Developing new fluorophores for applications in protease detection and protein labeling

DEVELOPING NEW FLUOROPHORES FOR APPLICATIONS IN PROTEASE DETECTION AND PROTEIN LABELING LI JUNQI NATIONAL UNIVERSITY OF SINGAPORE 2010 DEVELOPING NEW FLUOROPHORES FOR APPLICATIONS IN PROTEASE DETECTION AND PROTEIN LABELING LI JUNQI A THESIS SUBMITTED FOR THE DEGREE OF MASTER OF SCIENCE DEPARTMENT OF CHEMISTRY NATIONAL UNIVERSITY OF SINGAPORE 2010 ACKNOWLEGEMENTS This thesis is not the result of a sole experimenter working in isolation, but the culmination of efforts of all who have supported the individual in her search for greater knowledge. The journey as a graduate student in NUS may have ended, but it is the beginning of a path leading to a boundless world of scientific pursuits. My utmost gratitute to the following people who have made it possible: Prof Yao Shao Qin – supervisor, mentor, teacher and a friend in need – has been instrumental in shaping my development both as a scientist and as an individual. The years spent under his tutelage have had the most profound impact on my life as a student of science. It is with his enthusiasm and insight in scientific research, as well as confidence in my abilities that have led me to my accomplishments. My parents and my brother have been a silent pillar of support, showing their care and concern in their own ways even when I hardly spent time with them throughout the course of this degree. I can only reciprocate their love by dedicating to them every small accomplishment I make, including this thesis. The members of the Yao Lab, both past and present, have guided and accompanied me throughout these years. I thank particularly the following people: Jinzhan, Souvik and Candy for being great friends who shared my frustrations; Mingyu and Jingyan who have been great companions in the lab; and Mahesh and Wang Jun who have mentored me when I was learning the ropes of research. i I thank my old pals Aileen, Ke Ming and Zhiying who have not forgotten me during the time I disappeared into the lab. It is certainly comforting to know that our friendship has weathered these years. Last but not least, I thank National University of Singapore for funding my studies through the research scholarship, and the President’s Graduate Fellowship. ii TABLE OF CONTENTS Acknowledgements i Table of Contents iii List of Figures vii List of Schemes ix List of Tables x Index of Abbreviations xi List of Amino Acids xiv List of Publications xv Abstract xvii Chapter 1 INTRODUCTION 1.1 Detecting Enzyme Activity 1 1.2 Small Molecule-Based Fluorogenic Enzyme Substrates Chapter 2 1.2.1 FRET and Internally Quenched Substrates 3 1.2.2 Fluorophore Release after Enzymatic Cleavage 6 1.2.3 Fluoromorphic Probes 11 1.2.4 Fluorescence Detection of Binding Events 12 DEVELOPING A NEW FLUOROGENIC PROBE FOR PROTEASE ACTIVITY 2.1 Fluorogenic Protease Substrates for Detecting Protease Activity 15 on the Microarray and in Live Cells 2.2 Design of a New Fluorophore for Microarray and Bioimaging iii 18 Applications 2.3 Chemical Synthesis of SG and SG-Conjugated Peptides 20 2.4 Profiling Protease Activity on the Microarray 32 2.5 Imaging Caspase-3 and -7 Activities in Live Cells 38 2.6 Conclusions 39 Chapter 3 FLUOROGENIC PROBES FOR DETECTING PROTEASE ACTIVITY AT SUBCELLULAR LOCATIONS 3.1 Targeted Delivery of Molecules into Intracellular Locations 41 3.2 Design of Cell-Permeable Protease Substrates Targeting 46 Different Organelles 3.3 Chemical Synthesis of Peptide Substrates and Localization 51 Peptides 3.4 Bioimaging of Control Peptides 61 3.5 Current Work 65 Chapter 4 DISCOVERY AND DEVELOPMENT OF FLUOROGENIC LABELS FOR BIOMOLECULES 4.1 Fluorogenic Labeling of Biomolecules 69 4.2 Combinatorial Discovery of Fluorophores 74 4.3 Design of Xanthone- and Xanthene-Based Fluorophores 76 4.4 Chemical Synthesis of Xanthone- and Xanthene-based “Click” 77 Fluorophores 4.5 Spectroscopic Analysis of the “Click” Fluorophore Library iv 87 4.6 Conclusions 97 Chapter 5 EXPERIMENTAL SECTION 5.1 General Information 98 5.2 Solution-Phase Synthesis of Fluorophores, Linkers and Azides 99 5.2.1 Synthesis of SG1 and SG2 and related derivatives 5.2.2 Synthesis of alkynes A – F 111 5.2.3 Synthesis of Linkers 123 5.2.4 Synthesis of Azides 128 5.3 Solid-Phase Synthesis of Peptides and SG-Peptide Conjugates 99 132 5.3.1 General Information 128 5.3.2 General Procedures 128 5.3.3 Synthesis of Ac-DEVD-SG1 129 5.3.4 Synthesis of SG2-Peptide Conjugates 130 5.3.5 Synthesis of Alkyne-Functionalized SG2-Based 132 Substrates 5.3.6 Synthesis of Azido-Peptides and Control Peptides 133 5.4 Synthesis of Fluorophores Using “Click” chemistry 135 5.5 Spectroscopic Analysis 140 5.5.1 General Information 140 5.5.2 Determination of Molar Extinction Coefficients and 140 Quantum Yields 5.6 Microplate-Based Fluorescence Assays 141 5.6.1 General Information 141 5.6.2 Enzymatic Assays with SG-Peptide Conjugates 142 v 5.6.3 Fluorescence Analysis of “Click” Fluorophores 142 5.7 Microarray Experiments 143 5.8 Bioimaging 145 5.8.1 General Information 145 5.8.2 Detecting Caspase-3 and -7 Activity in Live HeLa 146 Cells 5.8.3 Evaluating the subcellular locations of the localization 146 peptides 148 Chapter 7 REFERENCES vi LIST OF FIGURES Figure Page 1.1 Enzyme assays with fluorescence detection methods 3 2.1a Protease and protease substrate nomenclature 16 2.1b 2 common types of synthetic peptide substrates 16 2.2 Structures of common fluorophores used in fluorogenic peptide 19 substrates 2.3 Resonance stabilization of phenolate anion resulting from TBS 21 deprotection 2.4 The 2 major resonance structures of the asymmetric xanthene 21 2.5 Formation of the undesired N-acylurea from Fmoc-Asp-SG1 24 and DIC 2.6 LC-MS profile of Ac-DEVD-SG1 25 2.7 LC-MS profiles of the 10 SG2-peptide conjugates 29 2.8a Fluorescence spectra of SG1 33 2.8b Fluorescence increase from cleavage of Ac-DEVD-SG1 33 2.9 Detecting protease activity on the microarray 34 2.10a Enzyme “fingerprints” obtained 36 2.10b Time-dependent kinetic profiles from microarray 36 2.11 Selected kinetic data from microplate and microarray 37 enzymatic assays 2.12 Detecting caspase-3/-7 activity in live HeLa cells with Ac- 39 DEVD-SG1 3.1 Overall strategy for imaging protease activity in subcellular vii 47 organelles 3.2 Acylation of resin-bound secondary amine by Fmoc-SG2- 52 COOH and possible side reaction 3.3 General structures and LC-MS