminary deacetylation with sodium methoxide had no effect on reaction yields or on side product formation. Only switching the hydrazine source to the less reactive hydrazine acetate somewhat attenuated the prevalence of the diacetyl acylhydrazide and afforded product in consistent – if moderate – yields (Figure 2.13). Nevertheless, several “bad batches” and unsuccessful flash chromatography purifications compelled us to seek an alternative glycosyl donor to forgo the use of hydrazines entirely. 41 Figure 2.13 Attempted deprotection of the phthalimide group of compound 16. We know from previous research done by Dr. Hadi in our lab 71 that the glycosyl bromide of peracetylated GlcNAc, while trivial to synthesize, is insufficiently reactive toward a sugar’s secondary alcohol during glycosylation: the glycosyl donor simply collapses into a stable oxazoline and is inert to further reaction (Figure 2.14). Fortunately, modifications which place electron withdrawing atoms on the acetamido group have been demonstrated to improve reactivity towards glycosyl acceptors. Previously, Dr. Hadi designed a trichloroacetyl-bearing glycosyl trichloroacetimidate that reacts successfully with regular MurNAc under mild Koenigs Knorr conditions. The trichloroacetyl group still undergoes neighbouring group participation to ensure that the -anomer is produced, but the more reactive trichloromethyl oxazoline is readily opened by the acceptor nucleophile. The trichloroacetyl group could subsequently be converted to the naturally occurring acetyl via the reducing agent tributyltin hydride and the radical initiator AIBN. 42 Figure 2.14 Dr. Hadi’s use of anchimeric assistance from a trichloromethyl group to prompt successful glycosylation. The glycosyl donor 20 inspired by Dr. Hadi’s glycosyl donor served our purpose exceptionally well – even increasing our glycosylation yield (Figure 2.15). Additionally, the protected trichloroacetylsugar 19 could be dependably produced in multigram quantities from GlcNH2•HCl instead of the more expensive tetra-acetyl GlcNH2•HCl, by treatment with trichloracetylchloride followed by acetic anhydride. Compound 19 could be converted into glycosyl bromide 20 as a 3:1 ratio of α:β anomers by treatment with HBr in HOAc. The subsequent coupling of 20 and 13 occurred in good yield by treatment with AgOTf. The appearance and subsequent disappearance of the trichloroacetyl oxazoline intermediate could easily be monitored by TLC if desired. The oxazoline could even be produced separately (absent coupling partner), isolated, and stored for later use if desired. We furthermore concluded that at least 2 days of reaction time are necessary for optimal glycosylation yields of 21 – a slow but consistent reaction. 43 Figure 2.15 The final glycosylation method used in the disaccharide synthesis. The foregoing radical dehalogenation 71 soon proved to be difficult to scale up, perhaps due to the difficulty of completely purging large reaction vessels of interfering oxygen. Furthermore, the use of freshly prepared Bu3SnH was essential for best results. Consequently, we opted to leverage a Raney nickel dehalogenation method based on previous literature (Figure 2.16). 81 Commercially available Raney nickel slurry is basic and would ordinarily rapidly saponify any esters in situ; however, it can conveniently be added to neat acetic acid with only minor and gradual expenditure of the reducing metal via hydrogenesis. The peracetylated disaccharide 21 can be quickly dehalogenated to give compound 18 under these non-basic conditions without any risk of saponification. Moreover, this experimental setup can be designed to suppress any unwanted Fischer-type deacetylation with the simple addition of ethyl acetate to act as a “buffer” against this side reaction. As an aside, we demonstrated that dehalogenation could also be achieved by palladium-catalyzed hydrogenolysis; however, this produced a mixture of partially dehalogenated products and we were not able to drive the reaction to completion. After successful dehalogenation of 21 in good yield, saponification quantitatively produced pure GlcNAc-1,6-anhydro-MurNAc, 1, to serve as the minimal AmpG substrate and to be used in coupling to stem peptides. 44 Figure 2.16 Dehalogenation and synthesis of GlcNAc-1,6-anhydro-MurNAc 1. 2.2.3 Peptide Coupling We then proceeded to append peptides to disaccharide 1. L-Ala benzyl ester could be coupled to the disaccharide fairly consistently (Figure 2.17); nevertheless, the yield of 22 was unusually low – especially when compared to those of more conventional peptide coupling reactions. Furthermore, successful purification of the glycopeptide could only be achieved via an initial silica plug and subsequent C-18 reverse-phase chromatography. We tested our conventional coupling reagents and methodology on the condensation between N-Boc-L-Ala and D-Glu-α,γ-(OBn)2•HCl. As expected, the yields were semi-quantitative (~95%) and purification was trivial. Therefore, it is specifically the creation of an amide bond between the lactate of 1 and a peptide coupling partner that is troublesome. Compound 22 was readily deprotected by hydrogenolysis to give our second putative AmpG substrate, compound 2. 45 Figure 2.17 Synthesis of GlcNAc-1,6-anhydro-MurNAc-L-Ala 2. Alternatively, 1 could be coupled to L-Ala benzyl ester, partially purified via a C-18 reverse-phase silica plug, and the crude eluate condensed with acetone under acidic conditions to give acetonide-protected disaccharide 23 (a key building block in later Chapters 3 and 4) in similar yields to the deprotected disaccharide 22 (Figure 2.18). 23 was then reacted with ammonia in methanol to give 24 and deprotected via Dowex to give our third putative substrate, compound 3. It should be noted that our attempt to produce 3 via the L-alanine(NHBn)-coupled disaccharide 25 failed due to the relative stability of 25’s amide benzyl to hydrogenation conditions: no debenzylation occurred even after a week of hydrogenation. 46 Figure 2.18 Elaboration of 1 into GlcNAc-1,6-anhydro-MurNAc-L-Ala-NH2 3. Our attempts to couple L-Ala-D-Glu dibenzyl ester to disaccharide 1 failed: the disaccharide dipeptide 4 could only be synthesized in a stepwise fashion (and in low yields) by first preparing compound 2, and then coupling it to D-Glu dibenzyl ester to give 26. Deprotection of compound 26 by hydrogenolysis gave our fourth putative AmpG substrate, compound 4 (Figure 2.19). 47 Figure 2.19 Elaboration of 1 into GlcNAc-1,6-anhydro-MurNAc-L-Ala-iso-D-Glu 4. We next wished to prepare the GlcNAc-1,6-anhydro-MurNAc pentapeptide, compound 5. As Dr. Soni in our lab had extensive experience in preparing meso-oxa-Dap containing peptides, he undertook the synthesis of the peptide portion of this molecule. His initial attempt involved embedding the aziridine within a pentapeptide and using the ring opening reaction to form meso-oxa-Dap as the final step (Figure 2.20). 72 While small quantities of the protected pentapeptide Fmoc-P5 were formed in this reaction, the yields were unacceptably low. He rationalized that the number of potential BF3 coordination sites were too great in a pentapeptide and the catalyst was likely non-productively binding to the substrate. 48 Figure 2.20 Dr. Soni’s initial meso-oxa-Dap pentapeptide synthesis. 72 Since the ring opening reaction had previously worked on a tripeptide susbstrate (Figure 2.8), 68 Dr. Soni next attempted the coupling on an aziridine embedded with a tripeptide XIV (Figure 2.21). This attempt was successful and gave comparable yields (33%) to the previous study. 68 Once again, 37% of the Heine rearrangement product was formed in this reaction. 68 69 70 From there, the rest of the protected pentapeptide Boc-P5 could be assembled by “capping” by an N-terminal D-Ala and a C-terminal L-Ala. 72 Treatment with HCl in dioxane would then finally liberate the N-terminus of the pentapeptide P5 for coupling of the lactate to the disaccharide. 49 Figure 2.21 Dr. Soni’s final meso-oxa-Dap pentapeptide synthesis. 72 Despite the modest yields of some individual steps within Dr. Soni’s pentapeptide synthesis, using meso-oxa-Dap greatly simplifies the synthesis and increases the overall yield of the pentapeptide. Compared to other contemporarily synthetic routes, 48 64 66 82 Dr. Soni’s produces functional meso-Dap in the fewest overall steps (Figure 2.22). Furthermore, our oxa-Dap peptides have been successfully used as a substrates by wild-type peptidoglycan processing enzymes; 68 therefore the isostere is readily accepted by biological catalysts. 50 Figure 2.22 Comparison of yields between our meso-oxa-Dap pentapeptide synthesis and other meso-Dap peptide syntheses in the literature. 48 64 66 82 We next attempted to couple our disaccharide 1 with Dr. Soni’s pentapeptide P5 (Figure 2.23). We dared not risk lowering the yields of this reaction with an initial extraction of what we expected may be a water-soluble product; consequently, purification via flash chromatography was tedious and inefficient, with coupling agent and/or additive routinely co-eluting with our product. Eventually, some final material 27 was successfully purified – however, the yield was only 28%. Although we are not entirely certain why this particular coupling reaction is inefficient 51 in comparison to conventional peptides, we suspect that the disaccharide’s steric bulk retards the formation of the peptide bond. Furthermore, the L-Ala-D-Glu motif that we employ may be problematic due to its propensity to spontaneously cyclize into a diketopiperazine in situ if not readily sequestered by a coupling partner (Figure 2.23). Previous groups in the field of peptidoglycan chemical synthesis attest to such a side reaction lowering yields. 83 84 85 Although we never found any direct evidence for such a side-product, it is reasonable to expect its formation may occur if coupling to our disaccharide were indeed unusually slow. The low yields and inelegant purification of this latest glycopeptide were deemed unacceptable; we therefore aspired to improve yields and workup efficiency via a slightly revised synthetic procedure. Figure 2.23 Initial coupling of disaccharide 1 to meso-oxa-Dap pentapeptide P5. We therefore resolved to decrease the polarity of the coupling product by retaining the acetyl groups masking the hydroxyls of the disaccharide post glycosylation. To this end, we 52 deprotected the methyl ester of 18 in a nucleophilic mode via the action 4-aminothiophenol under reflux and mildly basic conditions (Figure 2.24). 86 This selective demethylation formed our desired carboxylic acid 28, which could reliably undergo preliminary purification via solvent-solvent extraction without adversely affecting yields – a promising observation for the eventual workup of our desired protected glycopeptide. To support the utility of our new protected disaccharide, we again attempted to append both L-Ala benzyl ester and L-Ala-D-Glu dibenzyl ester onto its exposed carboxylate. To our chagrin, coupling yields were still low (19% and 17%, respectively); nevertheless, purification of 29 and 30 was indeed easier. Moreover, we successfully directly coupled the L-Ala-D-Glu dipeptide onto our disaccharide for the first time. Figure 2.24 Nucleophilic demethylation of disaccharide 18 and subsequent peptide coupling. We then undertook the coupling of compound 28 to P5 (Figure 2.25). Although acetylation of the polar hydroxyl groups made separation of the glycopeptide product from coupling reagents and additives possible via an extraction step – mass spectral analysis suggested that EDCI was forming an amphiphilic guanidinium side-product with the pentapeptide, which was difficult to separate via flash chromatography. Ultimately, dual rounds of flash chromatography gave pure 53 protected disaccharide pentapeptide 31 in 27% yield. Replacing EDCI with PyBOP may have ameliorated the issue of side product formation, but we were unable to optimize this coupling reaction due to our limited quantitiy of material. We did, however, try replacing the additive HOBt with NHS to perhaps monitor the reaction progress by the formation of the more stable NHS-ester. The results of this additive replacement were unremarkable: some NHS ester could indeed be seen by mass spectrometry, but always in minority to the starting carboxylate signal, and product yields were always lower (i.e. 14%) than those of the prior HOBt-dependent reactions. Figure 2.25 Protected disaccharide 28 coupling to P5 and proposed structure of guanidinium side-product. 54 To test whether the formation of a diketopiperazate side product was responsible for the low coupling yields, Dr. Soni provided us with a tetrapeptide P4 as this compound cannot cyclize (Figure 2.26). We first subjected compound 29 to hydrogenolysis to generate per-acetylated Glc-NAc-1,6-anhydro-MurNAc-L-Ala which was immediately coupled to the tetrapeptide. We opted for the NHS additive in this instance and although the reaction was successful, the yields of 31 were lower (24%) than those of our pentapeptide coupling reactions. Figure 2.26 Protected disaccharide-L-Ala coupling to meso-oxa-Dap tetrapeptide P4. Ultimately, a lack of material prevented us from optimizing the coupling reactions further and the direct coupling of P5 to compound 28 provided our highest yield (Figure 2.25). During the final global deprotection steps, glycopeptide 31 presented a parting challenge: overexposure to base results in the accumulation of an elimination side-product at the meso-oxa-Dap locus (Figure 2.27). This issue is well-documented when dealing with haloalanines, 87 cyclic sulfamidates, 88 and even cysteine disulfide bridges, 89 90 91 but it has not yet (to the best of our knowledge) been observed in the oxa-Dap series. Therefore, we found that it is necessary for the hydrogenolysis of the protecting groups to proceed before base treatment – and that the base be relatively mild – due to the tendency of the oxo-Dap eliminating dehydroalanine under basic conditions: evidently, 55 formation of a carboxylate sufficiently raises the pKa of the (R)-stereocenter to suppress deprotonation and subsequent elimination. Finally, we carefully quenched our carbonate base with incremental amounts of the heterogeneous acidic Dowex resin for ease of workup; only lowering the pH to ~4 to limit the glycopeptide product 5 binding to the resin. This provided working quantities of the fourth AmpG substrate to be used in binding and transport studies. Figure 2.27 Deprotection of disaccharide pentapeptide 31 – initial base treatment leads to buildup of elimination side-products. In conclusion, we synthesized five potential AmpG substrates. Notably, GlcNAc-1,6-anhydro-MurNAc bearing a meso-oxa-Dap-containing pentapeptide 5 was prepared in 12 steps starting from D-glucal with a ~1.6% overall yield (Figure 2.28). In a similar manner, GlcNAc-1,6-anhydro-MurNAc-L-Ala 2 was made in 11 steps with a ~6.1% overall yield, GlcNAc-1,6-anhydro-MurNAc-L-Ala-NH2 3 was made in 12 steps with a ~4.6% overall yield, and GlcNAc-1,6-anhydro-MurNAc-L-Ala-D-Glu 4 was made in 13 steps with a ~0.7% overall yield. Multimilligram quantities of these anhydro-PG fragments are excellent assets for our collaborators’ crystallographic, calorimetric, and/or electrophysiological experiments on purified recombinant AmpG. 56 Figure 2.28 Summary of disaccharide meso-oxa-Dap pentapeptide chemical synthesis. 57 2.3 Our Collaborators’ Work 2.3.1 Expressing and Isolating AmpG Extensive efforts have been made by our collaborators in the Strynadka group at UBC to express and isolate AmpG. Unsurprisingly, a large amphiphilic permease that requires a lipid bilayer environment for structural stability is an extremely challenging protein expression target. Moreover, overexpression of AmpG is apparently toxic to cells; thereby requiring larger volumes of bacterial broth to be processed. Furthermore, AmpG displays general instability at room temperature (and even at 0 ºC if left too long) and must be both prepared and expended “fresh”. Integral membrane proteins like AmpG are usually isolated and purified in detergent micelles, which approximate the native hydrophobic environment of the bacterial bilayer (Figure 2.29) 92 93 – these membrane proteins are not viable outside of a membrane-mimetic environment. The micelle detergent chosen may be ionic (i.e. SDS), zwitterionic (i.e. CHAPS), or non-ionic (i.e. Triton-X 100) and must be kept slightly above its critical micelle concentration (CMC) when sequestering membrane proteins: either too much or too little of the detergent will cause the protein to denature. 92 94 Once stabilized in this amphiphilic environment – a so-called “peptide-detergent complex” (PDC) – the integral membrane protein may be purified by chromatography and its micelle detergent may be gradually exchanged for other surfactants more suitable for the desired methodology. If desired, micelle-dissolved protein may be reconstituted into artificially-generated liposomes – thereby generating “proteoliposomes” (Figure 2.29) – via the gradual exchange of micellar detergent for surfactants such as phosphatidic acid (PA) or dipalmitoylphosphatidylcholine (DPPC), 95 which instead tend to form 2-D bilayers in solution. 93 58 96 These same bilayer-assembling phospholipids may also be mixed with either a so-called “membrane scaffold proteins” (MSPs) 97 or styrene-maleic acid lipid particles (SMALPs) 98 to generate lipid nanodiscs (Figure 2.29), solubilized 2-D circular “rafts” of bilayer lipids with uniform diameters. 93 99 Upon exposure of the protein-micelle complex to a mixture of nanodisc ingredients and subsequent removal of micelle detergent via dialysis or lipophilic resin treatment, membrane protein-nanodisc complexes may form spontaneously. These nanodiscs are known to most closely resemble the native environment of integral membrane proteins and may be isolated as structures of a homogenous and predictable size via affinity chromatography. 99 Unfortunately, the process of peptide reconstitution from micelle to nanodisc necessarily sacrifices some material; and therefore lowers the overall yield of protein preparation. Figure 2.29 Isolating, purifying, and reconstituting integral membrane proteins from transformed bacteria. Note that the strings of blue circles represent the recombinant protein hexahistidine tag. Members of the Strynadka group successfully isolated multimilligram samples of E. coli AmpG modified with an N-terminal thrombin-labile hexahistidine tag stabilized in n-dodecyl-β-maltoside (DDM) micelles. The recombinant protein was overproduced in E. coli and then purified via immobilized metal affinity chromatography (IMAC) followed by size exclusion 59 chromatography; protein size and structural integrity were verified by SDS-PAGE (Figure 2.30, left). They also successfully reconstituted AmpG into peptide MSP-nanodiscs; albeit only in multimicrogram amounts (Figure 2.30, right). However, neither AmpG’s in vitro functionality nor its tertiary structure has yet been demonstrated. Figure 2.30 Verifying the identity of AmpG in micelles (left) and in nanodiscs (right) by SDS-PAGE. 2.3.2 AmpG Crystallography These protein micelles could be leveraged to attempt protein crystallization. Crystallization of integral membrane proteins usually occurs via a “lipid cubic phase” (also known as “in meso” crystallization), 100 92 wherein particular bilayer-mimetic surfactants (i.e. monoacylglycerols or “MAGs”) bearing the reconstituted protein partition themselves into cubic 60 arrays of interlocking lipid bilayers (often in the Pn3m space group, Figure 2.31). 101 Once embedded in the laminar matrix, the integral membrane proteins, driven by various electrostatic interactions between each other, gradually coalesce into uniform crystal lattices within the amphiphilic medium. In theory, AmpG should be crystallizable in meso; moreover, as has been shown with other integral membrane proteins, 102 103 104 these crystal structures may be co-crystallized with any of our synthesized sugars to potentially elucidate key substrate recognition residues. As of now, only AmpG apoprotein “microcrystals” have been produced within the lipid cubic phase – these crystals are too small for structural determination. Further crystallographic results are pending – including the possibility of creating AmpG chimerae fused to protein domains which encourage crystallization. 105 There are, however, alternative methods of structural resolution that our collaborators fully intend to exploit. Figure 2.31 Simplified illustration of integral membrane protein crystallization from the lipid cubic phase. 61 2.3.3 AmpG Electron Microscopy The aforementioned peptide-nanodisc complexes have demonstrable utility for cryogenic electron microscopy (cryo-EM), 97 a type of scanning electron microscopy done at cryogenic temperatures to limit molecular vibration; thereby increasing structural resolution. 106 107 108 Cryo-EM can be thought of as a complement and/or alternative to protein crystallography: it is not necessary to grow perfect crystals or to have perfect structural uniformity, and it predominantly gives information about surface topology as its resolution is currently lesser than that of protein X-ray crystallography. Typically, cryo-EM resolves the structures of exceptionally large proteins, which are ordinarily tremendously difficult to crystallize, with asymmetric topologies (i.e. the protein is known to have different morphology on one “end” than on another). 