profiles of desired peptides and 54 side products 3.4 LC-MS profiles of azido-localization peptides and control 57 peptides 3.5 Fluorescent images of control peptides and corresponding 63 organelle stains 4.1 Fluorophore types which have been synthesized using “click” 75 chemistry 4.2 Design of xanthone- and xanthene-based “click” fluorophores 77 4.3 Undesired products obtained during the nucleophilic aromatic 79 substitution of 2b and 1ii with different nucleophiles 4.4 Structures of azides used in this study 81 4.5 Selected LC-MS profiles of “click” fluorophores 83 4.6 Emission spectra of selected fluorophores from microplate- 89 based fluorescence screening 4.7 Heat map showing fluorescence intensities of each “click” 91 product 4.8 Structures of fluorophores selected for quantitative fluorescence 93 analysis 4.9 Excitation and emission spectra of “hit” fluoropohores and their corresponding alkynes viii 94 LIST OF SCHEMES Scheme Page 2.1 Initial proposed synthesis of SG 20 2.2 Synthesis of SG1 and SG2 22 2.3 Derivatization of SG1 and solid-phase synthesis of Ac-DEVD- 25 SG1 2.4 Synthesis of Fmoc-SG2-CHO for peptide synthesis 26 2.5 Solid-phase synthesis of aldehyde-functionalized SG2-peptide 28 conjugates 2.6 Functionalization of glass slides with alkyoxyamines 35 3.1 Synthesis of Fmoc-SG2-COOH (3-1) and Fmoc-SG2-COCl (3- 51 2) 3.2 Solid-phase synthesis of alkyne-functionalized substrates, Ac- 53 X-SG2-alkyne 4.1 General synthetic strategy towards alkynes A, B, D and E 78 4.2 Synthesis of 4-2a and 4-2b from 4-1 78 4.3 Synthesis of alkynes C and F 80 4.4 Synthesis of aromatic azides from anilines 81 4.5 “Click” assembly of fluorophores 82 5.1 Synthesis of linker 2-12 used in the preparation of SG2 124 5.2 Synthesis of azide z15 127 ix LIST OF TABLES Table Page 2.1 Peptide sequences synthesized and their target proteases 35 3.1 Alkyne-functionalized SG2-based substrates and their target 49 enzymes 3.2 Azide-functionalized localization peptides selected and their 50 target organelles 4.1 λex and λem for each “click” fluorophores 88 4.2 Summary of spectroscopic properties of “hit” fluorophores 93 5.1 Reagent concentrations and volumes used per “click” reaction 135 5.2 Volumes of solvents used for scale-up “click” chemistry 136 5.3 Concentrations and buffers for proteases used in microarray 144 experiments x INDEX OF ABBREVIATIONS ABP Activity-based probe ACC 7-Aminocoumarin-4-acetic acid AMC 7-Amino-4-methylcoumarin aq. Aqueous Boc t-Butoxy carbonyl br Broad CPP Cell-penetrating peptide dd Doublet of doublets DIC N,N′-Diisopropylcarbodiimide (as a reagent) / Differential interference contrast (in bioimaging) DIEA N,N′-Diisopropylethylamine DCE 1,2-Dichloroethane DCM Dichloromethane DMAP 4-Dimethylaminopyridine DMF Dimethylformamide DMP Dess-Martin Periodinane EA Ethyl acetate EDT 1,2-ethanedithiol equiv Equivalent ESI Electron spray ionization Et Ethyl EtOH Ethanol FLIP Fluorescence loss in photobleaching xi Fmoc 9-Fluorenylmethoxycarbonyl FP Fluorescent protein FRAP Fluorescence recovery after photobleaching g Gram GFP Green fluorescent protein HeLa Human cervical adenocarcinoma HBTU 2-(1-H-benzotriazol-1-yl)-1,1,3,3-tetrauroniumhexafluorophosphate HOBt N-hydroxybenzotriazole Hz Hertz h Hours λem Wavelength of excitation maximum λex Wavelength of emission maximum LC-MS Liquid chromatography-mass spectrometry M Molar MeOH Methanol m Multiplet mg Milligram min Minute mM Millimolar µM Micromolar mmol Millimole MMP Matrix metalloproteases NLS Nuclear localization sequences NMR Nuclear magnetic resonance nM Nanomolar xii OTf Trifluoromethane sulfonyl / Triflate OTs p-Toluenesulfonyl / Tosylate PDC Pyridinium dichromate Ph Phenyl PL-FMP Polystyrene – 4-formyl-3-methoxyphenoxy resin ppm Parts per million PTD Protein transduction domain PyBrOP Bromo-tris-pyrrolidino phosphoniumhexafluorophosphate q Quartet RFP Red fluorescent protein SG Singapore Green SMM Small molecule microarray s Singlet sat. Saturated SV40 Simian virus 40 t Triplet TBS/TBDMS tert-Butyldimethylsilyl tBuOH tert-Butyl alcohol TFA Trifluoroacetic acid THF Tetrahydrofuran TIS Triisopropylsilane TLC Thin layer chromatography TMS trimethylsilyl UV Ultraviolet xiii LIST OF AMINO ACIDS One Letter Three Letter Amino Acid A Ala Alanine C Cys Cysteine D Asp Aspartic acid E Glu Glutamic acid F Phe Phenylalanine G Gly Glycine H His Histidine I Ile Isoleucine K Lys Lysine L Leu Leucine M Met Methionine N Asn Asparagine P Pro Proline Q Gln Glutamine R Arg Arginine S Ser Serine T Thr Threonine V Val Valine W Trp Tryptophan Y Tyr Tyrosine r D-Arg D-Arginine Fx - Cyclohexylalanine xiv LIST OF PUBLICATIONS 1. Li, J.; Hu, M.; Yao, S. Q. Rapid synthesis, screening and identification of xanthone- and xanthene-based fluorophores using click chemistry. Org. Lett. 2009, 11, 3008-3011. 2. Li, J.; Yao, S. Q. “Singapore Green” – a new fluorescent dye for microarray and bioimaging applications. Org. Lett. 2009, 11, 405-408. 3. Hu, M.; Li, J.; Yao, S. Q. In situ “click” assembly of small molecule matrix metalloprotease inhibitors containing zinc-chelating groups. Org. Lett. 2008, 10, 5529-5539 4. Uttamchandani, M.; Li, J.; Sun, H.; Yao, S. Q. Activity-based profiling: new developments and directions in protein fingerprinting. Chembiochem 2008, 9, 667-675 5. Srinivasan, R.; Li, J.; Ng, S. L.; Kalesh, K. A.; Yao, S. Q. Methods of using click chemistry in the discovery of enzyme inhibitors. Nat. Protocols 2007, 2, 26652664. 6. Lee, W. L.; Li, J.; Uttamchandani, M.; Sun, H.; Yao, S. Q. Inhibitor fingerprinting of metalloproteases using microplate and microarray platforms – an enabling technology in Catalomics. Nat. Protocols 2007, 2, 2126-2138. xv 7. Uttamchandani, M.; Wang, J.; Li, J.; Hu, M.; Sun, H.; Chen, K. Y.-T.; Liu, K.; Yao, S. Q. Inhibitor fingerprinting of matrix metalloproteases using a combinatorial peptide hydroxamate library. J. Am. Chem. Soc. 2007, 129, 1311013117. 8. Wang, J.; Uttamchandani, M.; Li, J.; Hu, M.; Yao, S. Q. “Click” synthesis of small molecule probes for activity-based fingerprinting of matrix metalloproteases. Chem. Commun. 2006, 3783-3785 9. Wang, J.; Uttamchandani, M.; Li, J.; Hu, M.; Yao, S. Q. Rapid assembly of matrix metalloproteases (MMP) inhibitors using click chemistry. Org. Lett. 2006, 8, 3821-3824 xvi ABSTRACT The design and synthesis of a new bi-functional fluorophore with emission and excitation wavelengths similar to fluorescein, and the utility of the fluorophore in microarray and bioimaging applications are described herein. We demonstrate the compatilibity of the fluorophore to solid-phase peptide synthesis for the assembly of various fluorophore-peptide conjugates which are used fluorogenic substrates for detecting protease activity on the microarray and in live cells. With the objective of expanding the bioimaging applications of the fluorophore to detecting protease activity in specific organelles, we synthesized, via solid phase synthesis, peptide conjugates functionalized with an alkyne which can be attached to cellular localization sequences via “click chemistry”. The use of a single fluorophore for these applications obviates the need for re-designing and synthetic evaluation of peptide conjugates for potetntial substrate profiling on the microarray and the live-cell imaging of enzyme activity separately. Based on the scaffold of our new fluorophore, we designed and synthesized a panel of new fluorophores with emission wavelengths from blue to yellow region by the “click” reaction of alkyne-functionalized xanthones and xanthenes with various azides. Screening of these fluorophores led to the identification of “hit” fluorophores which showed a fluorescence increase upon triazole formation. These “click”activated fluorogenic dyes could potentially be used for bioconjugation and bioimaging purposes. xvii CHAPTER 1 INTRODUCTION 1.1 Detecting Enzyme Activity Enzymes – macromolecular catalysts in biological reactions – are the life force of the cell, providing it with energy and function. Numerous pathological conditions are caused by aberrant enzymatic activity, leading researchers to seek the “magic bullet” for the specific inhibition or activation for each disease-associated enzyme [1]. These enzymes constitute more than twenty percent of the drug targets [2], underscoring the importance of finding small molecule modulators with either the aim of gaining a fundamental understanding of enzyme function or with the ultimate purpose of drug discovery. The development of enabling tools that could quantitatively assess the efficacy of these modulators in a reliable fashion is thus of tantamount importance. In vitro assays for various classes of enzymes have evolved from the labor-intensive, use of liquid chromatography and radio-labeled enzyme substrates to operationally simple methods allowing high-throughput and image-based analysis. In vivo tracking of enzymatic activity has advanced rapidly from the landmark discovery and applications of the green fluorescent protein (GFP), a milestone development in molecular biology that was awarded the Nobel Prize in 2008. Assays employing fluorescence detection methods have seen widespread use in both the academics and the industry. The appeal of fluorescence methods stems from their compatibility in both in vivo and in vitro settings, as well as their suitability for both quantitative analyses for real-time monitoring of enzyme kinetics and for 1 visual tracking of enzymatic activity. The proven utility of these assays has driven active research in designing and/or modifying fluorescent proteins, inorganic nanoparticles and small molecule organic fluorophores for use in these assays. Enzyme assays with fluorescence-based detection methods are based on a common principle – the synthetic substrate containing a fluorophore or pro-fluorophore is acted upon by the enzyme which results in a significant change in the fluorescence property of the substrate. This change could be achieved with the following mechanisms: 1) fluorescence resonance energy transfer (FRET) between a donor and acceptor fluorophore and other fluorophore-fluorophore interactions leading to quenching; 2) a fluorogenic dye which displays no or low fluorescence until enzymatic action on the substrate; and 3) the use of a metal sensitive-fluorogenic dye which is fluorescent only when chelated to metals, or an environment-sensitive fluorophore which display different spectral properties in different media (Figure 1.1). A formidable arsenal of organic fluorophores that display fluorescence changes through these mechanisms has been developed. Coupled with their amenability to structural changes through chemical synthesis, organic fluorophores now constitute an important component of the fluorescent toolbox. Their versatility has led to the development of synthetic substrates for enzymes that are not readily assayed using genetically encoded biosensors assembled from fluorescent proteins. The following section surveys the strategies in designing small molecule-based fluorogenic substrates for detecting enzyme activity. 2 a) b) c) Figure 1.1. Enzyme assays with fluorescence detection methods. a) In FRET substrates, fluorescence emission is observed from the acceptor fluorophore (red) when excited at the donor excitation wavelength until enzymatic cleavage of the substrate separates the donor and acceptor. Thereafter, emission is observed at the donor emission wavelength. b) the fluorogenic substrate is not fluorescent with the enzyme recognition head is attached. Upon enzymatic cleavage which removes the recognition head, fluorescence is restored. c) Addition of a phosphate group to the substrate by a kinase allows chelation of a metal ion by the fluorophore and phosphate group. The fluorescence is enhanced by the chelation event. 3 1.2 Small Molecule-Based Fluorogenic Enzyme Substrates 1.2.1 FRET and internally quenched substrates These fluorogenic substrates have fluorophores that are quenched by the interaction with an adjacent fluorophore or a fluorescently silent acceptor. While both types of interactions result in the decrease of the parent fluorophore, quenching and fluorescence resonance energy transfer are mechanistically distinct [4]. Quenching arises from the interaction of the electron cloud of the fluorophore and the quencher, and since molecular contact falls off rapidly with distance, most quenching mechanisms are operative only at short distances. This phenomenon was utilized in the design of synthetic graft polymers for selective tumor imaging by the Weissleder group [5]. The polymer consists of poly-L-lysine, which contains Cy5.5 (a nearinfrared cyanine dye) conjugated to some of the lysine residues, with the remaining residues either bearing free amines or protected with methoxypolyethylene glycol. In the intact polymer, the cyanine dyes