97 There are numerous examples in literature 109 110 111 of integral membrane proteins bound as peptide nanodiscs whose structures are successfully resolved by cryo-EM. Our collaborators in the Strynadka lab are in fact renowned experts in this sub-field. Their initial cryo-EM studies on AmpG embedded within nanodiscs have provided the images shown in Figure 2.32. Although the 54 kDa AmpG is thought to be too small to be adequately analyzed by cryo-EM, there are methods that can alleviate this problem. Our collaborators are constructing an AmpG chimera with the BRIL structural motif – a domain used previously as a cryo-EM fiducial 112 – thereby both increasing protein mass and differentiating one “side” of AmpG for facile micrograph interpretation and structural elucidation (Figure 2.32). 62 Figure 2.32 Current cryo-EM micrograph of AmpG in nanodiscs and proposed AmpG-BRIL chimera to increase structural resolution. 2.3.4 Planar Lipid Bilayer Studies with AmpG Our collaborators also aim to demonstrate the activity of AmpG in vitro and to test its specificity towards our substrates with a “planar lipid bilayer” experiment. A planar lipid bilayer – also known as a “bilayer lipid membrane” or a “black lipid membrane” (BLM) – is an artificial lipid bilayer meant to imitate the properties of cell membranes in vitro and is spread across a small circular opening (50-100 µM wide) of a TeflonTM divider between two separate chambers (Figure 2.33). 113 114 One technique to form this artificial bilayer barrier between the two chambers is the so-called “painting” method established by Montal and Mueller. 115 Bilayer-generating surfactants are dissolved in an organic solvent (usually n-decane) and “painted” across the submerged opening between the chambers by means of a brush; the surfactant molecules within the applied thin-film spontaneously arrange to create a bilayer with polar ends facing the two aqueous compartments and hydrophobic ends facing the organic solvent spanning the divider hole. An electrical potential 63 can be applied across the two compartments both to verify the integrity of the nascent artificial bilayer “capacitor” and to measure dissolved ion passage as the basis for the “signal” detection in generic planar lipid bilayer experiments. When the two solvent chambers are segregated, their individual chemical/ionic environments can be modulated in accordance with experimental protocol. AmpG may then be reconstituted into this artificial membrane, either directly through the addition of its PDC, or indirectly via its reconstitution from proteoliposomes. 116 Once definitively embedded, AmpG’s substrate transport efficacy can be monitored under an electric potential by loading one of the bilayer chambers with a PG fragment and an excess of ions: as material is recognized by and passes through the permease, ions such as K+ and H+ are simultaneously transported across the gradient to produce measurable signals as spikes in current (pA in scale). These ion current traces effectively report on the rate of pore opening and can be used to compare the efficiency of different PG fragments and eventually to analyze experiments run with AmpG “plugs”: if these potential inhibitors successfully compete with PG fragments and block their passive transport, then there should be an obvious decrease in the amount of measured passage events. 117 Unfortunately, although our collaborators succeeded in producing a black lipid membrane, initial attempts to integrate AmpG via a liposome carrier were unsuccessful – further attempts are currently underway. 64 Figure 2.33 Illustration of proposed AmpG reconstitution into a lipid planar bilayer and detection of substrate transport. 65 2.3.5 Calorimetric Studies on AmpG A potential means of determining AmpG substrate binding affinity is isothermal titration calorimetry (ITC). 118 119 Micelle-solvated or nanodisc-embedded AmpG can be isolated in a chamber which measures and maintains a constant temperature (Figure 2.34). Substrate can be added in intervals to this stirring solution and the change in heat (ΔH) associated with substrate-protein binding events can be quantified (heater power output measured as µcal/s) 120 (Figure 2.35 left); binding events may be endothermic or exothermic, but a larger signal indicates tighter binding. Readings can be compared to a reference cell wherein identical detergent micelles/nanodiscs lacking AmpG are subject to identical interval additions of binding material to account for changes in heat due to mixing and/or substrate-detergent interactions. Figure 2.34 Simple illustration of an isothermal titration calorimetry (ITC) setup. 120 Integrated heat data may be represented as a graph of heat per mol of injectant (substrate) vs. molar ratio (substrate/protein), thereby yielding a more conventional-looking titration curve (Figure 2.35, right). If substrate binding is exotheric, this curve bends “bottom-up”; if substrate binding is endothermic, this curve bends “top-down”. Certain thermodynamic properties of the binding interaction can be gleaned from this plot: binding enthalpy from the height of the curve, 66 binding stoichiometry from the position of the inflection point, and (most critically) the dissociation constant (KD) from the slope of the curve at said inflection point (Figure 2.35, right) – with both the values of ΔG and ΔS inferred once both KD and ΔH are known via the elementary equation ΔG = -RTln(1/KD) = ΔH – TΔS (Eqn. 1). The formula describing the shape of this curve is called the “Wiseman Isotherm.” 121 122 In practice, the titration curve is analyzed by software that derives these key constants from a line of best fit to a theoretical Wiseman Isotherm. Eqn. 1: Figure 2.35 Heat change upon addition of sample to protein solution (left); corresponding integrated heat as molar ratio of substrate/protein (right) – this titration curve yields thermodynamic parameters. 123 An ideal titration curve should have a clear sigmoidal shape for ease of software processing and there is a key practical consideration to achieve this end. The value of the protein concentration relative to the KD value of the substrate is tremendously important to the formation 67 of a clear sigmoidal titration curve (Figure 2.36). This relationship is described by the “c-value”, which is defined as c = [P]N/KD (Eqn. 2) where “[P]” is the protein concentration in the cell and “N” is the number of substrate binding sites on the protein. 118 Empirically, the optimal c-value for deriving all three key thermodynamic constants (ΔH, stoichiometry, and KD) is c = 30; therefore all ITC experiments should, when possible, carry out the titration at a protein concentration that is 30 times greater than the substrate dissociation constant. Eqn. 2: Figure 2.36 Calorimetry titration curves at different c-values. 118 The substrate concentration at which a protein reaches 50% saturation is known as the Michaelis constant (Km), which is an indicator for the enzyme’s substrate affinity. Since integral membrane proteins enact no chemical transformations – and, specifically, in vitro AmpG cannot even express an equilibrium between two partitions – they lack catalytic constants kcat’s; therefore, substrate Km’s can be approximated as their KD’s (Eqn. 3 and 4), the fundamental dissociation 68 constant between a substrate and its recognition protein. The apparent Km of AmpG for the simple GlcNAc-1,6-anhydro-MurNAc 1 is 100 uM according to Park. 13 This same apparent Km was inferred from Vocadlo’s spheroplast assay via the Cheng-Prusoff equation (Eqn. 5), 124 wherein KI ≡ Km for 1, and found to be 70 µM. 43 We also assume a single substrate binding site on AmpG; a single gated channel that recognizes anhydro-PG fragments only on its periplasmic side. Eqn. 3: Eqn. 4: Eqn. 5: The goal for executing an ideal AmpG calorimetric titration against our sugars such that c = 30 necessitates AmpG concentrations of 30 x 100 µM, or 3 mM – a challenging if not impossible proposition given the difficulty both in expressing reasonable quantities of the integral membrane protein and in concentrating the solution thereof. Moreover, should any of our compounds prove to have lower affinity to AmpG than GlcNAc-1,6-anhydro-MurNAc, even higher protein concentrations would be necessary to construct this ideal sigmoidal titration curve. Despite the challenges associated with running an ITC experiment with AmpG, our collaborators have obtained some promising preliminary results (Figure 2.37). The binding of substrate 1 (GlcNAc-1,6-anhydroMurNAc) to AmpG was shown to be endothermic and an initial experiment with 25 µM of the membrane protein solubilized in DDM gave an extrapolated KD of 542 µM ± 35 µM (Figure 2.37, left). The c-value in this experiment was only 0.046 (assuming one binding site per protein) which leads to significant error in the extrapolated thermodynamic values. This dissociation constant is somewhat higher than what has been reported in the literature, 69 perhaps because the protein is solubilized by detergent as opposed to being embedded within a membrane. In contrast, substrate 2 (GlcNAc-1,6-anhydroMurNAc-L-Ala) gave a larger extrapolated KD of 1780 µM ± 86 µM (Figure 2.37, middle) under similar experimental conditions (c-value = 0.012) – the presence of an alanine on the lactate evidently decreases binding affinity. One possible explanantion for this decreased affinity is that the alanine carboxylate bears a negative charge in a position that normally is occupied by a neutral amide amine in the natural PG substrates – running ITC on our substrates 3, 4, and 5 would help to validate such a claim. Finally, our collaborators have obtained initial results with AmpG embedded within nanodiscs (Figure 2.37, right). These experiments are quite challenging as they require a very large 12 L protein preparation to complete one experiment. In fact, we are unaware of any other groups carrying out ITC studying on integral membrane proteins embedded within nanodiscs. The experiments were run with substrate 1 and the results provided a dissociation constant of 176 µM ± 23 µM, which is more closely in agreement with the literature reported values, and a c-value of 0.06. This may be due to the notion that the environment surrounding the protein embedded with a nanodisc more closely resembles that of a protein within a membrane. It should be noted that when nanodisk-embedded AmpG was studied in the presence of substrate 2, ITC did not register any binding response whatsoever. 70 Figure 2.37 Our collaborators’ ITC results: titration curve and software-extrapolated chemical constants of AmpG-PDC interacting with substrate 1 (left), AmpG-PDC interacting with substrate 2 (middle), and AmpG-nanodisc interacting substrate 1 (right). Clearly, more work will need to be done in optimizing these ITC experiments and efforts are being made to increase the c-values used in these experiments and to use specific protocols for analyzing data obtained with low c-values. It will be inormative to run a negative control with a truncated substrate, 1,6-anhydroMurNAc, that is known not to be recognized by AmpG – this work is currently underway. Similarly, analyzing inactive mutants of AmpG could provide insight into the role of specific residues in binding versus transport. Experimental Constant Measured Value KD 542 µM ± 35 µM Assumed N 1 c-value 0.046 Experimental Constant Measured Value KD 1780 µM ± 86 µM Assumed N 1 c-value 0.012 Experimental Constant Measured Value KD 176 µM ± 23 µM Assumed N 1 c-value 0.06 71 Despite the challenges of using ITC with AmpG, it should be noted that there are literature examples of similar integral membrane proteins with even higher dissociation constants analyzed in this fashion. For example, the xylose transporter XylE (also solvated in DDM micelles) displayed an extrapolated KD = 350 ± 30 M with c ≈ 0.29 towards its natural substrate (Figure 2.38). 103 The truncated derivative of the putative bacterial homologue of the iron-exporting ferroportin, BbFPNΔC, gave an extrapolated KD = 195 µM with c ≈ 0.26 towards Co2+ – as expected, point mutations made to key residues in BbFPNΔC’s substrate-binding pocket drastically raised the KD to values ranging from 500-900 µM (Figure 2.39). 125 Likewise, the arginine-agmatine antiporter AdiC yielded an extrapolated KD = 93 µM with c ≈ 1.6 towards arginine and KD = 29 µM with c ≈ 5.2 for agmatine – these C-value are higher than the previous examples, but still nowhere near the ideal c = 30 sigmoidal titration curve (Figure 2.40). 126 Thus, while our current AmpG ITC results may not be optimal, with more precise datapoints, they may be publishable. 127 Figure 2.38 ITC curve of D-xylose binding to XylE reported in the literature. 103 72 Figure 2.39 ITC curves of Co2+ binding to BbFPNΔC (leftmost) that have been reported in the literature. Binding affinity decreases as point mutations are introduced to BbFPNΔC (right three traces). 125 Figure 2.40 ITC curves of arginine (left) and agmatine (right) binding to AdiC as reported in the literature. 126 In summary, we have synthesized a panel of four potential AmpG substrates, either lacking, or bearing peptides of various lengths. Notably, we have prepared an isosteric analog of the GlcNAc-1,6-anhydroMurNAc pentapeptide in which meso-oxa-Dap replaces meso-Dap. This allows for a much more rapid and higher yielding synthesis than has been currently reported for other PG fragments bearing meso-Dap. Our collaborators have been successful in purifying recombinant AmpG for the first time and can obtain the protein either detergent-solubilized or embedded within protein nanodiscs. Using ITC, we have obtained the first experimental results showing that purified AmpG is functional and recognizes our synthetic compounds as binding partners. 73 Chapter 3: Synthesis of Small Molecule AmpG Plug 3.1 Obstructing a Permease 3.1.1 Proposed Small Molecule Plug We envisioned that GlcNAc-1,6-anhydro-MurNAc-Ala could be appended to a sufficiently large water-soluble polymer and behave as a “plug” for AmpG. In theory, such a macromolecule would be received by the periplasmic side of AmpG, partially embed itself within the permease pore and remain “trapped” within the protein. The steric bulk of the water-soluble polymer would prevent passage through the pore, and the only fate of this complex would be release of the “plug” back to the periplasmic side of AmpG. Depending on how tightly such a compound binds to AmpG, it may serve as a competitive inhibitor of substrate transport or provide an excellent tool for crystallographers to “catch” this permease “in the act” of transferring substrate from one cellular environment to another; thereby lending new insights into the conformational change of the active protein. The structure of the target compound 6 is shown in Figure 3.1. The branched polymer we chose to employ was a dendron based on the 2,2-dihydroxymethylpropionic acid subunit. Branched polymers of varying “generations” of this subunit have been previously made in literature. 128 Such dendrons are very polar, while not imitating any naturally-occurring polar moieties (i.e. amino acids). They can tentatively be thought of as “biocompatible” 129 and their dipolar interactions with proteins are analogous to those of water. Moreover, such polar dendrons would not be expected to sequester themselves within any phospholipid bilayer 130 and/or permeate through these bilayers without a transport protein. 74 We also chose a moderately long and flexible PEG-based linker that would allow the disaccharide to access the binding site of AmpG, while keeping the bulky dendron portion projected out into solution. Vocadlo’s studies, which showed that fluorescent probes could be linked in a similar fashion and still serve as substrates, 43 44 led us to suspect that our compound 6 might also bind to AmpG. Figure 3.1 Structure of our proposed AmpG “plug,” 6, with linker highlighted in blue and dendron subunit highlighted in red. 3.1.2 Protein Plug Domains The idea of membrane protein “plugs” is readily found in nature as so-called “plug domains”: protein subunits that evolved to buffer a given integral membrane protein against a premature/undesirable “open” conformation. Such domains are more often found regulating β-barrel pore proteins; often (but not always) as α-helical segments of the appropriate size so as to fully embed themselves within and to occlude the β-barrel aperture (Figure 3.2). 131 132 133 75 Figure 3.2 Literature models of β-barrel transmembrane proteins with plug domains: NalP from Neisseria meningitidis with plug domain as blue helix (left), 131 BtuB from Escherichia coli with only plug domain highlighted (middle), 132 and SusCD from Bacteroides thetaiotaomicron with plug domain highlighted purple (right). 133 Nevertheless, there are examples in Nature of plug domains present in α-helix integral membrane proteins – much larger but similar to AmpG. A notable and thoroughly researched example of such a plug-occluded integral membrane protein is SecY, a subunit of the prokaryotic SecYEG complex, responsible for the translocation (active transport) of proteins from the cytosol across the inner membrane and into the periplasm. 134 When the plug portion of the subunit is deleted via mutagenesis, SecY allows certain defective proteins to be translocated more efficiently than their intact counterparts; 135 moreover, electrophysiological measurements show the plug deletion mutant intermittently opening to “leak” small molecules and ions (Figure 3.3) 136 – in contrast, WT SecY is always closed in its resting state. Admittedly, our proposed sugar-linked polymer would not be a true “plug”: it wound not be covalently-bound to AmpG, but would instead behave more as a competitive inhibitor that binds to the recognition site in an intermolecular fashion. 76 Figure 3.3 Structure of WT SecY from Escherichia coli with plug domain highlighted in red (top left), periplasmic view of the obstruction the WT SecY pore by the same red plug domain (top right) in contrast to the periplasmic view of the same pore in a plug deletion mutant (bottom right). 135 BLM experiment of similar Escherichia coli plug deletion mutant shows intermittent ion passage (bottom left). 136 3.1.3 Planar Lipid Bilayer Inhibition Experiments There are numerous examples in the literature of the inhibition of integral membrane protein function being detected during planar lipid bilayer studies. The classic acetylcholine receptor (AcChoR) was reconstituted into a BLM 137 and demonstrated both the ready passage of sodium ions in the presence of the agonist carbamoylcholine (CbmCho) 138 and the complete blocking of the channel in the presence of the competitive antagonist, curare. 139 Subsequently, conductivity was re-established by the addition of more substrate (Figure 3.4, top). More recently, 77 the vesicle-reconstituted human large conductance voltage- and calcium-activated potassium channel (hBK channel) was shown to be directly inhibited by Alzheimers-indicative amyloid beta (Aβ) particles both on the “intracellular” (Ca2+-transporting) and on the “extracellular” (K+-transporting) side. 140 Often in the literature, these pA readings over a given trial time are tabulated as “current (pA) probabilities” (Figure 3.4, bottom left). These same Aβ particles have demonstrated the capacity to coalesce into a membrane-permeating potentially neurotoxic Ca2+ channel. 141 142 When incorporated in planar lipid bilayers, these Aβ channels can be irreversibly inhibited by promising enantiopure small molecule drug targets (Figure 3.4, bottom right). 143 78 Figure 3.4 Literature examples of permease inhibition measured by planar lipid bilayer experiments: acetylcholine receptor (AcChoR) opened by carbamoylcholine (CbmCho) and reversibly inhibited by curare (top), 137 hBK channel inhibited by amyloid beta (Aβ) particles from both the Ca2+ and K+ “sides” of the protein with current readings normalized as probabilities (bottom left), 140 self-assembling Aβ Ca2+ channels irreversibly inhibited by the small molecule drug “MRS2485” (bottom right). 143 79 3.2 Synthesis of Compound 6 3.2.1 Dendron Framework Our plan was to use a literature-known, water soluble, dendron as the “plug” portion of our inhibitor. We chose a 2,2-dihydroxymethylpropionic acid-based system as it has been widely used and is easy to synthesize. 128 144 145 2,2-Dihydroxymethylpropionic acid can be benzylated at its carboxylate to form the “stem” or “first generation” (G1) of the dendron 32 from which other 2,2-dihydroxymethylpropionates may branch (Figure 3.5). 2,2-Dihydroxymethylpropionic acid can have its alcohols masked as an acetonide so that indiscriminate polymerization cannot occur and dendron size can only ever increase one “generation” at a time. This acetonide-protected subunit can be converted to its anhydride via the action of DCC. After filtering off DCU, the anhydride is used as the active building block for the dendron. 145 146 The benzylated “stem” is doubly acylated by the action of the anhydride under mildly basic conditions with DMAP catalyst to give the protected 2nd generation (G2) dendron 33. Following deprotection by acidic Dowex in methanol, the process may be iterated once more to give the protected G3 dendron 34 or twice to give the protected G4 dendron 35. Hydrogenation then exposes the “stem” carboxylates of 33, 34, or 35 for subsequent coupling to the mono-benzoylated PEG linker to give protected Cbz-NH-linker-G2 dendron 36, Cbz-NH-linker-G3 dendron 37, and protected Cbz-NH-linker-G4-linker 38; respectively. 80 Figure 3.5 Synthesis of various 2,2-dihydroxymethylpropionate dendrons and their attachment to a flexible water-soluble linker. 81 3.2.2 Linking Sugar and Dendron Meanwhile, we appended an O-benzyl alanine onto the carboxylate of our GlcNAc-1,6-anhydro-MurNAc disaccharide 1 via a PyBOP mediated coupling (Figure 3.6). We then incorporated an acetonide between the 4-OH and 6-OH of the GlcNAc moiety via the action of tosic acid in acetone and dimethoxypropionic ketal to give compound 23. 147 This modification decreases the polarity of the disaccharide and allows its facile separation from residual peptide coupling material by flash chromatography. The acid-labile acetonide of 23 mirrors the acetonides of the dendron – thereby permitting global deprotection after coupling – and is a far more appropriate sugar protecting group than the acetyl groups used in Chapter 2 given the base-sensitivity of the dendron product’s many esters. Figure 3.6 Synthesis of disaccharide-alanine protected with an acid-labile acetonide. Absent any experimental data, we surmised that a G3 dendron would be a sufficiently large preliminary “plug” for AmpG when appended to our disaccharide. Both the disaccharide 23 and the linker-G3 dendron 37 were first deprotected by hydrogenolysis to reveal a carboxylate and amine, respectively (Figure 3.7). Coupling between the disaccharide and the linker-G3 dendron proceeded via the action of EDCI (PyBOP was a poor choice in this case) in good yields to give compound 39. Subsequent global deprotection via treatment with acidic Dowex-H proceeded 82 without issues to give the target compound GlcNAc-1,6-anhydro-MurNAc-Ala-linker-G3 dendron 6. Overall, compound 6 was produced from D-glucal via 1 in 13 steps and ~3.6% yield. Figure 3.7 Convergent synthesis of disaccharide-alanine linked to a G3 dendron. 83 3.3 Assays for Transport Inhibition by Small Molecule Plug Compound 6 was sent to the Vocadlo lab to test against their fluorescent AmpG probe in their spheroplast assay (Figure 3.8). 43 Results are still pending. Furthermore, this potential inhibitor was given to the Strynadka lab to test in a planar lipid bilayer experiment against the GlcNAc-1,6-anhydro-MurNAc AmpG natural substrate. Efforts are currently underway to reconstitute AmpG from vesicles into the black lipid membrane. Thereafter, the first ever electrophysiological measurements of AmpG activity – and indeed the first measurement of AmpG activity in vitro – will be conducted. Figure 3.8 Competitive inhibition of AmpG by dendron “plug” prevents transport of fluorescent probe in Vocadlo’s spheroplast assay 43 – fluorescence signal should decrease in spheroplasts exposed to both inhibitor and probe relative to a spheroplast “control” group only exposed to the probe. Unlike certain ion channels studied in literature, 148 149 150 we anticipate no voltage-gated behavior; therefore, AmpG should stay “closed” (i.e. give no current signal) even when 84 incorporated into the black lipid membrane and under electric potential. When substrate is introduced to one chamber of the bilayer setup, symport of both the substrate diffusing to a lower concentration gradient and ion(s) (presumably H+) travelling to a lower electric potential across the membrane should be detected as picoamp (pA) spikes in current between the two chambers. If our “plug” were also added along with substrate, we might expect that the frequency of these current spikes should significantly decrease (Figure 3.9), and at sufficiently high inhibitor concentrations, passage event signals could disappear entirely. Alternatively, it may be possible that the binding of compound 6 would hold the AmpG pore in an “open” state and that one could observe current passage even in the absence of any substrate. In this case it would be possible to measure the affinity of compound 6 to AmpG using titration experiments. Finally, if the attachment of a linker/dendron to our disaccharide prevents the binding of compound 6 to AmpG, then the outcome of either assay would be unaffected by its presence. 85 Figure 3.9 Proposed detection of AmpG inhibition by the dendron plug in a planar lipid bilayer experiment. 86 Chapter 4: Synthesis and Evaluation of An AmpG Photoaffinity Probe 4.1 Background 4.1.1 Photoaffinity Theory For our final project, we aspired to design and to test an anhydro-PG-derived photoaffinity probe for AmpG. Photoaffinity probes, as their names would imply, are compounds whose affinity for their target proteins increase in the presence of light. More specifically, they are molecules with functional groups that, upon irradiation with specific wavelengths of UV light, undergo reactions that generate highly reactive intermediates. These intermediates form covalent bonds with any molecule in their vicinity – ideally the target proteins to which they bind. 151 152 153 These photoreactive molecules are excellent tools both in vivo, when screening for drug targets, and in vitro, when elucidating protein binding domains. Although the latter application is of course more relevant to our project, the general procedure for a photoaffinity experiment is the same (Figure 4.1). 154 155 The organism or solvated protein is “fed” a pharmacophore of interest modified with a photoreactive moiety or “tag”. The solution is then irradiated with UV light to activate the photoaffinity tag, which irreversibly links to residues within the binding site of the protein that recognizes the pharmacophore. Thereafter, photolabelled proteins are separated from the rest of the experimental solutes via SDS-PAGE and/or affinity chromatography (when applicable) 156 157 158 159 160 161 Most photoaffinity tags also incorporate a bioorthogonal “handle” that allow for the attachment of fluorescent reporters and/or affinity groups (i.e. biotin for streptavidin columns) to greatly simplify the search for and purification of photoaffinity target “hits”. 152 155 162 Finally, purified photolabelled proteins are digested via commercially available proteases, and the resultant 87 fragments may resolved and analyzed via HPLC-coupled mass spectrometry for identification of tagged amino acids within their primary structure 151 155 (relative to the mass spectrum of untagged and digested protein). Binding domains of the protein target are identified when there is a preponderance of photoaffinity “hits” among only a cluster of amino acid residues. Figure 4.1 General procedure for the use of photoaffinity probes in both proteomics and enzymology experiments. 88 Literature examples of photoaffinity methodology being applied to ascertain the binding sites of sugars, 163 glycopeptides, 164 cholesterol recognition sites for porin proteins, 165 and key enzyme active site residues 155 166 167 have all been reported. Perhaps the most famous of the major facilitators, the bacterial lactose symporter LacY, had its recognition site residues tagged with a photoaffinity probe in the 1980’s. 160 161 Since then, an almost identical photoaffinity probe was used to locate key substrate binding domain residues of analogous bacterial melibiose symporter MelB. 168 We believe that we might be able to label the glycopeptide recognition residues of another membrane-spanning major facilitator, AmpG. 4.1.2 “Big Three” Photoaffinity Groups Photoaffinity groups are moieties that form highly reactive intermediates when exposed to ultraviolet light; these intermediates then covalently bind to any other molecules in their immediate vicinity. Examples of moieties which can behave as photoaffinity tags include benzophenones, which form carbonyl triplet diradicals; azides, which release N2 gas and give nitrenes; and diazirines, which release N2 gas to form carbenes (Figure 4.2). In the presence of proteins, carbonyl triplet diradicals tend to undergo C-H bond insertion reaction with amino acid residues (with a particular affinity for methionine methyls). 152 Nitrenes and carbenes usually covalently bind to the most Lewis basic functional groups available to them in typical buffered conditions: i.e. carboxylates, amides, and alcohols in decreasing priority. 169 As one would expect, not all photoaffinity tags are equally reactive or equally appropriate for any given project and we will give a brief overview of the aforementioned “big three” photoaffinity groups. 89 Figure 4.2 The “big three” functional groups in photoaffinity studies. Diazirines are small, ring-strained, and surprisingly non-polar functional groups. Among the “big three,” diazirines are the most sensitive to ambient conditions. 170 They tend to be activated at ~360 nm (well above be the 300 nm threshold at which proteins start to incur radiation damage) 171 and their carbenes tend to be the most reactive of the “big three,” but diazirines themselves tend to be the most difficult to append onto a given molecule. Often photoaffinity probes have their diazirine flanked by both a trifluoromethyl and a phenyl group; this ubiquitous modification prevents the premature “quenching” of carbene intermediates via intramolecular rearrangements (i.e. via a 1,2-hydride shift) but necessarily increases the photoaffinity tag size (Figure 4.3). Aliphatic diazirines are typically made by condensing a ketone with hydroxylamine-O-sulfuric acid in the presence of ammonia and then oxidizing the resultant diaziridine (Figure 4.4). Similarly, trifluoromethyl phenyl diazirines are made by condensing 2',2',2'-trifluoroacetophenone with the hydroxylamine, activating the ensuing oxime with tosyl chloride, generating a diaziridine with ammonia, and finally oxidizing (Figure 4.4). 152 154 172 90 Figure 4.3 Diazirine reactivity: aliphatic (top) and trifluoromethyl phenyl (bottom). Figure 4.4 General diazirine synthesis: A) aliphatic and B) trifluoromethyl phenyl. Benzophenones are sterically bulky moieties which are nevertheless exceptionally stable in ambient conditions and easy to make and append to a probe. 173 Triplet carbonyl diradicals are formed at the non-damaging wavelength of ~360 nm (Figure 4.5) and tend to react far more readily with organic molecules than with water solvent but are generated reversibly and are therefore slow to react. While benzophenones can be assembled “from scratch” if desired via a Fridel-Crafts acylation between a benzoyl chloride and another phenyl group (Figure 4.5), 174 175 they can also simply be purchased as benzoylbenzoic acids and coupled to the amines 176 177 or alcohols 177 178 of substrate mimetics. The combination of benzophenone’s limited reactivity and large size often 91 results in poor protein labeling for similar irradiation times; therefore, compensatory high substrate affinity, longer irradiation times, and/or high target protein concentrations become necessary. Figure 4.5 Benzophenone reactivity (top) and general synthesis (bottom). Aliphatic azides are both small and stable but require a shorter wavelength (250 nm) to activate, 179 which of course risks damaging the protein being studied. For this reason, researchers often opt to employ phenylazides, which can liberate N2 gas to give nitrene intermediates at longer wavelength (300 nm) UV light – again at the expense of tag size (Figure 4.6). 152 153 Admittedly this wavelength may be considered to be at the limit of what is acceptable for protein studies, but this potential problem may be ameliorated by further substituting these aryl azides. In general, azides can be thought of as the photoaffinity “middle ground” between diazirines and benzophenones: while not as potent as carbenes, nitrenes are also generated irreversibly; thereby making them more reactive than benzophenones. Azide synthesis is also trivial: aliphatic azides are made via an SN2 reaction between an alkyl-leaving group and NaN3 and aryl azides are made via Sandmeyer reaction of an aryldiazonium salt with NaN3 180 or via a diazotransfer between an aniline and an electrophilic azide (i.e. imidazole-1-sulfonyl azide) (Figure 4.7). 181 92 Figure 4.6 Azide reactivity: alkyl azide (top) and aryl azide (bottom). Figure 4.7 General azide synthesis: aliphatic (top), via SNAr (middle), and via diazo transfer (bottom). Less commonly used photolabile functional groups have also been investigated (Figure 4.8): 153 172 i.e. α-diazo carbonyl compounds, 156 o-nitrobenzyl alcohols, 182 pyrones, and pyrimidones. 183 Diazirines may also be used to generate THF oxocarbeniums as a reactive intermediate when flanked by a γ-ketone. 184 Recently, 2-aryl-5-carboxytetrazoles (ACT’s) have been developed as photoaffinity tags. 185 ACT’s generate carboxynitrile imines with the release of N2 at 302 nm and are touted as being “less reactive but more selective” than conventional carbene sources – seemingly preferring to intercept proximal carboxylate nucleophiles – and are obviously more stable to ambient conditions than diazirines. Nevertheless, azides, benzophenones, and diazirines constitute the vast majority of photoaffinity tags published in the literature. 93 Figure 4.8 Alternative photofunctional moieties and their reactive intermediates. 4.2 The Design of an AmpG Photoaffinity Probe 4.2.1 Choice of Photofunctional Group Of particular interest to our project is the diazirine moiety; specifically, an aliphatic diazirine. We reasoned that its small size and high reactivity are perfectly suited to tag the recognition site of AmpG. We need the probe to bind to the recognition site, the tentative KD of which has already been measured to be at best 100 µM; 13 43 therefore, we dare not risk adding the steric bulk of a trifluoromethyl phenyl diazirine, phenyl azide, or a benzophenone. Moreover, since the recognition site does not “tightly” bind to its PG substrate, we require an exceptionally reactive intermediate (i.e. a carbene) to immediately react with whatever Lewis basic moieties (i.e. carboxylates and alcohols) are responsible for H-bonding to the natural substrate prior to transmembrane passage. Although aliphatic diazirines are at the most risk of “off target/indiscriminate” reactions with solvated proteins, we intend to optimize photolabeling conditions until we can demonstrate a decreased photoaffinity trend in the presence of competitive 94 inhibitor such as disaccharide 1 or 2. If the regular substrate does indeed interfere with photoaffinity tagging, then the diazirine probe must surely be competing with these “inhibitors” for the same AmpG recognition site. This technique of “testing” photolabeling specificity with a known competing molecule is routinely implemented throughout the literature. 186–191 To be able to analyze such a trend, we must also plan to somehow conveniently and reliably quantify the degree of AmpG photolabeling. To this end, our probe should be able to accommodate some fluorescent reporter post-photoreaction. One of the most convenient and reliable methods of appending a fluorescent reporter onto any molecule is via CuAAc (or “Click”) chemistry – our need for an alkyne/azide pair to link a fluorescent molecule onto our photoaffinity probe further precludes the use of an aryl-azide as a photofunctional moiety. The pair of the smallest possible aliphatic diazirines that satisfies our reporter handle criteria are the literature-known compound with a functionalizable C-terminus, which we will henceforth call C-diaz, 192 193 and the literature-known compound with a functionalizable N-terminus, which we will henceforth call N-diaz (Figure 4.9). 194 Figure 4.9 Two literature-known small molecule diazirines with both a peptide coupling and a CuAAC handle. Some notable examples of the C-diaz probe used in the literature (Figure 4.10) include it labeling the binding pocket of human insulin-like growth factor 1 receptor (IGF-1R) when coupled to the 1,3,4-oxadiazin-5(6H)-ones (OPALs) anticancer agents; 193 “hitting” several proteins seemingly indiscriminately in cultured human lung cancer cells when coupled to the potent natural 95 product cytotoxin illudin S (ILS), thereby validating illudin’s DNA-modification cytotoxicity hypothesis; 195 and labeling both the phthalidimide binding pocket of the substrate engagement of cereblon (CRBN) and a novel target protein, eukaryotic translation initiation factor 3 subunit i (eIF3i), when coupled to the chemotherapeutic agent lenalidomide (Len). 188 Likewise, literature examples of the N-diaz probe’s utility (Figure 4.10) include it labeling the transmembrane protein CD147, a member of the immunoglobulin superfamily heavily expressed on the surfaces of tumor cells, when coupled to pseudolaric acid B (PAB); 187 a bevy of different cytosolic proteins implicated in (among other things) reactive oxidation species regulation and endosomal trafficking within leukemic macrophages when coupled to various poly-unsaturated fatty acids (PUFAs); 196 and binding to the active site of prostaglandin-endoperoxide synthase 2 (COX-2) when coupled to the known inhibitor indomethacin, a nonsteroidal anti-inflammatory drug (NSAID). 197 Figure 4.10 Examples of both the C-diaz tag (highlighted in red) used in the literature 188 193 195 and the N-diaz tag (highlighted in blue) used in the literature. 187 196 197 96 Perhaps most relevant to our current work is a recent paper published by the Hang group that sought to photolabel both nucleotide-binding oligomerization domains 1 and 2 (Nod1 and Nod2). 164 Nod1 and Nod2 are cytosolic proteins ubiquitous to eukaryotes that are thought to be responsible for recognizing bacterial PG fragments and inducing an immune response to them. Therein, the C-diaz was coupled to the glutamate nitrogen of iso-D-glutamate-DAP – yielding x-alk-iE-DAP – and to the glucosamine nitrogens of two muramyl dipeptides (MDPs) – yielding x-alk-MDP-L,D and x-alk-MDP-D,L (Figure 4.11). N-diaz was coupled to the carboxylate of the iso-glutamine sidechain to give MDP-D,L-x-alk (Figure 4.11). These probes were fed to mammalian cells, which were subsequently UV-irradiated, lysed, and their lysates analyzed. The group demonstrated that iE-DAP directly binds to Nod1; but not to Nod2. Likewise, only x-alk-MDP-L,D with the correct dipeptide stereoconfiguration and with its diazirine probe situated remotely from its stem peptide had demonstrably bound to Nod2; but not to Nod1. Inactive mutants of both Nod’s were unlabeled by any probes and MDP-probe specificity towards Nod2 was further confirmed via substrate competition with both MDP and GlcNAc-MDP (Figure 4.11). Moreover, x-alk-MDP-L,D had covalently bound to, and led to the discovery of, a novel MDP-binding protein, ADP-ribosylation factor 6 (Arf6), a GTPse whose unexpected labeling implies perhaps some role in the regulation of Nod2 in the presence of PG fragments. 97 Figure 4.11 Literature example of muramyl dipeptide (MDP) photoaffinity probes (top) binding directly to Nucleotide-binding oligomerization domain 2 (Nod 2), visualized by a fluorescent tag on and experiencing competitive inhibition on SDS-PAGE (bottom left); the same competitive inhibition by various concentrations of MDP and GlcNAc-MDP represented as a bar graph (bottom right) 164 98 4.2.2 Choosing an AmpG Substrate Group Based on the results of the original Park paper, 13 the molecule primarily recognized by AmpG is GlcNAc-1,6-anhydro-MurNAc; stem peptides of three, four, or five amino acids in length are also tolerated. As such, our photoaffinity probe must at least contain this disaccharide to preferentially interact with AmpG’s recognition site. We elected to add a L-alanine to the probe – unlike with larger peptides, a simple alanine is readily coupled onto the lactate. We surmise that this single amino acid makes the molecule more similar to AmpG’s natural glycopeptide substrates without incurring the synthetic complexity of working with a meso-Dap/meso-oxa-Dap stem peptide. Furthermore, given the aforementioned difficulties in coupling an amine to the disaccharide lactate, the alanine carboxylate may be a preferable platform on which to couple our N-diaz photoaffinity tag. An appropriate azide-bearing fluorescent reporter may be bound to the alkyne “tail” of the probe 7 (Figure 4.12) via Cu(I)-catalyzed azide–alkyne cycloaddition (CuAAC or “Click”) chemistry. Figure 4.12 Our proposed AmpG photoaffinity probe, 7, with the diazirine and alkyne-bearing N-diaz highlighted in blue. 99 4.2.3 Choosing a Fluorescent Reporter Our fluorescent reporter of choice (Figure 4.13) was sulforhodamine B-PEG3-azide (Abs/Em = 568/584 nm) – also known as “lissamine rhodamine B PEG3 azide” – recommended and provided to us by the Perrin group. While somewhat obscure, this particular fluorophore has been used successfully before in the literature 190 198 – interestingly, the aforementioned poly-unsaturated fatty acid (PUFA) photoaffinity study paired this reporter with an N-diaz-based probe 196 The distributer of the dye, Tenova Pharmaceuticals (https://tenovapharma.com/products/lissamine-rhodamine-b-peg3-azide, accessed Dec. 1, 2023), boasts of the reporter’s superior water solubility relative to even carboxyrhodamine, vibrant fluorescence emission, and a long spacer to reduce any steric effects on the labeled molecule – and a publication from the Perrin lab backs up these claims. 