are held in close proximity relative to each other and are quenched. The biocompatible polymer is known to accumulate in tumor cells and is internalized by fluid-phase endocytosis. Following endocytosis, endosomal proteases such as the cathepsins which are upregulated in tumor cells rapidly cleave the polymer by virtue of enzymatic recognition of the free lysine residues. Upon cleavage, the polymer backbone disintegrates and the Cy5.5 dyes are separated spatially. The static quenching is disengaged and the tumors are illuminated with the resultant fluorescence. This enzyme-responsive, selective tumor imaging probe was also successful in the in vivo imaging of matrix metalloprotease 2 (MMP2) - secreting tumor cells by modification of the polymer side chain to include an MMP2 substrate 4 [6]. More significantly, the fluorogenic polymer was used to assess the in vivo MMP inhibition of known inhibitors by directly detecting MMP activity in tumors. The work by Weissleder and co-workers is considered an important advance in clinical molecular imaging and set the stage for developing similar imaging strategies and techniques targeting other enzymes. In contrast to quenching, fluorescence resonance energy transfer (FRET) is a result of long range dipole-dipole interaction between the donor and acceptor, resulting in the excess energy from the excited donor fluorophore being transferred to an acceptor in the ground state without emission of a photon during the transfer. The transfer efficiency is dependent on the distance between the donor and acceptor, the extent of overlap of the donor emission spectrum and the acceptor absorption spectrum, and the relative orientation between the donor and acceptor. FRET is usually efficient up to 100 Å between the donor and acceptor. The acceptor may or may not be fluorescent. The use of a fluorescent acceptor results in a construct that absorbs at the donor excitation wavelength and emits at the acceptor wavelength when the two fluorophores are in close proximity, enabling a ratiometric fluorescence response to the distance separating the fluorophores. While enzyme substrates utilizing fluorescent donors and acceptors are typically not termed as fluorogenic substrates, enzymatic action does result in a fluorescence change in both the donor and acceptor emission wavelengths. If a non-fluorescent acceptor is used (“dark quencher”), the substrate is optically silent until an enzymatic event causes the departure of the quencher from the fluorophore, giving rise to a fluorescence increase. This class of substrates have emerged to become the most widely used and versatile in design among the different classes of enzymatic substrates used. 5 The first FRET substrate which was developed by Matayoshi and co-workers targeted the human immunodeficiency virus-1 (HIV-1) protease [7]. The FRET substrate, (DABCYL)-SQNYPIVQ-(EDANS), contains the 8-amino acid peptide sequence that is known to be cleaved by the HIV-1 protease, and a fluorophore EDANS which is quenched by the dark quencher DABCYL. Upon cleavage by the protease, the fluorophore is separated from the quencher, providing a direct read-out of enzymatic activity which could be monitored in a real-time fashion. This seminal work establishes a general design of fluorogenic substrates for other proteases, many of them are commercially available. Recent developments have focused on the use of FRET for the design of nonpeptidic, small molecule-based substrates. One of the first small molecule-based FRET substrate was designed for β-lactamases by Tsien and co-workers, with the aim of using enzymatically-amplified fluorescence readout for gene expression [8]. Mammalian cells which were stably transfected with the TEM-1 β-lactamase gene regulated by a promoter rapidly gave blue fluorescence from the β-lactamasecatalyzed hydrolysis of the FRET substrate when the promoter was added which led to upregulated gene expression. It was found that the fluorescence intensities correlated well with the number of β-lactamases expressed per cell, which could enable quantification of the readout. The group also showed that this β-lactamase reporter system could also be used for flow cytometry in engineering cell lines with targeted patterns of gene expression, and for screening drug candidates which affect gene expression. 6 Another important contribution to the use of small molecule-based FRET substrates in biological systems can be attributed to the groups of Farber and Pack, who synthesized internally quenched phospholipids as substrates for phospholipase A2 (PLA2) to assay lipid metabolism in living zebrafish larvae [9]. Ingestion of the PLA2 substrate results in cleavage by endogeneous phospholipases and accumulation of the fluorescent products in the gall bladder. Mutants that have severely impaired phospholipid processing were not fluorescent, thus enabling the researchers to generate, efficiently screen and identify genes that are critical in vertebrate digestive physiology. Further to the two examples highlighted, different groups have improved on or developed other small molecule-based FRET substrates for β-lactamases [10], other phospholipases [11] and proteases [12] for different applications with one of the following aims: enabling near-infrared or ratiometric imaging, or improving the selective detection of the target enzyme over others. It is important to note that these small molecule-based substrates are extremely useful for assaying small molecule metabolism, since there are no genetically encoded substrates that are traditionally used for other enzymes, such as proteases. 1.2.2 Fluorophore release after enzymatic cleavage Many fluorophores, including the coumarins, fluoresceins and rhodamines are characterized by an electron-donating aniline or phenol where the lone pair on the heteroatom is in conjugation with an extended π system. Reducing the lone pair availability for conjugation through acylation or phosphorylation of phenols at the 7 heteroatom dramatically reduces the fluorescence quantum yield and also leads to shifts in the wavelength of maximum absorption. In the case of phenols, alkylation of the hydroxyl group also leads to decreased fluorescence because it is the anionic phenolate form that is highly fluorescent. Enzymatic deacylation or dephosphorylation thus has the reverse effect of “turning on” the fluorescence. This unique property governs the design of fluorogenic substrates for hydrolytic enzymes: a known enzyme substrate, alternatively termed as the enzyme recognition head, is conjugated to the aniline or phenol moiety of the fluorophore which is released only upon enzymatic hydrolysis of the substrate-fluorophore bond. Fluorogenic substrates adopting this design are perhaps the earliest fluorescent assays developed for proteases and phosphatases, their subsequent development and have since branched into two major applications: i) high-throughput screening and ii) bioimaging. Fluorescent assays employing coumarin-based substrates in high-throughput screening for profiling proteases came into the spotlight with the work published by the groups of Thornberry, Ellman and Craik. Thornberry and co-workers constructed a combinatorial positional-scanning library of coumarin-linked fluorogenic peptide substrates suited for probing the P2-P4 amino acid preferences of caspases, keeping the P1 position constant as aspartic acid [13]. The Ellman and Craik groups collectively devised a practical synthesis of a coumarin derivative, 7-amino-4carbamoylmethylcoumarin (ACC), and synthetic methods for including 20 proteinogenic amino acids in the P1 position, as well as the solid-phase synthesis of fluorogenic peptide libraries [14]. The ease at which large libraries could be generated enabled the same researchers to profile a diverse array of serine proteases. The screening platform thus established by these groups has since become a reliable tool 8 for probing protease substrate preferences and generating a “fingerprint” for each protease under study, thereby allowing the differentiation of closely-related enzyme. The Ellman group then took a step in the direction of assay miniaturization for highthroughput screening with large libraries. Leveraging on the ease at which these libraries can be synthesized, a fluorogenic substrate microarray was fabricated by immobilizing the individual hydroxylamine-tagged peptides in a spatially segregated fashion onto an aldehyde-functionalized glass slide via oxime formation [15]. In this work, the researchers probed the substrate specificity of the serine protease thrombin by examining its preferences for individual peptides on a miniaturized fluorogenic assay. This had the potential of generating a proteolytic fingerprint of each protease rapidly, using minimal amounts of enzymes and substrates, in a single experiment. At the same time, our group independently prepared a complementary microarray platform for the detection of proteases and other hydrolytic enzymes, such as alkaline phosphatases, epoxide hydrolases and acetylcholine esterase, thereby demonstrating the generality of the microarray approach for detecting enzyme activity [16]. Prior to microarray-based studies for phosphatises, traditional assays for certain classes of phosphatases (PTPs) typically use phosphorylated coumarins as fluorogenic enzyme substrates. In particular 6,8-difluoro-4-methylumbelliferyl phosphate (DiFMUP) is now routinely used as a general probe for phosphatase activity. The development of specific probes for a target phosphatase using these fluorogenic substrates however remain a formidable challenge due to the need for incorporation of a bulky fluorophore into the peptide substrate without affecting phosphatase binding and activity. Barrios et. al. modified a common phosphatase substrate, 4-methylumbelliferyl phosphate, into an amino acid which served as a 9 phosphotyrosine mimic [17]. Incorporation of the unnatural amino acid into a peptide substrate of the target tyrosine phosphatase thus furnishes a fluorogenic probe for the phosphatase of interest. This simple yet elegant approach holds some promise for profiling phosphatase activity using a combinatorial peptide library, analogous to protease ‘fingerprinting’ with fluorogenic peptide libraries [18]. Direct conjugation of the enzyme recognition head to the fluorophore limits the type of chemical functionality and consequently the type of enzyme substrates that could be constructed. An important contribution to extending the scope of enzymes that may be assayed using coumarin-based substrates came from Reymond and coworkers. The Reymond group added a linker between the enzyme recognition head and the fluorophore; upon enzymatic action on the substrate moiety, spontaneous βelimination or periodate oxidation of the free alcohol followed by β-elimination occurs and the fluorophore is released. This strategy was successfully applied to assays for various hydrolytic enzymes, including epoxide hydrolases, transalodolases, transketolases and Baeyer-Villigerases. This allowed the researchers to differentiate closely-related enzymes via their ‘fingerprints’ using chiral, non-racemic coumarinbased substrates, a subject that has been extensively reviewed [19]. Substrates for bioimaging however, seldom utilize coumarin as the fluorophore due to its excitation wavelength in the ultraviolet region, which results in high background signals from autofluorescence and is damaging to live cells. In place of coumarin, the fluoresceins and rhodamines have proven to be suitable fluorophores in the bioimaging of caspases and galactosidases. Recently, dual-function probes for caspases have been developed to expand the utility of these substrates in clinical 10 bioimaging. This new subgroup of fluorogenic substrates are based on precedent fluorogenic probes for caspases, but include an additional tag that may be detected via another molecular imaging technique which is more effective for diagnostic clinical imaging. Mizukami et al. constructed Gd-DOTA-DEVD-AFC, where the fluorescence and 19F magnetic resonance (MR) signals from the fluorine-containing 7amino-4-trifluoromethylcoumarin (AFC) fluorophore is suppressed in the intact probe [20]. Upon enzymatic cleavage by caspases 3 or 7, the fluorophore is released. The gadolinium complex which serves to attenuate the 19 F MR signal via paramagnetic relaxation enhancement is ineffective when the fluorophore diffuses away, leading to both an increase in fluorescence and MR signal in response to enzymatic activity. In a similar vein, Xiong and co-workers incorporated a radionuclide in a precedent fluorogenic caspase probe for both fluorescence and nuclear imaging in preliminary in vivo imaging experiments [21]. Given that substrates used in high throughput screening and those that are suited for bioimaging differ in the type of fluorophore, our group probed the possibility of developing fluorogenic probes that are suited for both in vitro assays and live cell imaging. We designed and synthesized a new green-emitting fluorophore which could be used as a coumarin substitute in microplate- and microarray-based assays, and also in live-cell imaging of apoptosis [22]. This work is the subject of Chapter 2 in this thesis. 