199 Once tagged with SulRho, any protein should be readily visualized on an SDS-PAGE gel with a fluorescence reader. Figure 4.13 Our proposed azide-bearing fluorescent tag. 100 4.2.4 Synopsis of AmpG Photolabeling Experiment Our plan was to receive AmpG from our collaborators – either in micelles or sequestered in peptide nanodiscs – irradiate it with UV light in the presence of our photoaffinity probe, “click” our sulforhodamine B reporter to it in the presence of protein denaturing sodium dodecyl sulfate (SDS) and return these photoreacted and fluorescently tagged samples to our collaborator in the Strynadka lab. We will also undertake photolabeling experiments in competition with natural substrate to verify probe specificity for AmpG’s substrate recognition site. Thereafter, our collaborator will run SDS-PAGE on our samples and analyze the resulting gel via fluorescence imaging. Once we can prove specific AmpG recognition domain binding, we will proceed in a joint effort to digest AmpG with trypsin and to analyze the ensuing fragments by LC-MS-MS. There, any fluorescently-tagged fragments would be clearly differentiated from their non-tagged counterparts and, knowing AmpG’s primary structure, we could identify the amino acids responsible for disaccharide recognition from the MS fragmentation patterns (Figure 4.14). Perhaps these discoveries, along with reference to previous theoretical models of the integral membrane protein’s structure, may finally identify AmpG’s glycopeptide binding domain. 101 Figure 4.14 Photoaffinity experimental design for both micellar and nanodisc-embedded AmpG. 102 4.3 Chemical Synthesis of the AmpG Photoaffinity Probe 4.3.1 Attempt to Follow Literature Synthesis of Diazirine Tag Our initial attempt to synthesize a photoaffinity probe utilized C-diaz as the labeling moiety. The carbon framework of C-diaz was assembled as described in literature. 164 Ethyl acetoacetate was converted to a dianion with ~2 equivalents of LDA at -78 ºC and alkylated at the more basic enolate with propargyl bromide to give compound 40 (Figure 4.15). 200 We originally planned to maintain an intact carboxylate on the tag and bind it to our disaccharide via a short linker (i.e. ethylene diamine), thereby assembling a full photoaffinity tag in only 3 steps. The ethyl ester could be readily saponified. However, despite repeated attempts at following the literature procedures that utilized hydroxylamine-O-sulfuric acid in the presence of ammonia, followed by oxidation, 193 196 we were unable to form any diazirine carboxylate. Figure 4.15 Initial attempt to make C-diaz. Realizing that the conversion of a ketone into a diazirine would be more troublesome than expected, we decided to change our synthetic target to the aforementioned amine-bearing N-diaz, (Figure 4.16) 194 201 which could also be coupled directly to our disaccharide without an intervening linker. 103 Figure 4.16 Literature synthetic route for N-diaz. 194 Ketone 40 was protected as an ethylene ketal by reaction with ethylene glycol and tosic acid catalyst (Figure 4.16). The ester 41 was then selectively reduced in the presence of LAH to give alcohol 42. Before proceeding with the rest of our planned synthetic route, we decided to optimize our diazirine-forming methodology via a test reaction: ketone 43 could be produced consistently and in large quantities from the Dowex-catalyzed deprotection of 42 and would serve as an excellent test reactant (Figure 4.17). The diazirine 44 is a literature-known photoaffinity tag, 194 201 202 203 and if N-diaz cannot be produced, 44 can instead be coupled to debenzylated disaccharide 23 (vide supra, Chapters 2 and 3). 203 We successfully produced a diazirine via the two-step literature process: fresh hydroxylamine-O-sulfonic acid in the presence of dissolved ammonia condenses to form crude diaziridine; then I2 oxidizes the diaziridine into diazirine 44, albeit in low yield. Figure 4.17 Our first successful synthesis of a test diazirine. 104 4.3.2 Alternative Synthetic Route to Diazirine Tag Although the literature methodology was simple to implement, the diazirine yields were much lower than we hoped (although not uniquely low throughout the literature) 193 201 and we chose not to exploit the high pressure liquid ammonia alternative reaction conditions. 194 204 205 As such, we sought another convenient method of forming aliphatic diazirines from the literature. We found a promising article describing the formation of diazirines 44 and C-diaz in high yields via an extended room temperature treatment of their corresponding ketones with an excess of the organic oxidant t-butyl hypochlorite in the presence of ammonia in methanol. 206 This t-butyl hypochlorite cannot be purchased due to its instability but can be produced as needed “in house” in multigram quantities from t-butanol reacting with bleach in the presence of acetic acid. 207 The hypochlorite method did produce the desired test diazirine 44 (Figure 4.18), but there was unfortunately no tremendous increase in yield relative to the aforementioned hydroxylamine-O-sulfonic acid method. Nevertheless, the cheapness and ease with which t-butyl hypochlorite is produced and subsequently reacted in the one-pot diazirine synthesis reaction makes the application of this more contemporary methodology preferable for our purposes. However, given our apparent difficulty in forming a diazirine and the relative sensitivity of a diazirine to ambient conditions, we aspired to modify the established synthetic route of the photoaffinity tag: the diazirine moiety would only be introduced at the last possible step in our tag synthesis. Figure 4.18 Successful synthesis of a test diazirine via the t-butyl hypochlorite reagent. 105 In our new approach, ketone 43 was converted to azide 45 via a mesyl intermediate (Figure 4.19) – the tosyl counterpart is insufficiently reactive and does not undergo SN2 with NaN3 even under reflux conditions. Conversion of 43 to the alkyl azide 45 via an alkyl mesylate intermediate and KI additive was accomplished in good yield. Thereafter, the azide underwent Staudinger reduction and subsequent DMAP-catalyzed t-butoxycarbonylation in the same pot. Both mono- and di-Boc products 46 and 47 were isolated, but yields were unexpectedly low. We surmised that having a free primary amine in the presence of a ketone may have been deleterious to the reaction (i.e. the formation of imine/enamine side-products), culminating in these poor yields. Figure 4.19 Synthesis of ketones poised for diazirine transformation. A simple adjustment was made to the synthetic route to circumvent any possibility of side-reactions (i.e. condensations or reductions) at this step: the ethylene ketal group of 42 would not be removed following LAH reduction, but instead be carried through the azidation reaction, Staudinger reduction, and Boc-protection steps (Figure 4.20). We attained azide 48 via another mesyl intermediate in a similarly good yield. As expected, the absence of a potentially reactive ketone greatly improved the yield of the one-pot reduction/carbamoylation step to give both mono- and di-Boc products 49 and 50. Figure 4.20 Protecting pro-diazirine ketones as ethylene ketals. 106 This initial success would be transient since we encountered another problem with the ethylene ketal group: under even the mildly acidic conditions required to remove a ketal, both Boc- groups of 50 would fall off before any of the ethylene ketal would (Figure 4.21). Although an ethylene ketal is a reliable protecting group, its high stability relative to that of a Boc was problematic. We felt that it was best to stay with a Boc protecting group on the primary amine as literature precedence showed an aziridine was stable to acidic deprotection conditions. 170 208 209 We therefore sought an alternative ketone protecting group. Figure 4.21 Boc groups are removed preferentially to ethylene ketal. Simultaneous to this finding, a colleague in the Tanner lab, Dmytro Voskoboinyk, had recently remedied a similar issue with his deprotection of an ethylene acetal: he had replaced the more stable ethylene acetal with the traceless, functionally identical, and far more labile dimethylacetal. Drawing inspiration from our colleague’s solution, we would again adjust our synthetic route to protect our ketone as a dimethylketal (Figure 4.22). Although there were no issues with either the ketal formation or LAH reduction step of 40 to give 51, the relative lability of the dimethylketal group complicated the workup of the subsequent azidation reaction. Yields of the azide 52 were unusually low (34%) via a mesyl intermediate; therefore, we opted to instead employ Appel methodology to accomplish azidation via an alkyl iodide intermediate. The Appel reaction gave azide 52 in higher yields (61%), although great care needed to be taken during the reaction workup because concentration of the iodoalkane intermediate must always be done in the 107 presence of base (i.e. NaHCO3) to prevent acid-catalyzed decomposition. The Staudinger reduction and subsequent Boc- protection of 52 proceed without issue to give either mono-Boc 53 without catalytic DMAP or di-Boc 54 with catalytic DMAP. As we predicted, the relative lability of the dimethylketal served our purposes well: this time, treatment of either 53 or 54 with PPTS cleanly exposed respective ketones 46 and 47 in quantitative yields with no loss of any Boc- groups in either case. Figure 4.22 Final successful syntheses of pro-diazirine ketones via the labile dimethylketal protecting group. Initially we attempted to convert the di-Boc ketone 47 into a diazirine. We thought that the absence of an amide N-H would help to limit unproductive hypochlorite side reactions (i.e. N-chlorination). Apparently, these assumptions were unfounded: the mono-Boc ketone 46 was converted to its corresponding diazirine 55 in higher yields than its di-Boc counterpart 56 (Figure 4.23). Coupling between the disaccharide 23 (vide supra in Chapters 2 and 3) post-hydrogenation and the diazirine 55 post-removal of Boc proceeded via PyBOP treatment in moderate yields to 108 give 57 (Figure 4.24). Acetonide deprotection was accomplished in quantitative yields by Dowex treatment to give our final product, photoaffinity probe 7. Relative to our starting D-glucal, 7 was synthesized via the route to 1 in 13 steps and ~2.6% yield. Figure 4.23 Synthesis of final diazirine tags via the t-butyl hypochlorite reagent. 109 Figure 4.24 Final coupling reaction to convergently assemble probe 7. 110 4.4 Reacting Photoaffinity Probe 7 with AmpG 4.4.1 Covalent Binding Between AmpG and Probe 7 The reactivity of probe 7 was tested with a Rayonet photochemical reactor: probe 7 was dissolved in distilled H2O and irradiated with 350 nm UV light for 50 mins until ESI-MS demonstrated complete disappearance of starting material. Under irradiation conditions, all of 7 was converted into compounds whose masses corresponded to either an elimination/cyclization product (loss of N2) or a H2O adduct product (loss of N2 with addition of H2O); in contrast, a “control” solution untreated with the photoreactor gave signals corresponding mainly to starting material with some minor signals corresponding to an elimination/cyclization product (presumably generated in the mass spectrometer during ionization since no H2O adduct was observed). We verified the efficacy of a CuAAC reaction in linking this photoreacted material to sulforhodamine B-PEG3-azide: the exceptionally polar [3+2] cycloaddition products were visible on TLC as a new low-lying purple band and absent in a control solution, which lacked the photoreacted alkyne. Furthermore, we tested the reliability of 7 linking to sulforhodamine B-PEG3-azide under both the buffer conditions proposed by our collaborators and the concentrations of the eventual denaturing SDS. In all cases, TLC displayed identical new UV-fluorescent product peaks. These results indicated that both the photoreaction and the click reaction were functional under the conditions we planned to use. Our collaborator in the Strynadka lab, Helena Sverak, began providing us with samples of E. coli AmpG – both as peptide-n-dodecyl-β-maltoside micelle detergent complexes (4 µM) and as peptide nanodiscs (1 µM) – isolated from E. coli and purified via affinity chromatography. According to our collaborator, the protein must be studied “fresh” and, to this end, requires ambient 111 temperatures of at most 0 ºC; as such, we would in all cases ensure the protein be kept on ice during photolabeling experiments. Our initial reactions sought to demonstrate successful protein tagging post-CuAAC reaction and SDS-PAGE. We arbitrarily decided to irradiate samples in the photoreactor for 20 mins in the presence of 100 µM of 7. These trials were compared to a control sample that lacked any irradiation. After irradiation, we proceeded to CuAAC tagging all of our samples with sulforhodamine B-PEG3-azide except for one, wherein we would test the efficacy of an alternative literature-known fluorescent tag, 3-azido-7-hydroxycoumarin (λabs 404 nm; λem 477 nm), 210 211 212 which we fully expected to be outperformed in both visibility and sensitivity by sulforhodamine B. Following SDS-PAGE of the irradiated and tagged samples, the resulting gels were visualized using a fluorescent reader (Figure 4.25 top images) to identifiy proteins that had been labeled, and by Coomassie blue staining (Figure 4.25 bottom image) to show total protein. Although the negative control behaved as expected – it gave almost no fluorescent signal with any labeling likely due to the reactivity of our probe under ambient light conditions – every other sample gave an intense and uniform signal (Figure 4.25). We had clearly labelled AmpG with probe 7 and the sulforhodamine B reporter successfully underwent [3+2] cycloaddition to the covalent complex regardless of whether the protein was first denatured via SDS. However, the probe had also reacted with the nanodisc membrane scaffold protein (MSP) seemingly as much as its intended target. To increase binding selectivity, we resolved to both lower our probe concentration a full order of magnitude (from 100 µM to 10 µM) and to change our irradiation source to a “Sun Lamp” in future experiments. The use of the 3-azido-7-hydroxycoumarin and visualization with a 532/28 emission filter (Figure 4.25, top right, lane 6) gave much less intense signals and was not used in further experiments. 112 Figure 4.25 SDS-PAGE of initial AmpG photolabeling with Rayonet photochemical reactor 350 nm irradiation for 20 mins and 100 µM of probe 7. Fluorescent tagging proceeds with sulforhodamine B-PEG3-azide. Sample 1 is a control: micelle-AmpG with probe 7 but no irradiation. Samples 2 and 3 have micelle-AmpG, but sample 2 used no SDS during “Click”. Samples 4 and 5 have nanodisc-AmpG, but sample 4 used no SDS during “Click”. Sample 6 has micelle-AmpG but is fluorescently tagged by 3-azido-7-hydroxycoumarin instead of sulforhodamine B-PEG3-azide. Sample visualization was achieved via a Biorad ChemiDoc MP imaging system with a 602/50 emission filter (top left), via a Biorad ChemiDoc MP imaging system with a 532/28 emission filter (top right), and via Coomassie blue staining that shows total protein regardless of labeling (bottom). 113 4.4.2 Selectivity of Probe Towards AmpG Active Site We next attempted to infer our probe’s selectivity for the recognition site of AmpG via a demonstration of competitive inhibition: we irradiated samples of AmpG with both probe 7 and disaccharide 2 in the hopes of lowering the extent of protein photolabeling with increasing concentrations of the compound 2. We also had the ancillary objective of demonstrating the integrity of AmpG even after being frozen, stored, and thawed; if AmpG were stable enough to be stored prior to use, researchers in general and our collaborators specifically would not need to prepare fresh protein immediately prior to each experiment. Since integral membrane protein reconstitution into nanodiscs necessarily sacrifices a significant amount of our collaborators’ precious (and already low) protein yield, we would forgo the use of nanodisc complexes until either evidence of active site binding is attained or a more native-like membrane environment proves to be an absolutely necessity. In our initial attempt, labeling of AmpG was still observed despite the lower concentration and less intense light source; however, there was seemingly no difference in photolabeling between any samples (data not shown), even at “competitive inhibitor” concentrations as high as 500 µM. We ran this preliminary inhibitory setup again – also with “frozen and thawed” protein – with competitive inhibitor 2 concentrations ranging from 0-500 M (closer to the reported KD of 100 M). We also ran a sample that lacked 2 but included bovine serum albumin (BSA) as a “dummy protein” to test for non-specific protein labeling (Figure 4.26). The labelling by 7 seemingly did preferentially target the micelle-complexed AmpG instead of the solvated BSA, since the visualization of the fluorophore (Figure 4.26 top image, lane 1) showed a more intense signal for AmpG than that observed with total protein staining (Figure 4.26 bottom image, lane 2). However, this may not be indicative of selective AmpG active site targeting as the literature has 114 reported a preference of various diazirine probes reacting with hydrophobic (i.e. membrane-associated) proteins instead of “hydrophilic” (i.e. cytosolic) ones. 169 As before, we did not see any obvious protection of AmpG due to increasing amounts of competitive inhibitor 2 indicating that the labeling we were seeing was non-specific. We attributed this indiscriminate photolabeling to our longer compensatory reaction times (50 mins) with our new “gentler” irradiation source. Moreover, we decided to use only “fresh/never frozen” protein in future experiments as the lack of selective binding could be explained by inactive protein that had been damaged by the freeze-thaw process. 115 Figure 4.26 SDS-PAGE of “frozen and thawed” micelle-AmpG photolabeling with 350 nm Sun Lamp irradiation for 50 mins, 10 µM of probe 7, and different concentrations of competitive inhibitor 2. The sample in band 2 contained bovine serum albumin (BSA) as a “dummy protein”. Sample visualization achieved via a Biorad ChemiDoc MP imaging system with a 602/50 emission filter on the (top) and Coomassie blue staining (bottom). 116 For “lower intensity” UV light sources, the literature recommends exposure periods of 5-10 mins for proper diazirine activation. 162 With “freshly harvested” AmpG, we would set up a series of experiments in which both irradiation times (1-30 mins) and competitive inhibitor 2 concentrations (0-500 µM) would vary. Additionally, we would carry out the labeling studies on crude membrane fractions prior to any detergent solubilization or affinity chromatography. These fractions contain a complex mixture of various membrane-bound proteins (including AmpG) and could be used to gauge target selectivity. Furthermore, the AmpG in these fractions is embedded in a more native-like environment and may have higher affinity to our probe. In the detergent solubilized samples, there was a definite dependence on UV exposure time and the intensity of reporter fluorescence (Figure 4.27, top left image) and we concluded that, in accordance with literature recommendations, a ~10 min exposure was sufficient to observe the fluorescence signal and hopefully minimize non-specific labeling. Unfortunately, there was once again no obvious correlation between fluorescence intensity and inhibitor concentration. Probe 7 also seemed to react non-specifically with the membrane fractions at all exposure times (Figure 4.27 lower images); therefore 7 was either completely unselective towards AmpG or in an environment of “target saturation” – i.e. there were too many off-target proteins relative to AmpG for 7 to reasonably associate with only its intended hydrophobic protein. A similar attempt was made by lowering probe concentration from ~10 µM to only ~2 µM and using both 10 min and 30 min irradiation times. As before, there was no obvious protection from compound 2 (Figure 4.28, top left image) and no obvious selection for AmpG in the membrane fractions (Figure 4.28, bottom left image). By this point we had come to suspect that probe 7 was not selectively adhering to its recognition site in AmpG regardless of irradiation time, protein “freshness,” or relative probe concentration. 117 Figure 4.27 SDS-PAGE of micelle-AmpG photolabeling via 350 nm Sun Lamp and 10 µM of 7 at different irradiation times and with different concentrations of competitive inhibitor 2: normalized fluorescence of AmpG in micelles (top left) and their Coomassie blue stained bands (top right); normalized fluorescence of AmpG in from crude cell membrane extracts (bottom left) and their Coomassie blue stained bands (bottom right). 118 Figure 4.28 SDS-PAGE of micelle-AmpG photolabeling via 350 nm Sun Lamp and 2 µM of 7 at different irradiation times and with different concentrations of competitive inhibitor 2: normalized fluorescence of AmpG in micelles (top left) and their Coomassie blue stained bands (top right); normalized fluorescence of AmpG in from crude cell membrane extracts (bottom left) and their Coomassie blue stained bands (bottom right). 