11 1.2.3 Fluoromorphic probes In contrast to assays for hydrolytic enzymes such as proteases and exoglycosidases, assays for other enzymes which catalyze other reaction types which cannot be monitored by the use of fluorescently-quenched peptide substrates or genetically encoded FP-based protein substrates. These transformations include isomerization reactions such peptidyl-prolyl cis/trans isomerases, or redox reactions of small molecule metabolites. There are no simple, intuitive guidelines for constructing fluorogenic probes for measuring activities of these enzymes. Suitable probes thus require separate design considerations tailored specifically for each enzyme. Sames and co-workers introduced the concept of ‘fluoromorphic’ molecules as part of their ongoing research in designing probes for redox enzymes. The group has successfully designed small molecule-based enzyme substrates which are structurally modified (“morphed”) by the enzyme of interest to a fluorescent product, hence the term ‘fluoromorphic’. In one of the earlier examples of these probes, Sames and coworkers desigened fluorogenic probes for monoamine oxidases (MAO) A and B which utilize a spontaneous indole formation following aerobic oxidation of the amine moiety by the MAO enzymes [23]. The indole formation switches on the fluorescence of the coumarin core, leading to a fluorescent response to enzymatic activity. The same group went on further to design probes for dehydrogenase enzymes by making use of the different electronic properties of ketone moiety in the probe and the alcohol resulting from dehydrogenase activity [24]. They demonstrated that these probes could distinguish the target enzymes in intact cells from numerous other redox 12 enzymes which could carry out the same functional group transformation. It should be noted that the success of the design and application of these probes depend largely on enzyme promiscuity; the construction of probes for highly specific enzymes which do not tolerate substrates which loosely resemble the endogeneous substrates still poses a formidable challenge. 1.2.4 Fluorescence detection of binding events Transferases such as the prenyltransferases and kinases which deliver small organic molecules such as lipids, or the inorganic phosphate group respectively to the substrate present a different problem in the design of synthetic substrates for reporting enzyme activities. While the addition of these chemical groups to the endogeneous substrates of these enzymes can have profound effects on protein conformation and consequently on protein function, these effects are often not translated to short peptides. At the molecular level, however, the addition of charged phosphates or highly hydrophobic lipids causes a dramatic change in the local environment surrounding the other amino acid residues on the peptide. This has been exploited in the design of chemosensors that translates this change into a fluorescent readout, effectively relating the level of fluorescence to enzymatic activity. One of the earliest examples of the design of such sensors was provided by Lawrence and co-workers in 2002, who constructed peptide-based probes for kinases, one of the most important enzyme classes extensively involved in cellular function, from cell signaling to apoptosis [25]. A peptide sequence known to serve as a substrate for protein kinase C (PKC) is appended with a fluorophore with fluorescent 13 properties tunable by metal chelation. Phosphorylation of the peptide by the kinase introduces a receptor site for a divalent metal comprising the dicarboxylate moiety on the fluorophore and the newly introduced phosphate. Coordination of a divalent metal (ATP-associated Mg2+) alters the electronic environment of the fluorophore and induces a marked increase in fluorescence. Using a similar strategy, Imperiali and coworkers incorporated a chelation-sensitive sulfamido-oxine (Sox) unnatural amino acid into a kinase probe [26]. This sensor sought to address some shortcomings of the probe dveloped by Lawrence by utilizing a less bulky sensor fluororphore which displayed a greater increase in fluroescence and spatially segregating the sensing moiety and the kinase recognition domain. These probes have recently shown to be competent in monitoring protein kinases in complex cellular media [27]. In conclusion, this section in this thesis serves to highlight the important advances that have been made in enzyme assays as a result of employing organic dyes in peptide- or small-molecule based substrates. Continued research is certainly necessary in pushing the frontiers of enzyme assays and bioimaging to include enzymes and applications that have not yet been accessible. 14 CHAPTER 2 DEVELOPING NEW FLUOROGENIC SUBSTRATES FOR DETECTING PROTEASE ACTIVITY 2.1 Fluorogenic Protease Substrates for Detecting Protease Activity on the Microarray and in Live Cells Proteases are enzymes that catalyze the breakdown of proteins through the hydrolysis of the peptide bond. Approximately 2% of the human genome codes for proteases, which translates to at least 500-600 proteases identified to date [28]. Proteases are classified according to the mechanism of hydrolysis. There are four major classes of human proteases: the cysteine, serine/threonine, aspartic proteases and the metalloproteases. The first two classes hydrolyses the substrate by using an active site residue (Cys, Ser/Thr respectively) for nucleophilic attack on the amide bond, while the aspartic and metalloproteases use an activated water molecule as a nucleophile. Protease function was initially thought to be limited to the degradation of proteins associated with the food digestion process or for the intracellular recycling of amino acids. However, studies have revealed the roles of proteases in more complex biological processes such as signaling cascades. Excessive or inappropriate proteolysis leads to unwanted activation of protease