119 Before abandoning our photoaffinity project, we made a final foray under the best possible conditions for selective recognition site labeling. We decided it was necessary to return to the use of AmpG embedded within peptide nanodisc rafts which should more closely mimic the protein in its native membrane environment. We would also retain the latest 2 µM concentration of probe 7 and a 10-minute Sun Lamp 350 nm irradiation period. Contemporaneous to this final attempt at proving probe selectivity, we attained ITC results that demonstrated AmpG’s association with disaccharide 1 (GlcNAc-anhMurNAc); therefore our inhibitor for most of these newest nanodisc samples would be 1 – only two samples would contain compound 2 (GlcNAc-anhMurNAc-L-Ala) at high (3 mM) and low (100 µM) concentrations for comparison. The results were similarly disappointing: no amount of either compound 2 or 1 (from 25 µM to 3 mM) ever changed the extent of protein photolabeling (Figure 4.29, left image). Moreover, 7 targeted the nanodisc membrane scaffold protein (MSP, lower band on gels) to the same extent as it did AmpG. Figure 4.29 SDS-PAGE of nanodisc-AmpG photolabeling via 10 mins of 350 nm Sun Lamp irraiation and 2 µM of 7 with different concentrations of either competitive inhibitor 1 (GlcNAc-anhMurNAc) or 2 (GlcNAc-anhMurNAc-L-Ala): normalized fluorescence of AmpG in nanodiscs (left) and their Coomassie blue stained bands (right). 120 Ultimately, it became clear that we were observing primarily non-specific protein labeling. Several explanations could be forwarded to explain this result. The most obvious is that probe 7 may not be recognized by AmpG due to the unnatural diazirine-alkyne moiety. Despite our attempts to keep the probe portion as sterically small as possible, and the precedence that Vocadlo’s fluorescent probe was accepted by AmpG, it may simply be the case that steric or electronic clashes with our compound and the AmpG’s binding site precludes binding. The binding of compound 7 to AmpG will ultimately be tested by ITC to see if this is the case. The observation of non-specific labeling of membrane-bound/solubilized proteins by compound 7 is likely due to the fact that it is quite non-polar and likely distributes within the detergent micelles preferentially to water. Favorable H-bonding interactions with the surfaces of proteins embedded within these micelles would localize the compound near these proteins and result in non-specific labeling. Other potential reasons for the lack of specific labeling could be that the protein is in an inactive conformation following purification or that a proton-gradient is required for substrate recognition. These possibilities are perhaps less likely given our preliminary ITC evidence for substrate binding by detergent-solubilized AmpG (Chapter 2). 121 Chapter 5: Overview and Future Work 5.1 Thesis Summary 5.1.1 Synthetic Contribution In summary, we have synthesized seven compounds to study the function of the periplasmic permease AmpG (Figure 5.1), which is implicated in both PG cell wall recycling and the biochemical pathway for the induction of β-lactam antibiotic resistance in Gram-negative bacteria. 25 Our first compound 1 is GlcNAc-1,6-anhydro-MurNAc, a known natural substrate for AmpG transport that has been chemically synthesized by many groups in the past. 43 49 73 We produced it starting from D-glucal in 9 steps with a ~12.2% overall yield. Compounds 2-7 are entirely novel and are also based off the GlcNAc-1,6-anhydro-MurNAc motif necessary for AmpG recognition. In fact, the synthetic routes of all these compounds (except for 5) go through 1 as a direct synthetic intermediate. Compounds 2 and 4 are putative natural substrates of AmpG and are synthesized from D-glucal in 11 steps (~6.1% yield) and 13 steps (~0.7% yield), respectively. Compounds 3 and 5 are putative natural substrate analogues of AmpG and are synthesized from D-glucal in 12 steps (~4.6% yield) and 12 steps (~1.6% yield), respectively. Intriguingly, compound 5 bears a stem pentapeptide with a meso-oxa-Dap isosteric analogue to the natural meso-Dap – the synthesis of this unnatural pentapeptide was developed in the Tanner lab. 68 72 Compound 6 is a possible competitive inhibitor and recognition site “plug” of AmpG that bears a 3rd generation (G3) branched 2,2-dihydroxymethylpropionyl-based water-soluble dendron. This glycodendron was synthesized from D-glucal in 13 steps (~3.6% yield). Compound 7 is a possible AmpG substrate and has a photoreactive diazirine moiety for the photoaffinity labeling of AmpG 122 as well as an alkyne “handle” for an azide-bound fluorescent tag. This photoreactive disaccharide was synthesized from D-glucal in 13 steps (~2.6% yield). Figure 5.1 Summary of key compounds 1-7 chemically synthesized in this work. 123 5.1.2 Biological Contribution Our collaborator in the Strynadka lab, Helena Sverak, has successfully expressed E. coli AmpG and isolated it both in detergent micelles and membrane scaffold protein nanodisc rafts. To measure the affinities of our compounds 1-5 towards AmpG, she intends to run ITC experiments for each compound. She found preliminary KD’s of 542 µM ± 35 µM for 1 towards AmpG in a PDC and 176 µM ± 23 µM for 1 towards AmpG in nanodiscs; supporting the notion that AmpG is functional. In contrast, she found a KD of only 1780 µM ± 86 µM for 2 towards AmpG in a PDC; extremely poor binding affinity. The ITC analyses of 3-5 are currently underway. Compound 6 was sent to both Stryndka and Vocadlo labs. The Strynadka group plans to measure in vitro AmpG inhibition by 6 with a planar lipid bilayer experiment against the background co-passage of ions along with one of our other substrates 1-5 through the artificial bilayer. Vocadlo plans to measure in vivo AmpG inhibition by 6 with his spheroplast-based fluorescent assay. 43 Results are pending. Photoaffinity experiments with diazirine 7 were undertaken by us using AmpG in both PDC’s and nanodiscs provided by the Strynadka lab. Analysis of the resultant photolabelled and fluorescently-tagged protein samples was completed by the Strynadka lab. Unfortunately, although we demonstrated photolabeling of AmpG by 7, we concluded that the labeling was nonspecific towards AmpG and its active site. Similarly hydrophobic membrane scaffold proteins were labelled as often as AmpG and we could never demonstrate any competitive inhibition between probe 7 and either substrates 1 or 2. 124 5.2 Future Work 5.2.1 AmpG Substrates Besides the aforementioned substitution of the carbodiimide-based EDCI for the phosphonium-based PyBOP in our peptide coupling reaction, we envision future attempts at producing the disaccharide pentapeptide 5 could be further improved by exchanging the acetyl groups of the glycosyl donor 20 with the even less polar butyryl groups of XV (Figure 5.2). These longer esters would greatly improve the efficiency of coupling product XVI purification: extraction would be even more efficient at eliminating polar impurities, and flash chromatography would provide cleaner separation from side-products. A per-butyrylated N-trichloroacetyl glucosamine could be produced initially and these butyryl groups may in theory survive the glycosyl bromination if not exposed to HBr in HOAc for too long. Otherwise, the acetyl groups may be removed after glycosylation and replaced by butyryl groups via butyric anhydride and pyridine/DMAP catalysis. 125 Figure 5.2 Proposed improved disaccharide pentapeptide chemical synthesis capitalizing upon butyryl protecting groups. 126 Although we never had a chance to integrate such a disaccharide into our synthetic schemes, we successfully produced and characterized tri-butyryl-GlcNAc-1,6-anhydro-MurNAc-L-Ala-OBn 58: post-alanine coupling, we treated the disaccharide 22 with butyric anhydride/pyridine with catalytic DMAP to get 58 in moderate yields – a reaction we surely would have optimized if the resultant product were more instrumental to our projects (Figure 5.3). Compound 58 would obviously need to be coupled to a meso-oxa-Dap tetrapeptide like P4 instead of a pentapeptide like P5; both of which were made previously in our lab. Figure 5.3 Synthesis and possible utility of butyryl-protected disaccharide-L-Ala 58. Our collaborators will continue to use the substrates we synthesized for their various planned calorimetric, crystallographic, and electrophysiological experiments to measure AmpG affinity and function. As procedures to reliably isolate AmpG and to test its affinity are developed – whether via ITC or some other technique like MicroScale Thermophoresis (MST) 213 – disaccharides bearing stem peptides of varying lengths will be required to gauge the necessity of each individual amino acid for AmpG’s substrate recognition. The disaccharide tripeptide GlcNAc-1,6-anhydro-MurNAc-L-Ala-D-iso-Glu-meso-oxa-Dap and the disaccharide tetrapeptide GlcNAc-1,6-anhydro-MurNAc-L-Ala-D-iso-Glu-meso-oxa-Dap-D-Ala (Figure 5.4) could be synthesized using our synthetic route and would complete the bevy of “natural” substrate isosteric 127 homologues available to the PG fragment permease. Of course, comparing the relative affinities of our meso-oxa-Dap isosteric analogues to those of true substrates bearing regular meso-Dap may also warrant investigation. Thereafter, it may be prudent to modify the configurations and/or identities of individual amino acids along the stem peptide; thereby fully understanding the structural requirements for AmpG binding site recognition, and hopefully explain Vocadlo and Holicek’s aforementioned perplexing and inconclusive findings in anticipation of introducing more exotic stem peptide molecular frameworks for uptake by AmpG. 44 Figure 5.4 Disaccharide tripeptides (left) and disaccharide tetrapeptides (right) as putative AmpG substrates: unnatural meso-oxa-Dap residues highlighted in red. 128 5.2.2 AmpG Plug Inhibitors If results from either electrophysiological or fluorescent assays are promising, dendrons of different sizes (i.e. G2 or G4) can be attached to GlcNAc-1,6-anhydro-MurNAc to find an optimally-sized “plug” (Figure 5.5, top). In all cases inhibitor results may be compared against those of GlcNAc-1,6-anhydro-MurNAc 1 and a linker-dendron (i.e. globally deprotected 36, 37, or 38) separately to verify that both an AmpG substrate portion and a dendron portion are necessary for a molecule to effectively “plug” the permease. Assuming successful acceptance of our sugar-dendron into AmpG’s recognition site, a true molecular plug may be envisioned wherein an electrophilic functional group (i.e. dehydroalanine) 214 or a photosensitive tag (i.e. a diazirine) may be incorporated into the linker or dendron to form a covalent bond between the inhibitor and the recognition site (Figure 5.5, bottom). Furthermore, if these dendron plugs are accepted by AmpG, we envision expanding the scope of macromolecular plugs to functionalized fullerene 215 or even gold nanoparticles. 216 Figure 5.5 Structures of possible AmpG “plugs” with different dendron sizes (top) or with reactive functional groups for covalent binding (bottom). 129 5.2.3 AmpG Photoaffinity Probes The use of probe 7 was shown to result in only non-specific labeling of AmpG. It is possible that probe 7 was too dissimilar to AmpG’s natural substrates to properly associate with its binding domain. One possible way to modify the probe would be to extend the stem peptide to include at least a meso-oxa-Dap-bearing tripeptide before coupling our diazirine tag to its C-terminus (Figure 5.6). Since AmpG has been shown to recognize disaccharides with tri- to pentapeptides, 13 the protein may instead tolerate a diazirine more remote from the disaccharide. Although there are numerous examples of integral membrane proteins being selectively photolabelled, there is also evidence to suggest that lipophilic photoaffinity probes such as ours tend to partition themselves within likewise hydrophobic portions of proteins (i.e. intramembrane domains) occasionally ignoring active/recognition sites in favour of these “off-target” associations. Regardless, a longer zwitterionic stem peptide would also increase probe polarity; thereby discouraging these “off-target” photoaffinity events. 130 Figure 5.6 Alternative AmpG photoaffinity probes bearing at least meso-oxa-Dap tripeptides for better protein affinity and the N-diaz tag (highlighted in blue). Assuming a certain amount of functional group tolerance between meso-Dap and the terminal D-Ala, we may envision appending other photolabile groups (i.e. a phenyl trifluoromethyl diazirine or even the aforementioned carboxytetrazole) onto the stem peptide terminus; perhaps another tag would be better suited for our purposes. Regardless, as our collaborators master the technique of ITC with AmpG, and assuming that an inordinate amount of protein is not required to attain accurate calorimetry readings, it may be prudent to first test any potential photoreactive substrate in a calorimetry experiment to see if their measured associations are sufficiently high. 131 5.2.4 Establishing a Proton Gradient Another reason for the failure to selectively label AmpG’s active site may instead be due to the mode of substrate uptake. Recall that in the Park paper, the proton decoupling agent CCCP had significantly decreased substrate uptake; 13 therefore AmpG may in fact co-transport a proton along with an anhydro-PG fragment. If indeed AmpG is a proton-substrate symporter, then its recognition site along with its cavity domain would be maximally functional along a proton gradient and proton-motive force may be required for the conformational change required for AmpG to engage in passive transport. Therefore, to study AmpG in vitro with a photoaffinity probe, the permease would first need to be reconstituted into a planar lipid bilayer (Figure 5.7). A pH or ion gradient can be established across the BLM prior to the introduction of PG fragments, i.e. 1, to “activate” AmpG and to verify its successful reconstitution and function. Then a photoaffinity probe, i.e. 7, may be added and both chambers irradiated by a UV lamp. The probe should irreversibly inhibit AmpG – perhaps even to complete inactivity – as it binds to its recognition and transport domains; thereafter, an electrical potential would be re-established to register this change in transport efficiency. Finally, after successful irreversible AmpG inhibition, the protein may be harvested from the artificial bilayer, reacted with sulforhodamine B-PEG3-azide under CuAAC conditions, digested by trypsin, and its fragments analyzed as mentioned in Chapter 4. 132 Figure 5.7 Establishing an in vitro proton gradient before photolabeling AmpG in a planar lipid bilayer setup. 133 Of course, an alternative route to circumvent the recalcitrance of AmpG using purified enzyme is to run a photoaffinity experiment on intact cells. The Vocadlo lab’s spheroplast assay 43 may in fact be the best way currently to achieve an AmpG substrate and proton gradient, and our probe is ideal for operating within cells since UV activation periods for aliphatic diazirines are neither too energetic nor too long to damage native proteins (Figure 5.8). Prior to any photoaffinity experiments, we might also be able to infer a KD for our probe when testing its ability to interfere with the uptake of Vocadlo’s fluorescent AmpG substrate. Like our collaborators in the Strynadka lab, Vocadlo expressed P. aeruginosa AmpG bearing hexahistidine tags within E. coli; therefore we could easily isolate photolabelled AmpG via metal affinity chromatography, append our fluorescent reporter to it, and proceed with trypsin digest and LC-MS-MS fragment analysis. Figure 5.8 Leveraging Vocadlo’s spheroplast assay to establish a proton gradient and attempting AmpG photolabeling in vivo. 134 Experimental General Information Chemicals were purchased from Millipore Sigma, Alfa Aesar, Fischer Scientific, Combi-blocks, Ark Pharm, and Chem-Impex Int. All chemicals were used directly without further purification unless noted. TLC plates were purchased from EMD Chemical Inc. Flash silica chromatography was conducted using silica gel SiliaFlash F60 (Silicycle, 230-400 mesh) or (where noted) high-purity Davisil Grade 633 (200-425 mesh, pore size 60 Å) silica gel. Strongly acidic hydrogen form Dowex® 50W X8 resin was purchased from Millipore Sigma. All reactions were carried out in Pyrex borosilicate glassware. 1H NMR spectra and proton-decoupled 13C NMR spectra were recorded on either a Bruker AV400sp spectrometer at a field strength of 400 MHz for 1H spectra and 101 MHz for 13C spectra or a Bruker AV600 spectrometer at a field strength of 600 MHz for 1H spectra and 151 MHz for 13C spectra. High-resolution spectra were recorded on a Waters/Micromass TOF spectrometer equipped with electrospray (ESI) ionization at the UBC Mass Spectrometry Facilities. 135 Photoaffinity Labeling Experiments We received all solutions of E. coli AmpG in EppendorfTM tubes from our collaborator and PhD candidate, Helena Sverak, in the Strynadka lab for our photoaffinity experiments: 100 µL of 4 µM AmpG solvated in DDM and HEPES buffer in the case of our AmpG-PDC samples or 60 µL of 1 µM AmpG solvated as lipid rafts bound together by a proprietary membrane scaffold protein in the case of our nanodisc samples – at the request of our collaborators, some of the details of solvating intact AmpG are left deliberately opaque. When not being prepared or used, all these received EppendorfTM tubes were kept in a covered Styrofoam ice box to limit possible thermal decomposition of the protein. Sulforhodamine B-PEG3-azide was taken with permission from the Perrin lab and originally purchased from Tenova Pharmaceuticals. Where applicable, we referred to previous photoaffinity literature for guidance with experimental setup and execution. 164 During irradiation, these EppendorfTM tubes were suspended in an ice-water 250 mL glass beaker via a thin perforated Styrofoam raft (Figure E1): this beaker with the sample array could either be inserted into a Rayonet RPR-100 immersion photoreactor with 15 lamps (RPR-3500Ǻ for 350nm emission) or placed 10 cm directly under a GE 275W SBMV "Sun Lamp" UV lamp – both UV sources were used with the permission of the Sammis lab. All EppendorfTM tubes were open during irradiation sessions and ice would be added to the beaker as necessary so that the bath never loses all its ice. The addition of all reagents and substrates to these samples were scaled with respect to their initial (either 100 µL or 60 µL) volumes. For simplicity, we will henceforth only refer to solution volumes added to 100 µL samples. Standard solutions of AmpG substrates were made and their concentrations varied depending on experiment parameters: in all cases probe 7 was dissolved in distilled water, substrate 1 was dissolved in distilled water, and substrate 2 was 136 dissolved in a 1:1 mixture of distilled H2O:MeOH. Appropriate volumes of these standard solutions were added to samples prior to irradiation such that desired concentrations would be achieved at a final volume of 130 uL: more buffer solution (either HEPES and micelle DDM buffer or nanodisc buffer) was added as necessary to reach these final volumes. After mixing the protein with the appropriate substrates, a period of “pre-incubation” was afforded for 15 mins prior to the irradiation session. After the photoreaction, samples were treated with SDS (10% w/v, 30 µL), CuSO4 (4 µL, 35.5 mM), sodium ascorbate (3 µL, 121 mM), and sulforhodamine B-PEG3-azide (7 µL, 2.6 mM dissolved in MeOH): the CuAAC reaction would then proceed at rt for 1 h. Finally, the reacted samples were taken back to our collaborator for further analysis. Figure E1 Illustration of photoaffinity experimental setup. 137 Synthetic Procedures Compound 2 To compound 22 (12 mg, 0.019 mmol) dissolved in 5 mL of MeOH was added 9 mg of 10% Pd/C (0.75 wt eq). This solution was stirred under a H2 atmosphere at rt for 4 h. This solution was then filtered through Celite® on top of a fine fritted glass funnel with MeOH eluent and subsequently concentrated under reduced pressure to give a clear colourless film (11 mg, quant). NMR samples were doped with MeOH as an internal standard for 13C NMR. 1H NMR (600 MHz, D2O with an internal standard of MeOH) δ 5.45 (s, 1H), 4.70 (d, J = 5.6 Hz, 1H), 4.66 (d, J = 8.4 Hz, 1H), 4.30 (d, J = 7.8 Hz, 1H), 4.16 (dq, J = 13.6, 7.0 Hz, 2H), 3.99 (s, 2H), 3.93 – 3.87 (m, 1H), 3.85 – 3.79 (m, 1H), 3.76 (td, J = 8.6, 3.3 Hz, 2H), 3.62 (s, 1H), 3.60 – 3.54 (m, 1H), 3.50 – 3.42 (m, 2H), 2.07 (s, 3H), 2.05 (s, 3H), 1.38 (d, J = 6.8 Hz, 3H), 1.36 (d, J = 7.2 Hz, 3H). 13C NMR (101 MHz, D2O) δ 174.33, 173.45, 172.62, 99.66, 98.88, 75.97, 74.99, 74.96, 73.78, 72.65, 72.33, 68.75, 63.80, 59.56, 54.52, 47.86, 47.77, 21.32, 20.90, 16.76, 16.66. HRMS (ESI): m/z calc for C22H35N3O13 [M+Na]+ 572.2068, found 572.2063. 138 Compound 3 Compound 24 (8.0 mg, 0.014 mmol) was diluted in 2 mL of MeOH and 2 mL of H2O and left stirring with 85 mg of 50W X8 hydrogen form Dowex resin at rt for 20 h. The solution was filtered through fritted glass via MeOH eluent and the filtrate was concentrated to give 3 as a clear colourless film (7.5 mg, quant). 1H NMR (600 MHz, Methanol-d4) δ 5.32 (s, 1H), 4.61 (d, J = 5.4 Hz, 1H), 4.49 (d, J = 8.4 Hz, 1H), 4.40 (q, J = 7.1 Hz, 1H), 4.31 (d, J = 7.5 Hz, 1H), 4.11 (q, J = 6.8 Hz, 1H), 3.98 (s, 1H), 3.89 (dd, J = 11.9, 2.2 Hz, 1H), 3.86 (s, 1H), 3.79 – 3.73 (m, 2H), 3.70 (dd, J = 11.9, 5.8 Hz, 1H), 3.56 – 3.53 (m, 1H), 3.44 (dd, J = 10.5, 8.5 Hz, 1H), 3.37 – 3.32 (m, 1H), 2.07 (s, 3H), 2.03 (s, 3H), 1.40 (d, J = 7.1 Hz, 3H), 1.37 (d, J = 6.8 Hz, 3H). 13C NMR (101 MHz, MeOD) δ 177.39, 174.98, 174.89, 173.36, 101.97, 101.78, 79.39, 78.17, 77.19, 75.61, 75.36, 74.59, 71.95, 65.84, 62.59, 57.08, 23.44, 22.63, 18.70, 18.33. HRMS (ESI): m/z calc for C22H36N4O12 [M+Na]+ 571.2231, found 571.2225. 139 Compound 4 To compound 26 (3.5 mg, 0.0041 mmol) dissolved in 5 mL of MeOH was added 22 mg of 10% Pd/C (6.3 wt eq). This solution was stirred under a H2 atmosphere at rt for 19 h. This solution was then filtered through Celite® on top of a fine fritted glass funnel with MeOH eluent and subsequently concentrated under reduced pressure to give 4 as a clear colourless film (2.8 mg, quant). NMR samples were doped with MeOH as an internal standard for 13C NMR. 1H NMR (600 MHz, D2O with an internal standard of MeOH) δ 5.48 (s, 1H), 4.72 (d, J = 5.4 Hz, 1H), 4.68 (d, J = 8.4 Hz, 1H), 4.43 (q, J = 7.2 Hz, 1H), 4.31 (d, J = 7.7 Hz, 1H), 4.23 – 4.15 (m, 2H), 4.04 – 3.99 (m, 2H), 3.91 (dd, J = 12.3, 1.8 Hz, 1H), 3.84 (dd, J = 7.9, 5.8 Hz, 1H), 3.81 – 3.75 (m, 2H), 3.65 (d, J = 2.8 Hz, 1H), 3.59 (dd, J = 10.5, 8.5 Hz, 1H), 3.50 (q, J = 9.9 Hz, 1H), 2.22 – 2.16 (m, 2H), 2.10 (s, 3H), 2.07 (s, 3H), 1.92 – 1.84 (m, 1H), 1.46 (d, J = 7.2 Hz, 3H), 1.41 (d, J = 6.8 Hz, 3H). 13C NMR (101 MHz, D2O) δ 182.66, 179.05, 175.97, 174.46, 174.29, 171.68, 101.36, 100.53, 82.37, 77.63, 76.69, 76.61, 75.41, 74.35, 73.98, 70.39, 65.46, 61.19, 56.17, 55.63, 50.01, 34.55, 29.33, 22.96, 22.54, 18.55, 17.98. HRMS (ESI): m/z calc for C27H42N4O16 [M-H]- 677.2596, found 677.2524. 140 Compound 5 To compound 31 (4.2 mg, 0.0028 mmol) dissolved in 2 mL of 1:1 MeOH:distilled H2O was added 5.8 mg of 10% Pd/C (1.4 wt eq). This solution was stirred under a H2 atmosphere at rt for 3 h. This solution was then filtered through a fine fritted glass funnel with distilled H2O eluent and subsequently concentrated under reduced pressure to give a clear colourless film. This resultant material was dissolved in 2 mL of 1:1 MeOH:distilled H2O and 15 mg of K2CO3 (0.11 mmol, 39 eq) was added. This basic solution was stirred for 3 h at rt; after which it was quenched with 50WX8 hydrogen form Dowex resin until the solution’s pH became 4-5. Finally, the mildly acidic solution was filtered through a fine fritted glass funnel with distilled H2O eluent and subsequently concentrated under reduced pressure to give compound 5 as a clear colourless film (1.7 mg, 62%). NMR samples were doped with MeOH as an internal standard for 13C NMR. 141 1H NMR (600 MHz, D2O with an internal standard of MeOH) δ 5.44 (s, 1H), 4.69 (d, J = 5.5 Hz, 1H), 4.65 (d, J = 8.4 Hz, 1H), 4.48 (t, J = 4.3 Hz, 1H), 4.37 (q, J = 6.1, 4.9 Hz, 1H), 4.35 – 4.31 (m, 1H), 4.27 (d, J = 7.9 Hz, 1H), 4.20 – 4.15 (m, 2H), 4.11 (q, J = 7.3 Hz, 1H), 3.99 (s, 1H), 3.97 (s, 1H), 3.96 – 3.84 (m, 4H), 3.82 – 3.72 (m, 6H), 3.60 (s, 1H), 3.56 (dd, J = 9.9, 8.1 Hz, 1H), 3.48 – 3.44 (m, 2H), 2.34 (t, J = 7.7 Hz, 2H), 2.15 (tt, J = 13.8, 6.7 Hz, 1H), 2.06 (s, 3H), 2.04 (s, 3H), 1.90 (dd, J = 14.6, 7.3 Hz, 1H), 1.42 (d, J = 7.2 Hz, 3H), 1.39 – 1.35 (m, 6H), 1.33 (d, J = 7.3 Hz, 3H). 13C NMR (151 MHz, D2O with an internal standard of CH3OH) δ 180.3, 178.2, 176.5, 176.2, 176.0, 174.5, 174.3, 174.3, 172.3, 172.3, 101.3, 100.5, 77.8, 76.7, 76.6, 75.2, 74.3, 74.0, 70.6, 70.4, 69.7, 65.5, 61.2, 56.2, 55.2, 54.8, 54.7, 51.5, 50.3, 50.3, 49.4, 32.3, 28.5, 23.0, 22.6, 18.7, 17.8, 17.7, 17.3. HRMS (ESI): m/z calc for C39H62N8O22 [M+H]+ 995.4051, found 995.4061. 142 Compound 6 Compound 39 (9 mg, 0.005 mmol) was diluted in 2 mL of MeOH and 2 mL of H2O and left stirring with 62 mg of 50WX8 hydrogen form Dowex resin at rt for 72 h. The solution was filtered through fritted glass via MeOH eluent and the filtrate was concentrated to give 6 as a clear colourless wax (8 mg, quant). 1H NMR (600 MHz, Methanol-d4) δ 5.33 (s, 1H), 4.63 (d, J = 5.4 Hz, 1H), 4.51 (d, J = 8.4 Hz, 1H), 4.40 (q, J = 7.1 Hz, 1H), 4.33 – 4.22 (m, 13H), 4.15 – 4.08 (m, 1H), 3.99 (s, 1H), 3.93 – 3.85 (m, 2H), 3.81 – 3.74 (m, 2H), 3.73 – 3.53 (m, 26H), 3.48 – 3.32 (m, 6H), 2.07 (s, 3H), 2.04 (s, 3H), 1.38 (dd, J = 9.2, 7.0 Hz, 6H), 1.31 (s, 3H), 1.30 (s, 6H), 1.15 (s, 12H). 13C NMR (151 MHz, MeOD) δ 175.95, 175.01, 174.93, 174.93, 174.59, 173.80, 173.38, 101.94, 101.79, 79.41, 78.13, 77.23, 75.64, 75.35, 74.62, 71.97, 71.31, 71.22, 70.45, 70.40, 68.14, 66.33, 65.86, 62.60, 57.09, 51.84, 50.08, 47.96, 47.77, 40.61, 40.38, 23.45, 22.66, 18.70, 18.45, 18.24, 18.13, 17.33. HRMS (ESI): m/z calc for C63H105N5O35 [M+H]+ 1492.6592, found 1492.6665. 143 Compound 7 Compound 57 (5.2 mg, 0.014 mmol) was diluted in 2 mL of MeOH and left stirring with 61 mg of 50WX8 hydrogen form Dowex resin at rt for 16 h. The solution was filtered through fritted glass via MeOH eluent and the filtrate was concentrated to give 7 as a clear colourless film (5.0 mg, quant). 1H NMR (600 MHz, Methanol-d4) δ 5.32 (s, 1H), 4.62 (d, J = 5.2 Hz, 1H), 4.50 (d, J = 8.4 Hz, 1H), 4.35 (q, J = 7.1 Hz, 1H), 4.31 (d, J = 7.5 Hz, 1H), 4.11 (q, J = 6.7 Hz, 1H), 3.98 (s, 1H), 3.89 (dd, J = 11.9, 1.8 Hz, 1H), 3.86 (s, 1H), 3.80 – 3.74 (m, 2H), 3.69 (dd, J = 11.9, 5.8 Hz, 1H), 3.54 (s, 1H), 3.44 (dd, J = 10.4, 8.6 Hz, 1H), 3.36 – 3.32 (m, 1H), 3.09 (ddt, J = 41.1, 13.8, 7.0 Hz, 2H), 2.27 (t, J = 2.6 Hz, 1H), 2.07 (s, 3H), 2.03 (s, 3H), 2.01 (d, J = 7.4 Hz, 2H), 1.66 – 1.60 (m, 4H), 1.40 (d, J = 7.1 Hz, 3H), 1.37 (d, J = 6.8 Hz, 3H). 13C NMR (101 MHz, MeOD) δ 175.02, 174.87, 174.71, 173.36, 101.99, 101.80, 79.45, 78.18, 77.25, 75.65, 75.38, 74.61, 71.99, 70.37, 65.87, 62.62, 57.10, 50.12, 35.43, 33.37, 33.24, 27.85, 23.43, 22.64, 18.46, 18.37, 13.84, 13.81. HRMS (ESI): m/z calc for C29H44N6O12 [M+Na]+ 691.2915, found 691.2905. 144 Compound 9 D-glucal (5.78 g, 39.6 mmol) was dissolved in 200 mL of MeCN and (Bu3Sn)2O (20 mL, 39.3 mmol, ~1 eq) was added. The solution was heated to 110 ºC and left to stir under Argon for 2 h. The solution was then cooled to rt and I2 (25g, 98 mmol, 2.5 eq) was added. This dark brown solution was left to stir at rt under Argon for 66 h. The iodine was quenched with the addition of 150 mL of sat aq Na2S2O3 and the MeCN was evaporated under reduced pressure while leaving behind the aqueous solution. This water solution was washed three times with 150 mL of Pet. Ether and then extracted four times with 150 mL of EtOAc. The EtOAc extracts were dried over Na2SO4, grav filtered and concentrated under reduced pressure to give a brown wax. This crude wax was dissolved in 40 mL of DMF and 10 mL of distilled H2O and NaHCO3 (5.15g, 61.3 mmol, 1.55 eq) and NaN3 (4.16g, 64.0 mmol, 1.62 eq) was added. This solution was heated to 100 ºC and stirred under Argon for 21 h. The resultant brown solution was cooled to rt and concentrated under reduced pressure to give a light brown solid. The solid was broken up and suspended as powdery clumps in 50 mL of EtOAc. Next, this solution was filtered through Celite® over fritted glass with EtOAc eluent and concentrated under reduced pressure to give a sticky brown film. The film was purified by silica gel chromatography. The column with step-wise gradient elution: 60% EtOAc:40% Pet. Ether → 70% EtOAc:30% Pet. Ether → 80% EtOAc:20% Pet. Ether. Compound 9 was isolated as light yellow crystals (4.34 g, 58% over 2 steps). In our hands, this modified synthesis of compound 9 gave similar yields to our rendition of the two-step procedure published 145 in literature without the need for either completely dry solvents or two separate large-scale chromatography purifications (63% vs. the current 58%). NMR and ESI mass spectral data were identical values reported in literature. 77 Compound 14 1,3,4,6-tetra-O-acetyl-β-D-glucosamine•hydrochloride (1.069 g, 2.79 mmol) was dissolved in 10 mL of pyridine and 10 mL of DCM and phthalic anhydride (497 mg, 3.36 mmol, 1.2 eq) was added. The solution was stirred at rt for 7 h. Ac2O (1.5 mL, 15.9 mmol, 5.7 eq) was then added and the solution was stirred at rt for a further 18 h. The solution was concentrated under reduced pressure then re-dissolved in 50 mL of DCM. This solution was washed twice with 30mL of aq 10% HCl, further washed twice with 30 ml of sat aq NaHCO3, dried over Na2SO4, filtered, and concentrated under reduced pressure to give a sticky light-yellow foam. The foam was purified by silica gel chromatography. The column with step-wise gradient elution: 40% EtOAc:60% Pet. Ether → 50% EtOAc:50% Pet. Ether. Compound 14 was isolated as pure white crystals (1.196 g, 90%). NMR and ESI mass spectral data were identical values reported in literature. 217 146 Compound 15 Compound 14 (292 mg, 0.61 mmol) was dissolved in 9 mL of degassed DMF and N2H5OAc (152 mg, 1.65 mmol, 2.7 eq) was added. The solution was stirred at rt for 1 h, after which it was diluted to 75 mL of EtOAc. This EtOAc solution was washed with 50 mL of 10% w/w LiCl, washed with 50 mL of sat aq NaHCO3, dried over Na2SO4, filtered, and concentrated under reduced pressure to give a pure white solid. This solid was dissolved in 1.5 mL and added dropwise over 15 min to a 1.5 mL DCM solution of 60% NaH in mineral oil (17 mg, 0.425 mmol, 0.88 eq) and Cl3CCN (0.3 mL, 2.99 mmol, 6.17 eq) vigorously stirring at rt under Argon. Once the slow addition was finished, the solution was quenched with the addition of pH 6.8 phosphate buffer and extracted twice with 50 mL of DCM. The organic extracts were dried over Na2SO4, filtered, and concentrated under reduced pressure to give a crude orange-brown oil. This oil was purified by silica gel chromatography. The column was run with step-wise gradient elution: 20% EtOAc:1% distilled NEt3:79% Pet. Ether → 30% EtOAc:1% distilled NEt3:69% Pet. Ether. Compound 15 was isolated as a light-yellow foam (140 mg, 40%). NMR and ESI mass spectral data were identical values reported in literature. 218 147 Compound 16 To 1 mL of distilled DCM was added 341 mg of pulverized flame-dried 4 Å molecular sieves, the glycosyl acceptor 13 (37 mg, 0.13 mmol), and the glycosyl donor 15 (315 mg, 0.543 mmol, 4.2 eq) under Argon. To this solution was slowly added 0.18 mL of 0.1 M BF3•OEt2 in DCM (0.018 mmol, 0.14 eq) was added, giving a yellow solution. The solution was left to stir at rt for 10 h. The solution was then quenched with 1 mL of NEt3, filtered through Celite® over fritted glass with MeOH eluent, and concentrated under reduced pressure to give a crude orange oil. This oil was purified by silica gel chromatography. The column was run with step-wise gradient elution: 60% EtOAc:40% Pet. Ether → 70% EtOAc:30% Pet. Ether → 80% EtOAc:20% Pet. Ether → 90% EtOAc:10% Pet. Ether → pure EtOAc. Compound 16 was isolated as a light-yellow foam (37 mg, 40%). NMR and ESI mass spectral data were identical values reported in literature. 73 148 Compound 18 To compound 21 (56 mg, 0.078 mmol) was added 1 mL of HOAc, 1 mL of EtOAc, and 1 mL of Raney Nickel slurry solution in H2O. The bubbling black mixture was stirred for 1 h at rt. The solution was then diluted with MeOH and filtered through Celite® over a fritted glass funnel. The filtrate was concentrated under reduced pressure to give a sticky bright green film. This material was dry loaded onto 150 mg of silica which was applied to a silica gel column. This column was run with step-wise gradient elution: pure EtOAc → 1% MeOH:99% EtOAc → 2% MeOH:98% EtOAc → 4% MeOH:96% EtOAc. Compound 18 was isolated as pure white crystals (35 mg, 73%). Alternatively, compound 21 (285 mg, 0.39 mmol) was dissolved in 12 mL of DMA and the solution was purged with Argon. AIBN (41 mg, 0.25 mmol, 0.64 eq) was added and the solution was stirred at 100 ºC for 1 h. The solution was cooled to rt and Bu3SnH (0.9 mL, 3.3 mmol, 8.6 eq) and more AIBN (9.6 mg, 0.059 mmol, 0.15 eq) was then added and the solution was stirred under Argon at 100 ºC for 2.5 h. The solution was then cooled to rt and concentrated under reduced pressure. The resultant wax was washed with 30 mL of hexanes to give a crude light yellow solid. This solid was purified by silica gel chromatography. The column was run with step-wise gradient elution: pure EtOAc → 1% MeOH:99% EtOAc → 2% MeOH:98% EtOAc → 4% MeOH:96% EtOAc. Compound 18 was isolated as pure white crystals (120 mg, 50%). 149 Moreover, compound 16 (85 mg, 0.12 mmol) may instead be dissolved in 5 mL of MeOH and N2H5OAc (115 mg, 1.25 mmol, 10 eq) was added. This solution was stirred at 90 ºC under an Argon atmosphere for 18 h. The solution was then cooled to rt, concentrated under vacuum to give a yellow wax. This wax and dissolved in 5 mL of pyridine and 1 mL of Ac2O and left to stir at rt for 68 h. This solution was then concentrated under reduced pressure to give a brown film. The crude film was purified by silica gel chromatography using high-purity Davisil Grade silica. This column was run with step-wise gradient elution: pure DCM → 2% MeOH:98% DCM → 3% MeOH:97% DCM. Compound 18 of the product was isolated as pure white crystals (41 mg, 55%). NMR and ESI mass spectral data were identical values reported in literature. 73 150 Compound 21 To a round bottom flask was added 1.41 g of pulverized flame-dried 4 Å molecular sieves and AgOTf (500 mg, 1.95 mmol, 2.09 eq) under an Argon atmosphere. Subsequently, the glycosyl donor 20 (1.10g, 2.14 mmol, 2.29 eq) 219 and the glycosyl acceptor 13 (270 mg, 0.934 mmol) 73 dissolved in 4 mL of distilled DCM were added. The solution was left to stir under light shielding conditions at rt for 48 h. The crude product mixture was diluted with MeOH and filtered through Celite® and a fritted glass funnel with ~60 total mL of MeOH. The light orange solution was concentrated under reduced pressure and the residue was purified by silica gel chromatography with step-wise gradient elution: 70% EtOAc:30% Pet. Ether → 80% EtOAc:20% Pet. Ether → pure EtOAc. Compound 21 was isolated as a light-yellow foam (525 mg, 78% yield). 1H NMR (600 MHz, MeOD) δ 5.41 (dd, J = 10.8, 9.2 Hz, 1H), 5.19 (s, 1H), 5.06 (t, J = 9.7 Hz, 1H), 4.90 (d, J = 8.5 Hz, 1H), 4.53 (d, J = 5.7 Hz, 1H), 4.32 – 4.27 (m, 2H), 4.18 – 4.13 (m, 2H), 4.05 (d, J = 9.0 Hz, 1H), 4.00 (dd, J = 10.8, 8.4 Hz, 1H), 3.88 – 3.82 (m, 2H), 3.73 (s, 3H), 3.67 (dd, J = 7.3, 5.8 Hz, 1H), 3.50 (s, 1H), 2.07 (s, 3H), 2.07 (s, 3H), 2.03 (s, 3H), 2.00 (s, 3H), 1.38 (d, J = 6.8 Hz, 3H). 13C NMR (101 MHz, MeOD) δ 174.8, 172.7, 172.3, 171.7, 171.2, 165.1, 102.0, 100.7, 78.5, 76.1, 75.5, 74.8, 73.2, 72.9, 70.0, 65.9, 63.1, 57.2, 52.6, 51.5, 22.8, 20.7, 20.6, 20.5, 18.7. HRMS (ESI): m/z calc for C26H35Cl3N2O15 [M+H]+ 721.1176, found 721.1176. 151 Compound 22 To compound 1 (20 mg, 0.042 mmol) was added DCC (43 mg, 0.21 mmol, 5 eq), HOBt•H2O (28 mg, 0.18 mmol, 4.4 eq), L-alanine(OBn)•hydrochloride (26 mg, 0.12 mmol, 2.8 eq), and DIPEA (0.04 mL, 0.23 mmol, 5.5 eq) in 1 mL of degassed DMF. This solution was left to stir at rt for 40 h, after which 1 mL of H2O and 1 mL of EtOAc was added and the solution was concentrated under reduced pressure to give an amber wax. The wax was partially purified via a quick silica plug. The plug was run with 1% Formic acid:99% EtOAc and the crude product was then flushed out with 30% MeOH:70% EtOAc. The flush fractions were concentrated under reduced pressure to give light yellow foam. This foam was then wet loaded with distilled H2O onto a 2 g reverse-phase C18 Sepack column. The column was run with step-wise gradient elution: 1% Formic acid:99% distilled H2O → 10% MeCN:90% distilled H2O → 20% MeCN:80% distilled H2O. The 20% MeCN fractions were concentrated and compound 22 was attained as a white solid (14 mg, 52%). 1H NMR (600 MHz, Methanol-d4) δ 7.42 – 7.29 (m, 5H), 5.30 (d, J = 2.0 Hz, 1H), 5.23 – 5.11 (m, 2H), 4.60 (d, J = 5.4 Hz, 1H), 4.50 (dd, J = 7.9, 6.2 Hz, 2H), 4.20 (d, J = 7.4 Hz, 1H), 4.12 (q, J = 6.7 Hz, 1H), 4.00 (s, 1H), 3.90 (dd, J = 12.0, 2.1 Hz, 1H), 3.84 (s, 1H), 3.77 (dd, J = 10.5, 8.3 Hz, 1H), 3.72 – 3.65 (m, 2H), 3.53 (s, 1H), 3.45 (dd, J = 10.5, 8.2 Hz, 1H), 2.06 (s, 3H), 2.03 (s, 3H), 1.44 (d, J = 7.3 Hz, 3H), 1.34 (d, J = 6.8 Hz, 3H). 152 13C NMR (101 MHz, MeOD) δ 175.29, 174.84, 173.66, 173.32, 137.24, 129.58, 129.33, 129.21, 101.99, 101.82, 79.40, 78.19, 77.18, 75.41, 75.35, 74.63, 72.01, 67.97, 65.78, 62.63, 57.09, 49.37, 49.33, 23.42, 22.63, 18.40, 17.59. HRMS (ESI): m/z calc for C29H41N3O13 [M+Na]+ 662.2538, found 662.2534. Compound 23 To compound 1 (37 mg, 0.077 mmol) was added PyBOP (66 mg, 0.13 mmol, 1.6 eq), HOBt•H2O (23 mg, 0.15 mmol, 2 eq), L-alanine(OBn)•hydrochloride (35 mg, 0.16 mmol, 2.1 eq), and DIPEA (0.04 mL, 0.23 mmol, 3 eq) in 2 mL of degassed DMF. This solution was left to stir at rt for 40 h, after which 1 mL of H2O and 1 mL of EtOAc was added and the solution was concentrated under reduced pressure to give an amber wax. This material was wet loaded with distilled H2O onto a 2 g reverse-phase C18 Sepack column. The column was run with step-wise gradient elution: pure distilled H2O → 10% MeCN:90% distilled H2O → 20% MeCN:80% distilled H2O. The 20% MeCN fractions were concentrated to give a clear colourless wax. To this clear wax was added 1 mL of degassed DMF, 1 mL of acetone, 1 mL of 2,2-dimethoxypropane, and HOTs•H2O (15 mg, 0.087 mmol, 1.1 eq). This solution was left to stir at rt for 24 h, after which 1 mL of EtOAc and several drops of sat aq NH4OH were added to quench the reaction and the solution was 153 concentrated under reduced pressure to give white precipitate. This crude product was wet purified by silica gel chromatography. The column was run with stepwise gradient elution: 5% MeOH:95% DCM → 6% MeOH:94% DCM → 7% MeOH:93% DCM. Compound 23 was attained as a white solid (23 mg, 44%). 1H NMR (600 MHz, Methanol-d4) δ 7.43 – 7.27 (m, 5H), 5.31 (s, 1H), 5.22 – 5.12 (m, 2H), 4.59 (d, J = 5.4 Hz, 1H), 4.54 (d, J = 8.4 Hz, 1H), 4.50 (q, J = 7.3 Hz, 1H), 4.20 (d, J = 7.4 Hz, 1H), 4.08 (q, J = 6.8 Hz, 1H), 3.98 (s, 1H), 3.92 – 3.78 (m, 3H), 3.77 (s, 1H), 3.69 (dd, J = 7.5, 5.7 Hz, 1H), 3.61 – 3.53 (m, 2H), 3.45 (s, 1H), 2.06 (s, 3H), 2.03 (s, 3H), 1.52 (s, 3H), 1.43 (d, J = 7.3 Hz, 3H), 1.39 (s, 3H), 1.32 (d, J = 6.8 Hz, 3H). 13C NMR (101 MHz, MeOD) δ 174.93, 174.86, 173.68, 173.29, 137.24, 129.58, 129.33, 129.21, 102.08, 102.05, 100.94, 79.47, 77.10, 75.67, 75.43, 74.42, 72.49, 68.66, 67.98, 65.75, 63.04, 57.61, 49.36, 49.12, 29.42, 23.43, 22.58, 19.30, 18.20, 17.65. HRMS (ESI): m/z calc for C32H45N3O13 [M+Na]+ 702.2850, found 702.2844. 154 Compound 24 Compound 23 (11 mg, 0.016 mmol) was dissolved in 2 mL of 7M NH3 in MeOH and left to stir at rt for 68 h. The solution was then concentrated under reduced pressure to give a clear colourless film. This film was purified by silica gel chromatography. The column was run with stepwise gradient elution: 7% MeOH:93% DCM → 8% MeOH:92% DCM. Compound 24 was attained as a white solid (8.0 mg, 85%). 1H NMR (600 MHz, Methanol-d4) δ 5.33 (s, 1H), 4.61 (d, J = 5.4 Hz, 1H), 4.55 (d, J = 8.4 Hz, 1H), 4.40 (q, J = 7.1 Hz, 1H), 4.30 (d, J = 7.5 Hz, 1H), 4.08 (q, J = 6.8 Hz, 1H), 3.97 (s, 1H), 3.91 – 3.74 (m, 5H), 3.61 – 3.52 (m, 2H), 3.46 (s, 1H), 2.08 (s, 3H), 2.04 (s, 3H), 1.53 (s, 3H), 1.40 (d, J = 7.4 Hz, 6H), 1.34 (d, J = 6.8 Hz, 3H). 13C NMR (101 MHz, MeOD) δ 177.37, 174.91, 174.65, 173.33, 102.05, 102.02, 100.94, 79.45, 77.07, 75.88, 75.44, 74.41, 72.51, 68.65, 65.83, 63.05, 57.61, 29.42, 23.43, 22.58, 19.28, 18.77, 18.10. HRMS (ESI): m/z calc for C25H40N4O12 [M+H]+ 589.2645, found 589.2717. 155 Compound 25 To compound 1 (41 mg, 0.086 mmol) was added PyBOP (72 mg, 0.14 mmol, 1.6 eq), HOBt•H2O (27 mg, 0.18 mmol, 2.1 eq), L-alanine(NHBn)•hydrochloride (38 mg, 0.18 mmol, 2.1 eq), and DIPEA (0.04 mL, 0.23 mmol, 2.7 eq) in 2 mL of degassed DMF. This solution was left to stir at rt for 46 h, after which 1 mL of H2O and 1 mL of EtOAc was added and the solution was concentrated under reduced pressure to give an amber wax. This material was wet loaded with distilled H2O onto a 2 g reverse-phase C18 Sepack column. The column was run with step-wise gradient elution: pure distilled H2O → 10% MeCN:90% distilled H2O → 20% MeCN:80% distilled H2O. The 20% MeCN fractions were concentrated to give a clear colourless wax. To this clear wax was added 1 mL of degassed DMF, 1 mL of acetone, 1 mL of 2,2-dimethoxypropane, and HOTs•H2O (15 mg, 0.087 mmol, 1.1 eq). This solution was left to stir at rt for 20 h, after which 1 mL of EtOAc and several drops of sat aq NH4OH were added to quench the reaction and the solution was concentrated under reduced pressure to give white precipitate. This crude product was purified by silica gel chromatography. The column was run with stepwise gradient elution: 6% MeOH:94% DCM → 7% MeOH:93% DCM → 8% MeOH:92% DCM. Compound 25 was attained as a white solid (23 mg, 40%). 156 1H NMR (600 MHz, Methanol-d4) δ 7.46 – 7.14 (m, 5H), 5.32 (s, 1H), 4.60 (d, J = 5.2 Hz, 1H), 4.55 (d, J = 8.3 Hz, 1H), 4.46 – 4.40 (m, 1H), 4.39 (d, J = 2.5 Hz, 2H), 4.29 (d, J = 7.6 Hz, 1H), 4.10 (t, J = 6.7 Hz, 1H), 3.97 (s, 1H), 3.93 – 3.70 (m, 5H), 3.63 – 3.50 (m, 2H), 3.46 (s, 1H), 2.07 (s, 3H), 2.03 (s, 3H), 1.53 (s, 3H), 1.40 (d, J = 7.6 Hz, 6H), 1.34 (d, J = 6.8 Hz, 3H). 