signaling pathways, which may lead to detrimental physiological and pathological conditions. Consequently, many proteases have emerged as potential drug targets in disease states where the modulation of protease activity can have a corrective effect on aberrant or insufficient protease activity [29]. Understanding the protease in its native environment and its role in protease cascades is of paramount importance in validating the protease as a 15 drug target. This involves the identification of the protease’s endogenous substrates and the downstream effects of cleavage of these substrates. To aid in the identification of endogenous protease substrates, researchers have developed several approaches to profile the substrate specificity of the proteases. The mapping of residue preferences at each binding pocket of the protease can enable the prediction of substrate sequences that are cleaved in vivo, which in turn help to identify the endogenous protein substrates. These typically involve the construction of peptide libraries, either synthesized chemically or displayed biologically, and assessing the residues (Pn – Pn’) that are most preferred at each position (Sn – Sn’) [30]. The standard nomenclature used to designate substrate/inhibitor residues that bind to corresponding enzyme subsites is shown in Figure 2.1a. The optimal peptide sequence derived from such studies may be converted to a fluorogenic peptide substrate to detect protease activity in inhibitor screening and in live cells. These fluorogenic substrates emit a fluorescence signal after it is cleaved by the protease of interest. Recording the fluorescence readout over a period of time gives the kinetics of the enzymatic reaction. This fluorescence signal is also often the mode of detection for imaging protease activity in whole cells. S3 a) S1 b) S2' P3 N-terminus P3 P2 P1' P1 S2 P3 H 2N O P2 P3' P1 N H H N O O P1' C-terminus Q N H S3' S1' O H N P2' P2' N H O H N O P2 P3 H2N O O H N H N P1 N H O O P1 N H O H N P1' H N O P2 O AMC: R = CH3 ACC: R = CH2 COOH OH P3' P2' N H H N O O O F P3' O R Protease substrate Figure 2.1. a) Protease and protease substrate nomenclature. b) 2 types of synthetic peptide substrates commonly used for assaying protease activity. i) AMC- or ACC-based substrates; ii) FRET-based substrates. 16 Fluorogenic peptide substrates, including those employing latent fluorophores, internally quenched and fluorescence resonance energy transfer (FRET)-based substrates, have emerged as indispensable tools in the profiling and visualization of protease activities both in vitro and in vivo [31]. Two types of synthetic fluorogenic peptides are widely used in high-throughput screening of protease inhibitors: i) extended FRET-based peptide substrates containing fluorophore and a dark quencher and ii) fluorogenic peptide substrates containing a C-terminally capped coumarin derivative (i.e. 7-amino-4-methylcoumarin (AMC) or 7-amino-4- carbamoylmethylcoumarin (ACC)). The fluorescence in FRET-based substrates is suppressed by the dark quencher which absorbs the light emitted by the fluorophore. Cleavage of the peptide substrate results in the spatial separation of the fluorophore and quencher. Energy transfer becomes extremely inefficient and negligible, leading to an increase in fluorescence from the fluorophore. AMC-/ACC-based substrates contain a coumarin moiety which is fluorescently silent when capped with a peptide sequence. They are arguably the most useful for substrate specificity profiling experiments, as only cleavage at the amide bond between the peptide and the coumarin moiety will release the highly fluorescent coumarin [32]. Consequently, coumarin-based fluorogenic peptide libraries have been employed to study the substrate specificities of numerous therapeutically important proteases, including caspases, thrombin, cathepsins and many others. In recent years, several attempts have been made to introduce these substrate libraries to microarray-based applications where further miniaturization and higher throughput of enzymatic assays can be achieved [15, 16]. We and others recently reported the immobilization of coumarinbased enzyme substrates onto microarray platforms and used them to profile substrate specificities of proteases. Since coumarin dyes are excited in the UV region 17 (maximum λex ~350 nm), these strategies have not been effective due to high fluorescence backgrounds and the lack of microarray scanners with UV light sources. For similar reasons, coumarin-based peptide substrates are rarely used in live-cell imaging experiments. The aim in this current work is thus to replace coumarin with a new fluorophore having excitation and emission wavelengths in the visible range, so that it can have dual utilities in both microarray and live-cell imaging applications. 2.2 Design of a New Fluorophore for Microarray and Bioimaging Applications In designing a suitable fluorophore, we turned to other fluorescein and rhodamine fluorophores that have been used for labeling reagents and enzymatic assays [34]. Of these, Rhodamine 110 (R110)-based substrates are well-established peptide probes for serine and cysteine proteases [35, 36]. Despite the desirable fluorescence properties of R110, several drawbacks hinder the direct use of these substrates: (1) R110-conjugated peptides require both peptide groups to be cleaved in order to generate maximum fluorescence, and thus are not suitable for quantitative studies of linear enzyme kinetics; (2) ‘asymmetric’ versions of these dyes containing a single peptide cleavage site lack an immobilization handle which is essential for both solid-phase peptide synthesis and microarray applications; (3) equilibrium between the quinone and the non-fluorescent spirolactone forms of R110 reduces fluorescence output. 