13C NMR (101 MHz, MeOD) δ 174.91, 174.78, 174.76, 173.33, 139.78, 129.54, 128.38, 128.20, 102.03, 102.01, 100.93, 79.53, 77.13, 75.83, 75.42, 74.39, 72.49, 68.64, 65.82, 63.03, 57.59, 50.18, 49.85, 44.01, 29.42, 23.44, 22.58, 19.28, 18.55, 18.19. HRMS (ESI): m/z calc for C32H46N4O12 [M+H]+ 679.3110, found 679.3183. Compound 26 To compound 2 (20 mg, 0.036 mmol) dissolved in 0.5 mL of degassed DMF was added EDCI (35 mg, 0.18 mmol, 5.1 eq), DMAP (21 mg, 0.17 mmol, 4.8 eq), D-glutamate(OBn)2•hydrochloride (14 mg, 0.039 mmol, 1.07 eq), and DIPEA (0.06 mL, 0.34 mmol, 9.4 eq). This solution was stirred at rt with light shielding for 40 h. The reaction mixture was concentrated under reduced pressure to give a sticky yellow film. This film was first purified by silica gel chromatography. The column was run with step-wise gradient elution: pure DCM→ 2% MeOH:98% DCM → 4% MeOH:96% DCM → 8% MeOH:92% DCM → 10% MeOH:90% DCM. The resultant white foam was still 157 impure and was therefore wet loaded with distilled H2O onto a 2 g reverse-phase C18 Sepack column. The column was run with step-wise gradient elution: 1% Formic acid:99% distilled H2O → 10% MeCN:90% distilled H2O → 20% MeCN:80% distilled H2O → 30% MeCN:70% distilled H2O → 40% MeCN:60% distilled H2O. The 40% MeCN fractions were concentrated and compound 26 was attained as a clear colourless wax (3.5 mg, 12%). 1H NMR (600 MHz, Methanol-d4) δ 7.42 – 7.23 (m, 10H), 5.31 (s, 1H), 5.20 – 5.11 (m, 2H), 5.10 (d, J = 2.6 Hz, 2H), 4.59 (d, J = 5.3 Hz, 1H), 4.44 (q, J = 7.1 Hz, 1H), 4.27 (d, J = 7.6 Hz, 1H), 4.11 (q, J = 6.7 Hz, 1H), 3.97 (s, 1H), 3.89 (dd, J = 11.9, 2.1 Hz, 1H), 3.84 (s, 1H), 3.80 – 3.63 (m, 4H), 3.54 (s, 1H), 3.47 – 3.41 (m, 1H), 3.34 (d, J = 9.1 Hz, 1H), 2.44 (ddd, J = 9.3, 7.4, 2.9 Hz, 2H), 2.26 – 2.18 (m, 1H), 2.06 (s, 3H), 2.03 (s, 3H), 2.02 – 1.95 (m, 1H), 1.35 (dd, J = 6.9, 4.9 Hz, 6H). 13C NMR (101 MHz, MeOD) δ 175.03, 174.88, 174.87, 173.94, 173.35, 172.60, 137.51, 137.18, 129.58, 129.54, 129.40, 129.32, 129.29, 129.22, 101.96, 101.79, 79.43, 78.18, 77.25, 75.59, 75.38, 74.59, 71.99, 68.05, 67.43, 65.86, 62.62, 57.10, 53.08, 31.08, 27.52, 23.43, 22.63, 18.58, 18.45. HRMS (ESI): m/z calc for C41H54N4O16 [M+H]+ 859.3535, found 859.3610. 158 Compound 27 Dr. Soni’s protected pentapeptide Boc-P5 (28 mg, 0.026 mmol, 1.4 eq) was stirred with 2 mL of 4 M HCl in dioxane at rt for 2 hr. This pentapeptide solution was then concentrated under reduced pressure, re-diluted several times with diethyl ether, and subsequently re-concentrated to give a white hydrochloride salt. To this hydrochloride salt was transferred compound 1 (8.5 mg, 0.018 mmol), DMAP (10 mg, 0.082 mmol, 4.6 eq), EDCI (18 mg, 0.094 mmol, 5.2 eq), and DIPEA (0.03 mL, 0.17 mmol, 9.6 eq) dissolved in 0.3 mL of degassed DMF. This solution was stirred at rt with light shielding for 44 h. The reaction mixture was concentrated under reduced pressure to give a sticky orange film. This film was first purified by silica gel chromatography. The column was run with step-wise gradient elution: pure DCM → 4% MeOH:96% DCM → 8% MeOH:92% DCM → 10% MeOH:90% DCM. The resultant light-yellow foam was still impure and was therefore wet loaded with distilled H2O onto a 2 g reverse-phase C18 Sepack column. The column was run with step-wise gradient elution: 1% Formic acid:99% distilled H2O → 10% MeCN:90% distilled H2O → 20% MeCN:80% distilled H2O → 30% MeCN:70% distilled H2O → 40% MeCN:60% distilled H2O. The 40% MeCN fractions were concentrated and compound 27 was attained as a clear colourless film (7 mg, 28%). 159 1H NMR (600 MHz, Methanol-d4) δ 7.31 (ddt, J = 21.6, 6.6, 3.5 Hz, 20H), 5.29 (s, 1H), 5.21 – 5.04 (m, 8H), 4.61 (d, J = 10.5 Hz, 2H), 4.50 (d, J = 8.3 Hz, 1H), 4.47 (t, J = 3.9 Hz, 1H), 4.45 – 4.38 (m, 4H), 4.36 (q, J = 7.1 Hz, 1H), 4.27 (d, J = 7.4 Hz, 1H), 4.11 (q, J = 6.7 Hz, 1H), 3.96 (s, 1H), 3.92 – 3.82 (m, 3H), 3.81 – 3.66 (m, 5H), 3.64 (dd, J = 9.8, 4.8 Hz, 1H), 3.53 (s, 1H), 3.45 (t, J = 9.4 Hz, 1H), 2.36 – 2.17 (m, 3H), 2.06 (s, 3H), 2.03 (s, 3H), 1.95 (td, J = 14.9, 5.5 Hz, 1H), 1.37 – 1.32 (m, 10H), 1.29 (d, J = 7.1 Hz, 3H). 13C NMR (151 MHz, MeOD) δ 175.36, 174.95, 174.54, 173.82, 173.35, 172.64, 171.93, 171.61, 171.00, 170.63, 158.62, 138.03, 137.22, 137.11, 129.62, 129.58, 129.57, 129.53, 129.34, 129.33, 129.31, 129.27, 129.16, 129.12, 129.08, 128.88, 101.93, 101.73, 79.42, 78.14, 77.20, 75.50, 75.32, 74.55, 71.95, 71.81, 71.49, 68.19, 68.06, 67.87, 67.81, 65.84, 62.60, 57.06, 55.92, 55.15, 52.98, 50.44, 50.32, 49.68, 32.34, 27.80, 23.47, 22.68, 18.49, 18.27, 18.08, 17.23. HRMS (ESI): m/z calc for C25H36N2O15 [M+Na]+ 1421.5653, found 1421.5699. 160 Compound 28 Compound 18 (87 mg, 0.14 mmol) was dissolved in 3.5 mL of DMA (previously degassed under high vacuum). Subsequently, 142 mg (1.13 mmol, 8.07 eq) of 4-aminothiophenol and 120 mg of Cs2CO3 (0.368 mmol, 2.63 eq) were added. The light-yellow solution was heated to 110 oC under an Argon atmosphere for 30 min; after which the solution became dark orange. The mixture was quenched by the addition of 1 mL of formic acid and thereafter concentrated under reduced pressure to give a sticky light brown film. This film was purified by silica gel chromatography. The column was run with step-wise gradient elution: pure EtOAc → 2% MeOH:1% formic acid:97% EtOAc → 4% MeOH:1% formic acid:95% EtOAc. Compound 28 was isolated as a light-yellow foam (67 mg, 85%). 1H NMR (600 MHz, MeOD) δ 5.26 – 5.21 (m, 2H), 5.01 (dd, J = 10.2, 9.3 Hz, 1H), 4.77 (d, J = 8.4 Hz, 1H), 4.51 (d, J = 5.3 Hz, 1H), 4.32 (dd, J = 12.4, 4.4 Hz, 1H), 4.21 (m, 1H), 4.16 (d, J = 7.3 Hz, 2H), 4.12 (dd, J = 12.4, 2.3 Hz, 1H), 3.99 (s, 1H), 3.96 (dd, J = 10.7, 8.4 Hz, 1H), 3.86 – 3.81 (m, 2H), 3.68 (dd, J = 7.3, 5.8 Hz, 1H), 3.54 (s, 1H), 2.06 (s, 3H), 2.04 (s, 3H), 2.01 (s, 3H), 2.00 (s, 3H), 1.96 (s, 3H), 1.37 (d, J = 6.6 Hz, 3H). 13C NMR (101 MHz, MeOD) δ 174.3, 172.3, 171.8, 171.3, 102.1, 101.3, 77.4, 77.0, 75.3, 73.5, 73.0, 70.0, 66.1, 63.1, 55.4, 52.3, 23.1, 22.7, 20.7, 20.6, 20.6. HRMS (ESI): m/z calc for C25H36N2O15 [M+H]+ 605.2188, found 605.2195. 161 Compound 29 To compound 28 (11 mg, 0.018 mmol) was added DCC (16 mg, 0.078 mmol, 4.3 eq), DMAP (11 mg, 0.090 mmol, 5 eq), L-alanine(OBn)•hydrochloride (6 mg, 0.03 mmol, 1.5 eq), and DIPEA (0.03 mL, 0.17 mmol, 9.6 eq) in 0.3 mL of degassed DMF. This solution was left to stir at rt for 68 h, after which 1 mL of H2O and 1 mL of EtOAc was added and the solution was concentrated under reduced pressure to give a light-yellow wax. This film was initially purified by silica gel chromatography. The column was run with step-wise gradient elution: pure EtOAc → 1% MeOH:99% EtOAc → 2% MeOH:98% EtOAc → 3% MeOH:97% EtOAc. Still impure, the resultant light-yellow oil was then wet loaded with distilled H2O onto a 2 g reverse-phase C18 Sepack column. The column was run with step-wise gradient elution: pure distilled H2O → 10% MeCN:90% distilled H2O → 20% MeCN:80% distilled H2O → 30% MeCN:70% distilled H2O → 40% MeCN:60% distilled H2O → 50% MeCN:50% distilled H2O. The 50% MeCN fractions were concentrated and compound 29 was attained as a clear colourless film (2.7 mg, 19%). 162 1H NMR (600 MHz, Methanol-d4) δ 7.46 – 7.27 (m, 5H), 5.33 (s, 1H), 5.23 – 5.12 (m, 3H), 5.00 (t, J = 9.7 Hz, 1H), 4.71 (d, J = 8.4 Hz, 1H), 4.58 (d, J = 4.9 Hz, 1H), 4.51 (q, J = 7.2 Hz, 1H), 4.30 (dd, J = 12.4, 4.4 Hz, 1H), 4.21 (d, J = 7.5 Hz, 1H), 4.17 – 4.09 (m, 2H), 4.01 (dd, J = 10.7, 8.5 Hz, 1H), 3.97 (s, 1H), 3.82 (ddd, J = 10.1, 4.3, 2.2 Hz, 1H), 3.79 (s, 1H), 3.73 – 3.66 (m, 1H), 3.56 (s, 1H), 2.05 (s, 6H), 2.01 (s, 2H), 2.01 (s, 3H), 1.97 (s, 3H), 1.43 (d, J = 7.3 Hz, 3H), 1.34 (d, J = 6.7 Hz, 4H). 13C NMR (101 MHz, MeOD) δ 174.92, 174.53, 173.70, 173.19, 172.24, 171.84, 171.23, 137.24, 129.58, 129.34, 129.22, 101.99, 101.50, 79.09, 77.15, 76.58, 74.78, 73.40, 73.05, 70.04, 67.98, 65.79, 63.07, 55.15, 49.36, 23.17, 22.59, 20.68, 20.58, 20.54, 18.28, 17.67. HRMS (ESI): m/z calc for C35H47N3O16 [M+Na]+ 788.2851, found 788.2846. Compound 30 To compound 28 (21 mg, 0.035 mmol) was added HATU (66 mg, 0.17 mmol, 5 eq), HOBt•H2O (34 mg, 0.22 mmol, 6.3 eq), L-alanine-D-Glu(OBn)2•hydrochloride (18 mg, 0.042 mmol, 1.2 eq), and NMM (0.06 mL, 0.55 mmol, 15.6 eq) in 0.6 mL of degassed DMF. This solution was left to stir at rt for 22 h, after which 50 mL of EtOAc was added and the solution was washed twice with 30 mL of sat aq NaHCO3, dried over Na2SO4, filtered, and concentrated to give a light-yellow 163 wax. This film was purified by silica gel chromatography. The column was run with step-wise gradient elution: pure EtOAc → 1% MeOH:99% EtOAc → 2% MeOH:98% EtOAc → 3% MeOH:97% EtOAc. Compound 30 was attained as a clear colourless film (5.6 mg, 16%). 1H NMR (600 MHz, Methanol-d4) δ 7.36 – 7.28 (m, 10H), 5.33 (s, 1H), 5.23 – 5.13 (m, 3H), 5.10 (d, J = 3.1 Hz, 2H), 5.00 (t, J = 9.7 Hz, 1H), 4.71 (d, J = 8.4 Hz, 1H), 4.58 (d, J = 5.4 Hz, 1H), 4.49 (dd, J = 9.1, 5.2 Hz, 1H), 4.45 (q, J = 7.2 Hz, 1H), 4.31 – 4.26 (m, 2H), 4.15 – 4.09 (m, 2H), 4.01 (dd, J = 10.8, 8.4 Hz, 1H), 3.82 (ddd, J = 10.0, 4.4, 2.3 Hz, 1H), 3.80 (s, 1H), 3.74 (dd, J = 7.6, 5.7 Hz, 1H), 3.57 (s, 1H), 2.49 – 2.41 (m, 2H), 2.25 – 2.16 (m, 2H), 2.06 (s, 3H), 2.05 (s, 3H), 2.01 (s, 3H), 2.00 (s, 3H), 1.98 (s, 3H), 1.35 (dd, J = 6.9, 2.5 Hz, 6H). 13C NMR (101 MHz, MeOD) δ 174.85, 174.65, 174.55, 173.92, 173.22, 172.59, 172.24, 171.82, 171.22, 137.51, 137.18, 129.58, 129.54, 129.32, 129.28, 129.22, 129.20, 101.94, 101.48, 79.08, 77.15, 76.73, 74.75, 73.40, 73.06, 71.59, 70.04, 68.04, 67.43, 65.89, 55.16, 53.08, 31.08, 30.76, 27.53, 23.18, 22.60, 20.69, 20.58, 20.54, 18.71, 18.30. HRMS (ESI): m/z calc for C47H60N4O19 [M+Na]+ 1007.3746, found 1007.3741. 164 Compound 31 Compound 28 (17 mg, 0.028 mmol) was dissolved in 0.3 mL of distilled DCM and treated with HOBt hydrate powder (19 mg, 0.12 mmol, 4.4 eq), EDCI (21 mg, 0.11 mmol, 3.9 eq), and 0.05 mL (0.3 mmol, 10 eq) of Hunig’s base at rt and light shielding for 4 h. Meanwhile, Boc-P5 (35 mg, 0.033 mmol, 1.2 eq) was stirred with 2 mL of 4M HCl in dioxane at rt for 2 hr. This pentapeptide solution was then concentrated under reduced pressure, re-diluted several times with diethyl ether, and subsequently re-concentrated to give P5 as a white solid to which was transferred the activated ester solution. This reaction mixture was stirred at rt with light shielding for 36 h. The solution was diluted with 50 mL of DCM, washed with 2x30 mL of 0.2 M aqueous NaH2PO4, then washed with 2x30 mL of sat aqueous NaHCO3. The organic layer was dried over Na2SO4. This organic extract was concentrated under reduced pressure to give a light brown sticky film. The crude product was purified by silica gel chromatography using high-purity Davisil Grade silica. This column was run with step-wise gradient elution: pure EtOAc → 1% MeOH:99% EtOAc → 2% MeOH:98% EtOAc → 3% MeOH:97% EtOAc → 4% MeOH:96% EtOAc. Compound 31 was attained as a clear colourless wax (11 mg, 26%). Alternatively, Compound 29 (23 mg, 0.031 mmol) was diluted to 5 mL of MeOH and treated with 12 mg of 10% Pd/C (~0.5 wt eq) under H2 atmosphere for 3 h. The catalyst was filtered through Celite® via MeOH eluent and the filtrate was concentrated to give crude debenzylated 29 as a 165 clear colourless wax. To this wax was added EDCI (9 mg, 0.047 mmol, 1.5 eq), NHS (6 mg, 0.052 mmol, 1.7 eq) and 0.15 mL of degassed DMF. This solution was left to stir at rt with light shielding for 20 h. Then Dr. Soni’s tetrapeptide Fmoc-P4 (36 mg, 0.037 mmol, 1.2 eq) was stirred with 2 mL of 5% piperidine in DMF for 3 min. This tetrapeptide solution was then concentrated under reduced pressure to give a white foam. This foam was diluted to 0.3 mL of degassed DMF and added to the activated ester solution along with DIPEA (0.03 mL, 0.17 mmol, 5.5 eq). This mixture was left to stir at rt with light shielding for 48 h, after which it was diluted to 50 mL of DCM, washed twice with 30 mL of sat aq NaHCO3, dried over Na2SO4, filtered, and concentrated. The crude product was purified by silica gel chromatography. This column was run with step-wise gradient elution: pure EtOAc → 2% MeOH:98% EtOAc → 4% MeOH:96% EtOAc. Compound 31 was attained as a clear colourless wax (11 mg, 24%). 1H NMR (600 MHz, MeOD) δ 7.38 – 7.24 (m, 20H), 5.31 (s, 1H), 5.21 (dd, J = 10.8, 9.2 Hz, 1H), 5.18 – 5.04 (m, 8H), 5.01 (t, J = 9.7 Hz, 1H), 4.71 (d, J = 8.4 Hz, 1H), 4.62 – 4.57 (m, 2H), 4.47 (t, J = 4.1 Hz, 1H), 4.45 – 4.32 (m, 4H), 4.33 – 4.26 (m, 2H), 4.15 – 4.09 (m, 2H), 4.02 (dd, J = 10.8, 8.4 Hz, 1H), 3.94 (s, 1H), 3.87 (dd, J = 9.8, 4.6 Hz, 1H), 3.84 – 3.79 (m, 2H), 3.77 – 3.70 (m, 3H), 3.64 (dd, J = 9.8, 4.7 Hz, 1H), 3.55 (s, 1H), 2.48 – 2.14 (m, 3H), 2.05 (s, 3H), 2.03 (s, 3H), 2.01 (s, 3H), 2.01 (s, 3H), 2.02 – 1.99 (m, 1H), 1.98 (s, 3H), 1.40 – 1.25 (m, 12H). 13C NMR (101 MHz, CDCl3) δ 173.4, 172.9, 172.9, 172.8, 172.2, 171.9, 171.9, 171.8, 171.2, 170.8, 170.3, 170.2, 169.3, 156.5, 136.5, 135.6, 135.4, 135.4, 128.8, 128.7, 128.7, 128.6, 128.6, 128.6, 128.4, 128.4, 128.3, 128.2, 128.2, 101.4, 100.2, 77.8, 75.7, 74.3, 72.7, 72.3, 72.0, 71.4, 70.6, 68.2, 67.6, 67.3, 67.2, 67.1, 64.6, 62.0, 54.6, 54.3, 53.4, 51.1, 49.6, 48.5, 47.0, 31.4, 29.8, 28.0, 23.8, 22.8, 20.9, 20.8, 20.7, 17.9, 17.7, 17.6, 17.4. HRMS (ESI): m/z calc for C74H92N8O27 [M+Na]+ 1547.5964, found 1547.5946. 166 Compound 36 Protected G2 dendron 33 (42 mg, 0.078 mmol) was diluted to 5 mL of acetone and treated with 15 mg of 10% Pd/C (0.36 wt eq) under H2 atmosphere for 1 h. Thereafter this solution was filtered through Celite® and fritted glass with acetone eluent and concentrated under reduced pressure. This crude debenzylated 33 was diluted to 2 mL of DCM and DIPEA (0.12 mL, 0.69 mmol, 8.8 eq), PyBOP (76 mg, 0.15 mmol, 1.9 eq), and the linker mono-N-carboxybenzoyl-2,2′-(ethylenedioxy)bis(ethylamine)•hydrochloride (41 mg, 0.13 mmol, 1.7 eq) was added. This reaction mixture was stirred at rt for 16 h. The reaction mixture was then concentrated under reduced pressure to give a light-yellow foam. This foam was purified by silica gel chromatography. The column was run with step-wise gradient elution: 40% EtOAc:60% Pet. Ether → 50% EtOAc:50% Pet. Ether → 60% EtOAc:40% Pet. Ether. Compound 36 was attained as a clear colourless wax (47 mg, 85%). 1H NMR (400 MHz, Chloroform-d) δ 7.39 – 7.27 (m, 5H), 6.61 (s, 1H), 5.37 (s, 1H), 5.09 (s, 2H), 4.30 (q, J = 11.2 Hz, 4H), 4.16 (d, J = 11.9 Hz, 4H), 3.64 (s, 2H), 3.61 (s, 2H), 3.60 – 3.50 (m, 8H), 3.41 (dt, J = 24.3, 4.8 Hz, 4H), 1.41 (s, 6H), 1.35 (s, 6H), 1.25 (s, 3H), 1.12 (s, 6H). 13C NMR (101 MHz, Acetone) δ 206.12, 174.23, 172.84, 157.20, 138.45, 129.19, 128.66, 128.59, 98.55, 70.95, 70.90, 70.56, 70.17, 67.48, 66.80, 66.46, 66.44, 47.15, 42.73, 41.56, 40.09, 25.60, 22.54, 18.75, 18.14. HRMS (ESI): m/z calc for C35H54N2O13 [M+H]+ 711.3624, found 711.3698. 167 Compound 37 Protected G3 dendron 34 (80 mg, 0.074 mmol) was diluted to 5 mL of acetone and treated with 12 mg of 10% Pd/C under H2 atmosphere for 1 h. Thereafter this solution was filtered through Celite® and fritted glass with acetone eluent and concentrated under reduced pressure. This crude debenzylated 34 was diluted to 2 mL of DCM and DIPEA (0.12 mL, 0.69 mmol, 9.3 eq), PyBOP (80 mg, 0.15 mmol, 2.1 eq), and the linker mono-N-carboxybenzoyl-2,2′-(ethylenedioxy)bis(ethylamine)•hydrochloride (47 mg, 0.15 mmol, 2 eq) was added. This reaction mixture was stirred at rt for 18 h. The reaction mixture was then concentrated under reduced pressure to give a light-yellow foam. This foam was purified by silica gel chromatography. The column was run with step-wise gradient elution: 60% EtOAc:40% Pet. Ether → 70% EtOAc:30% Pet. Ether → 80% EtOAc:20% Pet. Ether. Compound 37 was attained as a white solid (83 mg, 90%). 168 1H NMR (600 MHz, D6-Acetone) δ 7.43 – 7.27 (m, 5H), 7.19 (s, 1H), 6.30 (s, 1H), 5.07 (s, 2H), 4.38 – 4.26 (m, 13H), 4.12 (d, J = 11.9 Hz, 8H), 3.65 (d, J = 11.8 Hz, 8H), 3.58 (s, 4H), 3.54 (t, J = 5.7 Hz, 4H), 3.40 (q, J = 5.6 Hz, 2H), 3.34 – 3.28 (m, 2H), 1.37 (s, 12H), 1.35 (s, 3H), 1.32 (s, 7H), 1.29 (s, 13H), 1.13 (s, 12H). 13C NMR (101 MHz, D6-Acetone) δ 206.10, 174.15, 172.83, 172.44, 157.20, 138.45, 129.20, 128.67, 128.60, 98.53, 70.95, 70.88, 70.56, 70.15, 67.90, 66.44, 66.40, 65.76, 47.68, 47.08, 42.71, 41.57, 40.19, 25.61, 22.57, 22.54, 18.79, 18.08, 17.99. HRMS (ESI): m/z calc for C61H94N2O25 [M+H]+ 1255.6148, found 1255.6221. Compound 38 Protected G4 dendron 35 (80 mg, 0.034 mmol) was diluted to 5 mL of acetone and treated with 18 mg of 10% Pd/C under H2 atmosphere for 2 h. Thereafter this solution was filtered through Celite® and fritted glass with acetone eluent and concentrated under reduced pressure. This crude debenzylated 35 was diluted to 1.5 mL of degassed DMF and DIPEA (0.07 mL, 0.4 mmol, 12 eq), 169 PyBOP (39 mg, 0.075 mmol, 2.2 eq), and the linker mono-N-carboxybenzoyl-2,2′-(ethylenedioxy)bis(ethylamine)•hydrochloride (30 mg, 0.094 mmol, 2.8 eq) was added. This reaction mixture was stirred at rt for 72 h. The solution was diluted with 50 mL of EtOAc, washed with 2x30 mL of sat aqueous NaHCO3. The organic layer was dried over Na2SO4. This organic extract was concentrated under reduced pressure to give a light-yellow foam. This foam was purified by silica gel chromatography. The column was run with stepwise gradient elution: pure DCM → 99% DCM:1% MeOH → 98% DCM:2% MeOH → 97% DCM:3% MeOH. Compound 38 was attained as a clear colourless wax (54 mg, 62%). 1H NMR (600 MHz, D6-Acetone) δ 7.43 – 7.28 (m, 5H), 7.20 (t, J = 5.2 Hz, 1H), 6.30 (s, 1H), 5.07 (s, 2H), 4.43 – 4.28 (m, 31H), 4.13 (d, J = 11.8 Hz, 18H), 3.66 (d, J = 12.8 Hz, 18H), 3.59 (s, 4H), 3.54 (t, J = 5.8 Hz, 4H), 3.41 (q, J = 5.7 Hz, 2H), 3.32 (q, J = 5.7 Hz, 2H), 1.37 (s, 30H), 1.35 (s, 7H), 1.33 (s, 14H), 1.29 (s, 24H), 1.13 (s, 24H). 13C NMR (101 MHz, D6-Acetone) δ 174.15, 172.83, 172.58, 172.33, 157.22, 138.45, 129.22, 128.70, 128.63, 98.55, 70.96, 70.88, 70.59, 70.21, 68.16, 66.66, 66.46, 66.41, 65.64, 47.67, 47.12, 42.72, 41.58, 40.22, 25.68, 22.53, 18.82, 18.14, 18.00. HRMS (ESI): m/z calc for C112H174N2O49 [M+Na]+ 2366.1082, found 2366.1080. 170 Compound 39 Compound 23 (11 mg, 0.016 mmol) was diluted to 5 mL of MeOH and treated with 10 mg of 10% Pd/C under H2 atmosphere for 2 hr. The catalyst was filtered through Celite® via MeOH eluent and the filtrate was concentrated to give crude debenzylated 23 as a clear colourless wax. Meanwhile, compound 37 (38 mg, 0.030 mmol, 1.9 eq) was diluted to 5 mL of MeOH and treated with 20 mg of 10% Pd/C under H2 atmosphere for 2 hr. The catalyst was filtered through Celite® on top of a fine fritted glass funnel via MeOH eluent and the filtrate was concentrated to give crude debenzylated 37 as a clear colourless wax. This wax was diluted to 1 mL of degassed DMF and added to the debenzylated 23 wax along with PyBOP (23 mg, 0.044 mmol, 2.8 eq), HOBt•H2O (8 mg, 0.052 mmol, 3.3 eq), and DIPEA (0.03 mL, 0.17 mmol, 11 eq). This solution was left to stir at rt for 40 h, after which it was diluted to 50 mL of EtOAc and washed twice with 30 mL of 1:1 sat aq NaHCO3:10% w/w aq LiCl, dried over Na2SO4, filtered, and concentrated. The crude product was purified by silica gel chromatography using high-purity Davisil Grade silica. This column was run with step-wise gradient elution: 5% MeOH:95% DCM → 6% MeOH:94% DCM → 7% MeOH:93% DCM. Compound 39 was attained as a clear colourless wax (18 mg, 67%). 171 1H NMR (600 MHz, Methanol-d4) δ 5.33 (s, 1H), 4.62 (d, J = 5.4 Hz, 1H), 4.56 (d, J = 8.4 Hz, 1H), 4.40 (q, J = 7.1 Hz, 1H), 4.38 – 4.25 (m, 14H), 4.13 (d, J = 11.8 Hz, 8H), 4.08 (q, J = 6.7 Hz, 1H), 3.97 (s, 1H), 3.91 – 3.75 (m, 5H), 3.69 (d, J = 11.9 Hz, 8H), 3.62 (s, 4H), 3.59 – 3.54 (m, 6H), 3.46 (s, 1H), 3.44 – 3.36 (m, 4H), 2.08 (s, 3H), 2.04 (s, 3H), 1.53 (s, 3H), 1.41 (s, 12H), 1.39 (s, 3H), 1.38 (d, J = 7.1 Hz, 3H), 1.34 (d, J = 6.8 Hz, 3H), 1.31 (s, 22H), 1.10 (s, 13H). 13C NMR (101 MHz, MeOD) δ 175.17, 174.90, 174.88, 174.60, 174.44, 173.52, 173.30, 102.04, 100.95, 99.39, 79.50, 77.09, 75.85, 75.44, 74.40, 72.50, 71.36, 71.28, 70.49, 68.65, 68.01, 67.04, 67.00, 66.23, 65.86, 63.05, 57.61, 49.99, 48.20, 47.65, 43.35, 40.62, 40.39, 29.43, 26.65, 26.63, 23.46, 22.61, 21.57, 21.55, 19.30, 18.80, 18.65, 18.23, 18.18. HRMS (ESI): m/z calc for C78H125N5O35 [M+Na]+ 1714.8054, found 1714.8050. 172 Compound 45 To compound 43 (105 mg, 0.83 mmol) was added MsCl (0.08 mL, 1.0 mmol, 1.2 eq), NEt3 (0.24 mL, 1.7 mmol, 2.1 eq) in 1 mL of DCM. This solution was left to stir at rt for 6 h. It was then diluted to 60 mL of diethyl ether and washed twice with 20 mL H2O, once with 10 mL of brine, dried over Na2SO4, filtered, and concentrated under reduced pressure to give an orange oil. To the resultant oil was added NaN3 (253 mg, 3.9 mmol, 4.7 eq), KI (400 mg, 2.4 mmol, 2.9 eq) and 3 mL of degassed DMF. This solution was left to stir at rt under light shielding for 30 h. It was then diluted to 60 mL of diethyl ether and washed twice with 20 mL H2O, dried over Na2SO4, filtered, and concentrated to give a light-yellow oil. This oil was purified by silica gel chromatography. The column was run with step-wise gradient elution: 5% EtOAc:95% Pet. Ether → 10% EtOAc:90% Pet. Ether → 15% EtOAc:85% Pet. Ether. Compound 45 was attained as a colourless liquid (78 mg, 64%). 1H NMR (400 MHz, Chloroform-d) δ 3.56 (t, J = 6.4 Hz, 2H), 2.70 (m, 4H), 2.47 (td, J = 7.2, 2.6 Hz, 2H), 1.96 (t, J = 2.7 Hz, 1H). 13C NMR (101 MHz, CDCl3) δ 205.72, 82.77, 69.09, 45.77, 41.80, 41.69, 12.98. HRMS (ESI): m/z calc for C7H9N3O [M+Na]+ 174.0644, found 174.0612. 173 Compound 46 To compound 53 (195 mg, 0.72 mmol) was added PPTS (37 mg, 0.15 mmol, 0.2 eq) in 5 mL of acetone and 1 mL of distilled H2O. This solution was left to stir at rt for 18 h, after which it was partially concentrated to remove acetone. Thereafter, the remaining aqueous solution was diluted to 60 mL of diethyl ether and washed twice with 20 mL of 0.2 M NaH2PO4, dried over Na2SO4, filtered, and concentrated. Compound 46 was attained as a colourless wax (162 mg, quant.). 1H NMR (400 MHz, Methanol-d4) δ 3.28 (t, J = 6.6 Hz, 2H), 2.67 (dt, J = 13.4, 6.9 Hz, 4H), 2.40 (td, J = 7.2, 2.7 Hz, 2H), 2.21 (t, J = 2.7 Hz, 1H), 1.42 (s, 9H). 13C NMR (101 MHz, MeOD) δ 209.38, 158.29, 83.88, 80.08, 69.79, 43.36, 42.43, 36.37, 28.73, 13.52. HRMS (ESI): m/z calc for C12H19NO3 [M+Na]+ 248.1263, found 248.1257. 174 Compound 47 To compound 54 (20 mg, 0.054 mmol) was added PPTS (23 mg, 0.092 mmol, 1.7 eq) in 2 mL of acetone and 1 mL of distilled H2O. This solution was left to stir at rt for 8 h, after which it was partially concentrated to remove acetone. Thereafter, the remaining aqueous solution was diluted to 60 mL of diethyl ether and washed twice with 20 mL of 0.2 M NaH2PO4, dried over Na2SO4, filtered, and concentrated. Compound 47 was attained as a colourless wax (17 mg, quant.). 1H NMR (400 MHz, Methanol-d4) δ 3.88 – 3.78 (m, 2H), 2.73 (dt, J = 14.8, 7.3 Hz, 4H), 2.40 (td, J = 7.2, 2.7 Hz, 2H), 2.22 (t, J = 2.7 Hz, 1H), 1.50 (s, 18H). 13C NMR (101 MHz, MeOD) δ 208.83, 153.82, 83.93, 83.83, 69.89, 42.72, 42.68, 42.41, 28.29, 13.60. HRMS (ESI): m/z calc for C17H27NO5 [M+Na]+ 348.1791, found 348.1785. 175 Compound 48 To compound 42 (104 mg, 0.61 mmol) was added MsCl (0.08 mL, 1.0 mmol, 1.6 eq), NEt3 (0.24 mL, 1.7 mmol, 2.8 eq) in 1 mL of DCM. This solution was left to stir at rt for 1 h. It was then diluted to 60 mL of diethyl ether and washed twice with 20 mL H2O, once with 10 mL of brine, dried over Na2SO4, filtered, and concentrated under reduced pressure to give an orange oil. To the resultant oil was added NaN3 (232 mg, 3.6 mmol, 5.9 eq), KI (410 mg, 2.5 mmol, 4.0 eq), and 3 mL of degassed DMF. This solution was left to stir at rt under light shielding for 49 h. It was then diluted to 60 mL of diethyl ether and washed twice with 20 mL H2O, dried over Na2SO4, filtered, and concentrated to give a light-yellow oil. This oil was purified by silica gel chromatography. The column was run with step-wise gradient elution: 2% EtOAc:98% Pet. Ether → 5% EtOAc:95% Pet. Ether → 10% EtOAc:90% Pet. Ether. Compound 48 was attained as a light-yellow liquid (73 mg, 61%). 1H NMR (400 MHz, Methanol-d4) δ 3.95 (s, 4H), 3.35 (t, J = 7.3 Hz, 2H), 2.33 – 2.14 (m, 3H), 2.03 – 1.77 (m, 4H). 13C NMR (101 MHz, MeOD) δ 110.37, 84.67, 69.37, 66.14, 47.67, 37.53, 36.77, 13.74. HRMS (ESI): m/z calc for C9H13N3O2 [M+H]+ 196.1006, found 196.1079. 176 Compound 49 To compound 48 (51 mg, 0.27 mmol) was added PPh3 (147 mg, 0.56 mmol, 2.1 eq) in 2 mL of THF and 0.2 mL of distilled H2O. This solution was left to stir at rt for 17 h, after which it was concentrated under reduced pressure. To the resultant solid was added 2 mL of THF, 0.06 mL of distilled NEt3, and Boc2O (522 mg, 2.39 mmol, 8.9 eq), and DMAP (24 mg, 0.20 mmol, 0.73 eq). This solution was left to stir at rt for 4 h, after which it was quenched with the addition of 2 mL of a 1:1 mixture of H2O:acetone. This quenched solution was diluted to 60 mL of diethyl ether and washed twice with 20 mL of 0.2 M NaH2PO4, dried over Na2SO4, filtered, and concentrated to give a white solid. This solid was purified by silica gel chromatography. The column was run with step-wise gradient elution: 5% EtOAc:95% Pet. Ether → 10% EtOAc:90% Pet. Ether → 15% EtOAc:85% Pet. Ether → 20% EtOAc:80% Pet. Ether. Compound 49 was attained as a colourless wax (43 mg, 59%). 1H NMR (400 MHz, Methanol-d4) δ 3.94 (s, 4H), 3.16 – 3.08 (m, 2H), 2.27 – 2.15 (m, 3H), 1.90 – 1.76 (m, 4H), 1.43 (s, 9H). 13C NMR (101 MHz, MeOD) δ 158.36, 110.84, 84.72, 79.97, 69.28, 66.04, 37.52, 37.43, 36.86, 28.77, 13.76. HRMS (ESI): m/z calc for C14H23NO4 [M+Na]+ 292.1525, found 292.1519. 177 Compound 50 To compound 48 (51 mg, 0.27 mmol) was added PPh3 (147 mg, 0.56 mmol, 2.1 eq) in 2 mL of THF and 0.2 mL of distilled H2O. This solution was left to stir at rt for 17 h, after which it was concentrated under reduced pressure. To the resultant solid was added 2 mL of THF, 0.06 mL of distilled NEt3, and Boc2O (522 mg, 2.39 mmol, 8.9 eq), and DMAP (24 mg, 0.20 mmol, 0.73 eq). This solution was left to stir at rt for 4 h, after which it was quenched with the addition of 2 mL of a 1:1 mixture of H2O:acetone. This quenched solution was diluted to 60 mL of diethyl ether and washed twice with 20 mL of 0.2 M NaH2PO4, dried over Na2SO4, filtered, and concentrated to give a white solid. This solid was purified by silica gel chromatography. The column was run with step-wise gradient elution: 5% EtOAc:95% Pet. Ether → 10% EtOAc:90% Pet. Ether → 15% EtOAc:85% Pet. Ether → 20% EtOAc:80% Pet. Ether. Compound 50 was attained as a colourless wax (40 mg, 40%). 1H NMR (400 MHz, Methanol-d4) δ 3.94 (s, 4H), 3.68 – 3.62 (m, 2H), 2.30 – 2.13 (m, 3H), 1.96 – 1.81 (m, 4H), 1.51 (s, 18H). 13C NMR (101 MHz, MeOD) δ 153.83, 110.39, 84.64, 83.74, 69.36, 66.05, 43.28, 37.30, 36.74, 28.31, 13.70. HRMS (ESI): m/z calc for C19H31NO6 [M+Na]+ 392.2053, found 392.2047. 178 Compound 51 Compound 51: To compound 40 (722 mg, 4.3 mmol) was added HOTs•H2O (352 mg, 1.9 mmol, 0.43 eq) in 15 mL of MeOH and 1 mL of Trimethyl orthoformate. This solution was left to stir under Argon at rt for 70 h. The solution was then quenched with the addition of 2 mL of NH4OH and 10 mL of EtOAc and concentrated under reduced pressure. The resultant wax was diluted to 60 mL of diethyl ether and washed twice with 20 mL of H2O, dried over Na2SO4, filtered, and concentrated to give a light orange oil. This oil was diluted to 40 mL of THF and LAH (398 mg, 10.5 mmol, 2.44 eq) was added. This solution was left to stir under Argon at rt for 2 h, after which it was quenched, filtered, and concentrated as per the typical Fieser workup procedure to give a light orange oil. This oil was purified by silica gel chromatography. The column was run with step-wise gradient elution: 10% EtOAc:90% Pet. Ether → 20% EtOAc:80% Pet. Ether → 30% EtOAc:70% Pet. Ether. Compound 51 was attained as a colourless liquid (576 mg, 78%). 1H NMR (400 MHz, Methanol-d4) δ 3.58 (t, J = 7.3 Hz, 2H), 3.16 (s, 6H), 2.19 (ddt, J = 16.3, 6.3, 2.7 Hz, 3H), 1.85 (q, J = 7.3 Hz, 4H). 13C NMR (101 MHz, MeOD) δ 102.74, 84.44, 69.44, 58.55, 48.05, 35.97, 33.56, 14.00. HRMS (ESI): m/z calc for C9H16O3 [M+Na]+ 195.0999, found 195.0993. 179 Compound 52 To compound 51 (102 mg, 0.59 mmol) was added PPh3 (522 mg, 2.0 mmol, 3.4 eq), I2 (530 mg, 2.1 mmol, 3.5 eq), and imidazole (383 mg, 5.6 mmol, 9.5 eq) in 4 mL of DCM. This solution was left to stir at rt under light shielding for 26 h. It was then diluted to 60 mL of diethyl ether and washed twice with 20 mL of bleach, twice with 20 mL of sat Na2S2O3, dried over Na2SO4 with some NaHCO3, filtered, and concentrated to partial dryness with NaHCO3 present – care must be taken to keep the alkyl iodide intermediate in a mildly basic environment at all times. To the resultant light-yellow liquid was added NaN3 (342 mg, 5.3 mmol, 9.0 eq) and 3 mL of degassed DMF. This solution was left to stir at rt under light shielding for 42 h. It was then diluted to 60 mL of diethyl ether and washed twice with 20 mL H2O, dried over Na2SO4, filtered, and concentrated to give a light-yellow wax. This wax was purified by silica gel chromatography. The column was run with step-wise gradient elution: 2% EtOAc:98% Pet. Ether → 5% EtOAc:95% Pet. Ether → 10% EtOAc:90% Pet. Ether. Compound 52 was attained as a light-yellow liquid (71 mg, 61%). Alternatively, to compound 51 (198 mg, 1.15 mmol) was added MsCl (0.26 mL, 3.4 mmol, 2.9 eq), NEt3 (0.8 mL, 5.8 mmol, 3.8 eq) in 3 mL of DCM. This solution was left to stir at rt 1.5 h. It was then diluted to 60 mL of diethyl ether and washed twice with 20 mL H2O, once with 10 mL of brine, dried over Na2SO4 with some NaHCO3, filtered, and concentrated to partial dryness with NaHCO3 present – care must be taken to keep the alkyl mesylate intermediate in a mildly basic environment at all times. To the resultant light-yellow liquid was added NaN3 (558 mg, 8.6 mmol, 180 7.5 eq), KI (603 mg, 3.6 mmol, 3.2 eq) and 4 mL of degassed DMF. This solution was left to stir at rt under light shielding for 36 h. It was then diluted to 60 mL of diethyl ether and washed twice with 20 mL H2O, dried over Na2SO4, filtered, and concentrated to give a light-yellow wax. This wax was purified by silica gel chromatography. The column was run with step-wise gradient elution: 2% EtOAc:98% Pet. Ether → 5% EtOAc:95% Pet. Ether → 10% EtOAc:90% Pet. Ether. Compound 52 was attained as a light-yellow liquid (76 mg, 34%). 1H NMR (400 MHz, Methanol-d4) δ 3.42 – 3.34 (m, 2H), 3.20 (s, 6H), 2.29 (t, J = 2.7 Hz, 1H), 2.24 (td, J = 8.2, 7.7, 2.7 Hz, 2H), 2.00 – 1.93 (m, 2H), 1.93 – 1.87 (m, 2H). 13C NMR (101 MHz, MeOD) δ 100.49, 82.40, 67.71, 46.19, 45.86, 31.05, 30.45, 11.85. HRMS (ESI): m/z calc for C9H15N3O2 [M+H]+ 198.1160, found 198.1232. 181 Compound 53 To compound 52 (71 mg, 0.36 mmol) was added PPh3 (221 mg, 0.84 mmol, 2.3 eq) in 5 mL of THF and 0.5 mL of distilled H2O. This solution was left to stir at rt for 24 h, after which it was concentrated under reduced pressure. To the resultant solid was added 5 mL of THF, 0.5 mL of distilled NEt3, and Boc2O (618 mg, 2.83 mmol, 7.9 eq). This solution was left to stir at rt for 8 h, after which it was quenched with the addition of 2 mL of a 1:1 mixture of H2O:acetone and DMAP (41 mg, 0.34 mmol, 0.9 eq). This quenched solution was diluted to 60 mL of diethyl ether and washed twice with 20 mL of 0.2 M NaH2PO4, dried over Na2SO4, filtered, and concentrated to give a white solid. This solid was purified by silica gel chromatography. The column was run with step-wise gradient elution: 5% EtOAc:95% Pet. Ether → 10% EtOAc:90% Pet. Ether → 15% EtOAc:85% Pet. Ether → 20% EtOAc:80% Pet. Ether. Compound 53 was attained as a colourless wax (80 mg, 82%). 1H NMR (400 MHz, Methanol-d4) δ 3.16 (s, 6H), 3.07 – 2.99 (m, 2H), 2.23 (t, J = 2.6 Hz, 1H), 2.21 – 2.12 (m, 2H), 1.88 – 1.80 (m, 2H), 1.80 – 1.73 (m, 2H), 1.43 (s, 9H). 13C NMR (101 MHz, MeOD) δ 158.39, 102.71, 84.39, 79.95, 69.55, 36.79, 33.28, 33.12, 28.78, 28.74, 13.92. HRMS (ESI): m/z calc for C14H25NO4 [M+Na]+ 294.1683, found 294.1677. 182 Compound 54 To compound 52 (50 mg, 0.25 mmol) was added PPh3 (164 mg, 0.63 mmol, 2.5 eq) in 5 mL of THF and 0.5 mL of distilled H2O. This solution was left to stir at rt for 26 h, after which it was concentrated under reduced pressure. To the resultant solid was added 10 mL of THF, 0.5 mL of distilled NEt3, Boc2O (521 mg, 2.29 mmol, 9.6 eq), and DMAP (31 mg, 0.25 mmol, 1 eq). This solution was left to stir at rt for 14 h, after which it was quenched with the addition of 2 mL of a 1:1 mixture of H2O:acetone. This quenched solution was diluted to 60 mL of diethyl ether and washed twice with 20 mL of 0.2 M NaH2PO4, dried over Na2SO4, filtered, and concentrated to give a white solid. This solid was purified by silica gel chromatography. The column was run with step-wise gradient elution: pure Pet. Ether → 5% EtOAc:95% Pet. Ether → 10% EtOAc:90% Pet. Ether. Compound 54 was attained as a colourless wax (78 mg, 83%). 1H NMR (400 MHz, Methanol-d4) δ 3.61 – 3.53 (m, 2H), 3.18 (s, 6H), 2.21 (m, 3H), 1.89 – 1.80 (m, 4H), 1.52 (s, 18H). 13C NMR (101 MHz, MeOD) δ 153.96, 102.40, 84.14, 83.94, 69.71, 42.97, 33.24, 32.38, 28.35, 14.05. HRMS (ESI): m/z calc for C19H33NO6 [M+Na]+ 394.2209, found 394.2204. 183 Compound 55 Compound 46 (75 mg, 0.33 mmol) was diluted to 5 mL of 7 M NH3 in MeOH and left to stir under Argon and light shielding at rt for 24 h. Freshly-made t-butyl hypochlorite 206 (387 mg, 3.6 mmol, 10.8 eq) dissolved in 0.5 mL of t-butyl alcohol was added under ice cooling. This mixture was left to stir under Argon and light shielding at rt for 24 h. Thereafter, more t-butyl hypochlorite (368 mg, 3.4 mmol, 10.3 eq) was added to the mixture under ice cooling. This solution was left to stir under Argon and light shielding at rt a further 20 h. Thereafter, this solution was diluted to 60 mL of diethyl ether and washed twice with 20 mL of sat Na2S2O3, washed twice with 20 mL of 0.2 M NaH2PO4, dried over Na2SO4, filtered, and concentrated to give a light yellow oil. This solid was purified by silica gel chromatography. The column was run with step-wise gradient elution: pure Pet. Ether → 2% EtOAc:98% Pet. Ether → 5% EtOAc:95% Pet. Ether→ 7% EtOAc:93% Pet. Ether → 10% EtOAc:90% Pet. Ether. Compound 55 was attained as a colourless oil (31 mg, 39%) and was surprisingly non-polar relative to the starting ketone 46. 1H NMR (400 MHz, Methanol-d4) δ 2.94 (t, J = 7.1 Hz, 2H), 2.27 (t, J = 2.6 Hz, 1H), 2.02 (td, J = 7.5, 2.6 Hz, 2H), 1.62 (t, J = 7.5 Hz, 2H), 1.56 (t, J = 7.1 Hz, 2H), 1.44 (s, 9H). 13C NMR (101 MHz, MeOD) δ 158.26, 83.59, 80.10, 70.28, 36.26, 34.00, 33.39, 28.76, 27.79, 13.84. HRMS (ESI): m/z calc for C12H19N3O2 [M+H]+ 238.1474, found 238.1546. 184 Compound 56 Compound 47 (15 mg, 0.046 mmol) was diluted to 2 mL of 7 M NH3 in MeOH and left to stir under Argon and light shielding at rt for 22 h. Freshly-made t-butyl hypochlorite 206 (52 mg, 0.48 mmol, 10.4 eq) dissolved in 0.5 mL of t-butyl alcohol was added under ice cooling. This mixture was left to stir under Argon and light shielding at rt for 20 h. Thereafter, more t-butyl hypochlorite (62 mg, 0.57 mmol, 12.4 eq) was added to the mixture under ice cooling. This solution was left to stir under Argon and light shielding at rt a further 16 h. Thereafter, this solution was diluted to 60 mL of diethyl ether and washed twice with 20 mL of sat Na2S2O3, washed twice with 20 mL of 0.2 M NaH2PO4, dried over Na2SO4, filtered, and concentrated to give a light-yellow oil. This solid was purified by silica gel chromatography. The column was run with step-wise gradient elution: pure Pet. Ether → 1% EtOAc:99% Pet. Ether → 2% EtOAc:98% Pet. Ether→ 3% EtOAc:97% Pet. Ether → 4% EtOAc:96% Pet. Ether. Compound 56 was attained as a colourless oil (2.6 mg, 17%) and was surprisingly non-polar relative to the starting ketone 47. 1H NMR (400 MHz, Methanol-d4) δ 3.65 – 3.56 (m, 2H), 2.30 (t, J = 2.6 Hz, 1H), 2.03 (td, J = 7.4, 2.6 Hz, 2H), 1.66 – 1.57 (m, 4H), 1.51 (s, 18H). 13C NMR (101 MHz, MeOD) δ 153.64, 84.00, 83.51, 70.42, 42.46, 33.79, 33.13, 28.28, 27.55, 13.85. HRMS (ESI): m/z calc for C17H27N3O4 [M+Na]+ 360.1896, found 360.1888. 185 Compound 57 Compound 23 (8 mg, 0.012 mmol) was diluted to 5 mL of MeOH and treated with 9 mg of 10% Pd/C under H2 atmosphere for 1 hr. The catalyst was filtered through Celite® on top of a fine fritted glass funnel via MeOH eluent and the filtrate was concentrated to give crude debenzylated 23 as a clear colourless wax. Meanwhile, compound 55 (5 mg, 0.021 mmol, 1.8 eq) was treated with 1 mL of 4 M HCl in dioxane at rt for 30 mins and concentrated under reduced pressure to give a clear colourless foam. This foam was diluted to 1 mL of degassed DMF and added to the debenzylated wax of 23 along with PyBOP (8 mg, 0.015 mmol, 1.3 eq), HOBt•H2O (4 mg, 0.026 mmol, 2.2 eq), and DIPEA (0.02 mL, 0.11 mmol, 9.6 eq). This solution was left to stir at rt and under light shielding for 20 h, after which it was diluted to 60 mL of DCM and washed twice with 20 mL of 1:1 sat aq NaHCO3:10% w/w aq LiCl, twice with 20 mL of 0.2 M NaH2PO4, dried over Na2SO4, filtered, and concentrated. The resultant yellow foam was loaded with distilled H2O onto a 2 g reverse-phase C18 Sepack column. The column was run with step-wise gradient elution: distilled H2O → 10% MeCN:90% distilled H2O → 20% MeCN:80% distilled H2O. The latter 20% MeCN:80% distilled H2O fractions were harvested and concentrated. The resultant white foam was further purified by silica gel chromatography. The column was run with step-wise gradient elution: 4% MeOH:96% DCM → 5% MeOH:95% DCM → 6% MeOH:94% DCM → 186 7% MeOH:93% DCM → 8% MeOH:92% DCM. Compound 57 was attained as a clear colourless film (4 mg, 49%). 1H NMR (600 MHz, Methanol-d4) δ 5.33 (s, 1H), 4.61 (d, J = 5.1 Hz, 1H), 4.55 (d, J = 8.4 Hz, 1H), 4.35 (q, J = 7.1 Hz, 1H), 4.30 (d, J = 7.4 Hz, 1H), 4.08 (q, J = 6.7 Hz, 1H), 3.97 (s, 1H), 3.92 – 3.74 (m, 6H), 3.59 – 3.55 (m, 2H), 3.46 (s, 1H), 3.09 (ddt, J = 40.4, 13.8, 7.0 Hz, 2H), 2.27 (t, J = 2.6 Hz, 1H), 2.08 (s, 3H), 2.04 (s, 3H), 2.03 – 2.00 (m, 2H), 1.63 (td, J = 7.2, 2.7 Hz, 4H), 1.53 (s, 3H), 1.39 (d, J = 5.8 Hz, 6H), 1.34 (d, J = 6.7 Hz, 3H). 13C NMR (101 MHz, MeOD) δ 174.94, 174.70, 173.34, 102.01, 100.92, 83.60, 79.52, 77.11, 75.84, 75.42, 74.39, 72.48, 70.40, 68.63, 65.84, 63.04, 57.59, 50.08, 35.42, 33.34, 33.21, 29.42, 27.86, 23.45, 22.59, 19.27, 18.52, 18.16, 13.84, 13.82. HRMS (ESI): m/z calc for C32H48N6O12 [M+Na]+ 731.3227, found 731.3221. 187 Compound 58 To compound 22 (21 mg, 0.032 mmol) was added butyric anhydride (220 mg, 1.39 mmol, 43.5 eq), DMAP (9 mg, 0.07 mg, 2.3 eq) and 2 mL of pyridine. This solution was left to stir at rt under Argon atmosphere for 5 h. The solution was quenched with 4 mL of MeOH and concentrated under reduced pressure to give a yellow wax. This wax was purified by silica gel chromatography. The column was run with stepwise gradient elution: pure DCM → 99% DCM:1% MeOH → 98% DCM:2% MeOH → 97% DCM:3% MeOH → 96% DCM:4% MeOH → 95% DCM:5% MeOH. Compound 58 was attained as a clear colourless wax (19 mg, 68%). 188 1H NMR (600 MHz, Methanol-d4) δ 7.34 (dd, J = 23.6, 4.4 Hz, 5H), 5.33 (s, 1H), 5.25 – 5.21 (m, 1H), 5.21 – 5.11 (m, 2H), 5.05 (t, J = 9.7 Hz, 1H), 4.71 (d, J = 8.4 Hz, 1H), 4.59 (d, J = 5.3 Hz, 1H), 4.51 (q, J = 7.3 Hz, 1H), 4.26 (dd, J = 12.4, 4.2 Hz, 1H), 4.21 (d, J = 7.5 Hz, 2H), 4.12 (q, J = 6.7 Hz, 1H), 4.02 (dd, J = 10.8, 8.5 Hz, 1H), 3.97 (s, 1H), 3.83 (ddd, J = 10.1, 4.2, 2.3 Hz, 1H), 3.80 (s, 1H), 3.72 – 3.68 (m, 1H), 3.55 (s, 1H), 2.36 – 2.20 (m, 7H), 2.05 (s, 3H), 1.96 (s, 3H), 1.67 – 1.55 (m, 6H), 1.44 (d, J = 7.3 Hz, 3H), 1.34 (d, J = 6.8 Hz, 3H), 0.95 (t, J = 7.4 Hz, 3H), 0.92 (t, J = 7.4 Hz, 6H). 13C NMR (101 MHz, MeOD) δ 174.91, 174.68, 174.47, 174.27, 173.71, 173.55, 173.18, 137.23, 129.58, 129.34, 129.21, 102.00, 101.40, 79.17, 77.17, 76.45, 74.73, 73.17, 73.14, 69.64, 67.99, 65.78, 62.75, 55.20, 49.36, 36.86, 36.76, 23.21, 22.62, 19.44, 19.33, 19.24, 18.30, 17.68, 14.01, 13.93, 13.87. 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Carbohydrate Research 2001, 336 (1), 13–46. https://doi.org/10.1016/S0008-6215(01)00213-0. 204 Appendix: NMR Data Compound 2 – 1H NMR (D2O with an internal standard of CH3OH, 600 MHz) Compound 2 – 13C NMR (D2O with an internal standard of CH3OH, 101 MHz) 205 Compound 3 – 1H NMR (MeOD, 600 MHz) Compound 3 – 13C NMR (MeOD, 101 MHz) 206 Compound 4 – 13C NMR (D2O with an internal standard of CH3OH, 600 MHz) Compound 4 – 13C NMR (D2O with an internal standard of CH3OH, 101 MHz) 207 Compound 5 – 1H NMR (D2O with an internal standard of CH3OH, 600 MHz) Compound 5 – 13C NMR (D2O with an internal standard of CH3OH, 151 MHz) 208 Compound 6 – 1H NMR (MeOD, 600 MHz) Compound 6 – 13C NMR (MeOD, 101 MHz) 209 Compound 7 – 1H NMR (MeOD, 600 MHz) Compound 7 – 13C NMR (MeOD, 101 MHz) 210 Compound 21 – 1H NMR (MeOD, 600 MHz) Compound 21 – 13C NMR (MeOD, 101 MHz) Amide N-H proton slow to undergo Deuterium exchange 211 Compound 22 – 1H NMR (MeOD, 600 MHz) Compound 22 – 13C NMR (MeOD, 101 MHz) 212 Compound 23 – 1H NMR (MeOD, 600 MHz) Compound 23 – 13C NMR (MeOD, 101 MHz) 213 Compound 24 – 1H NMR (MeOD, 600 MHz) Compound 24 – 13C NMR (MeOD, 101 MHz) 214 Compound 25 – 1H NMR (MeOD, 600 MHz) Compound 25 – 13C NMR (MeOD, 101 MHz) 215 Compound 26 – 1H NMR (MeOD, 600 MHz) Compound 26 – 13C NMR (MeOD, 101 MHz) 216 Compound 27 – 1H NMR (MeOD, 600 MHz) Compound 27 – 1H NMR (MeOD, 151 MHz) 217 Compound 28 – 1H NMR (MeOD, 600 MHz) Compound 28 – 13C NMR (MeOD, 101 MHz) 218 Compound 29 – 1H NMR (MeOD, 600 MHz) Compound 29 – 13C NMR (MeOD, 101 MHz) 219 Compound 30 – 1H NMR (MeOD, 600 MHz) Compound 30 – 13C NMR (MeOD, 101 MHz) 220 Compound 31 – 1H NMR (MeOD, 600 MHz) Compound 31 – 13C NMR (CDCl3, 101 MHz) 221 Compound 36 – 1H NMR (CDCl3, 400 MHz) Compound 36 – 13C NMR (D6-acetone, 101 MHz) 222 Compound 37 – 1H NMR (D6-acetone, 600 MHz) Compound 37 – 13C NMR (D6-acetone, 101 MHz) 223 Compound 38 – 1H NMR (D6-acetone, 600 MHz) Compound 38 – 13C NMR (D6-acetone, 101 MHz) 224 Compound 39 – 1H NMR (MeOD, 600 MHz) Compound 39 – 13C NMR (MeOD, 101 MHz) 225 Compound 45 – 1H NMR (MeOD, 400 MHz) Compound 45 – 13C NMR (MeOD, 101 MHz) 226 Compound 46 – 1H NMR (MeOD, 400 MHz) Compound 46 – 13C NMR (MeOD, 101 MHz) 227 Compound 47 – 1H NMR (MeOD, 400 MHz) Compound 47 – 13C NMR (MeOD, 101 MHz) 228 Compound 48 – 1H NMR (MeOD, 400 MHz) Compound 48 – 13C NMR (MeOD, 101 MHz) 229 Compound 49 – 1H NMR (MeOD, 400 MHz) Compound 49 – 13C NMR (MeOD, 101 MHz) 230 Compound 50 – 1H NMR (MeOD, 400 MHz) Compound 50 – 13C NMR (MeOD, 101 MHz) 231 Compound 51 – 1H NMR (MeOD, 400 MHz) Compound 51 – 13C NMR (MeOD, 101 MHz) 232 Compound 52 – 1H NMR (MeOD, 400 MHz) Compound 52 – 13C NMR (MeOD, 101 MHz) 233 Compound 53 – 1H NMR (MeOD, 400 MHz) Compound 53 – 13C NMR (MeOD, 101 MHz) 234 Compound 54 – 1H NMR (MeOD, 400 MHz) Compound 54 – 13C NMR (MeOD, 101 MHz) 235 Compound 55 – 1H NMR (MeOD, 400 MHz) Compound 55 – 13C NMR (MeOD, 101 MHz) 236 Compound 56 – 1H NMR (MeOD, 400 MHz) Compound 56 – 13C NMR (MeOD, 101 MHz) 237 Compound 57 – 1H NMR (MeOD, 600 MHz) Compound 57 – 13C NMR (MeOD, 101 MHz) 238 Compound 58 – 1H NMR (MeOD, 600 MHz) Compound 58 – 13C NMR (MeOD, 101 MHz)
Thesis/Dissertation
eng
Chemistry
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Tanner, Martin E.
Synthesis of tools for studying the AmpG pore protein involved in antibiotic resistance
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