18 H2N O O H2N Cl- O OH NH2 O O COOH Rhodamine 110 O Single-step enzymatic cleavage for peptide conjugates O 7-aminocoumarin4-acetic acid (ACC) HN OH 2-Me Tokyo Green Singapore Green OMe • Microarray immobilization • Anchor for solid phase synthesis • Attachment of subcellular localization singals / PTDs Figure 2.2. Structures of common fluorophores used in fluorogenic peptide substrates (ACC and Rhodamine 110) and fluorophores from which Singapore Green was derived (Rhodamine 110 and Tokyo Green). Our new fluorophore, Singapore Green (SG), is a hybrid of R110 and the fluorescein analog 2-Me TokyoGreen [37], with a phenolic group on one end providing a chemical handle (for solid-phase peptide synthesis, microarray immobilization and potentially other applications in cell-based experiments), and an amino group on the other end serving as the point of conjugation to a peptide sequence (Figure 2.2). We reasoned that, amidation at the amino group of SG by a peptide should suppress the fluorescence of the dye, similar to coumarin-based peptide substrates. Cleavage of the amide bond by a protease should release the highly fluorescent SG, thus reporting protease activity accordingly. Herein, we report the synthesis and characterizations of SG, the solid-phase synthesis of SG-conjugated peptides, as well as their applications in microarray-based and live-cell imaging experiments. 19 2.3 Chemical Synthesis of SG and SG-Conjugated Peptides 2.3.1 Chemical Synthesis of SG1 and SG2 O O O H 2N O DMF OH H2N 2i TrtHN O DMF, rt OTBS O TrtN Br O TrtHN 80% 2ii O MgBr CPh3 Cl, pyridine TBS-Cl, imidazole, O CO2tBu CO2tBu 5 5 K2CO3, DMF O 2iii HN TFA, H2O OTBS THF, 50οC 36% O O COOH 5 DCM SG-COOH 2iv 2v Scheme 2.1. Initial proposed synthesis of SG-COOH. In designing a synthesis strategy to SG and its related derivatives, we first conceived a route that took advantage of a published xanthone intermediate [38] which was subsequently protected with a trityl group on the aniline (Scheme 2.1). Grignard addition of o-tolylmagnesium bromide to the ketone gave the corresponding xanthene which could undergo alkylation to install a linker functionalized with a protected carboxylic acid at the end for anchoring onto the resin for solid-phase synthesis. However, during the course of the synthesis, it was found that the TBS group was cleaved off during the Grignard reaction before acidic work-up. This was an unusual occurrence since this protecting group is well-known for its stability under strongly basic anhydrous conditions. We reasoned that deprotection of the silyl group resulted from the nucleophilic attack of the Grignard reagent with the phenoxide anion acting as a stable leaving group. The extra stability of the phenol is conferred by the delocalization of the negative charge into the neighbouring carbonyl group. This enol form is stabilized by the extended conjugated system of the planar xanthone unit (Figure 2.3). Due to this delocalization, the C=O carbon is less electrophilic and the 20 reactivity towards nucleophiles is greatly reduced. This led to a sluggish Grignard reaction in which the starting material remained unconsumed even after 3 days of heating 50οC, resulting in a low yield. O O less electrophilic than usual C=O carbon O TBS deprotection under basic conditions TrtHN O TrtHN OTBS O O TrtHN O O Figure 2.3. Resonance stabilization of phenolate anion resulting from TBS deprotection The subsequent alkylation also proceeded slowly as the stable xanthene core existed predominantly as the ketone form and O-alkylation required a shift in electron density to the oxygen atom in the imine form. The imine form is thermodynamically less stable and is disfavored due to increased steric clash between the xanthene rings and the three bulky phenyl rings of the trityl group resulting from the rigid C=N bond (Figure 2.4). Both elevated temperatures (60οC) and microwave heating (70οC) for 30 min did not significantly improve the yield. Ph Ph H N O O Ph N O Ph Ph OH TrtHN O OR R Br base Ketone form (major) Imine form (minor) Figure 2.4. The 2 major resonance structures of the asymmetric xanthene. The ketone form suffers less from steric clash between the 3 phenyl groups and the xanthene core due to a rotatable C-N bond, while the imine form has a rigid C=N bond and is thermodynamically less stable. Subsequent alkylation with an alkyl bromide is inefficient due to slow equilibration to the imine form. In face of these 2 synthetic problems involving advanced intermediates, we conceived another strategy in which the linker unit was installed early in the route. 21 We reasoned that the ether linkage on the phenol should be stable during the Grignard reaction and the carbonyl carbon will thus be more reactive in the nucleophilic addition. This synthesis route was tried and optimized as shown in Scheme 2.2. We decided to synthesize both SG1, a simple methyl ether as a green-fluorescing analog of AMC for spectroscopic analysis and SG2 with the same fluorophore structure with an additional linker for attachment onto solid support or for conjugation with other functionalities. Cl 1. NaNO2, 50% H2SO4, 0οC 1. K2CO3, Cu, DMF, 130oC OH + O 2N O O O HO NHAc 2. H2SO4, 80oC O2 N O (20% in 2 steps) O (MeO)2SO2 or I 2. H2O, 95οC O 2N O (93%) CPh3Cl, NEt3 SnCl2.2H2O, EtOH, reflux O 2N O OR or H2, 10% Pd/C, EtOAc 2-3a: R = CH3 (70%) 2-3b: R = (CH2)5OTBS (82%) O H2N O OR HN 1. O DCM 2-4a: R = CH3 (90%) 2-4b: R = (CH2)5OTBS (92%) O OMe HN O MgBr TrtHN OH 2-2 O OTBS 5 K2CO3, DMF NH2 2-1 O OH 5 , THF, 50οC OR 2-5a: R = CH3 (88%) 2-5b: R = (CH2)5OTBS (97%) 2. DCM/TFA/H2O 7:2:1 SG1 (68%) SG2 (56%) Scheme 2.2. Synthesis of SG1 and SG2. The synthesis of SG started with the formation of the asymmetric xanthone 1, which was generated by Ullman-type coupling between 3-acetamidophenol and 2chloro-4-nitrobenzoic acid based on a published procedure. Diazotization of the amino group and hydrolysis of the diazonium salt yielded the phenol 2 which underwent methylation or alkylation with a linker unit to give 3. The nitro group was subsequently reduced to give 4a and 4b. Subsequent trityl-protection and Grignard addition of o-tolyl magnesium bromide followed by deprotection of the trityl group afforded the fluorophores SG1 and SG2, respectively. 22 2.3.2 Derivatization of SG1 and SG2 and Solid-Phase Synthesis of SGConjugated Peptides as Fluorogenic Protease Substrates To test if peptide conjugates of SG1 can be used as fluorogenic probes to detect enzymatic activity, we synthesized Ac-DEVD-SG1 as an analog of Ac-DEVDAMC, a coumarin-based substrate for the enzymes caspase-3 and caspase-7 from commercial sources. We adopted a solid-phase approach for the synthesis of the tetrapeptide. To this end, SG1 was first coupled in solution to Fmoc-Asp(OtBu)-OH using standard peptide coupling reagents, HBTU, HOBt and DIEA, followed by TFA deprotection of the t-butyl ester on the Fmoc-Asp moiety, leaving a free acid, FmocAsp-SG1 for loading onto solid support for the subsequent peptide synthesis (Scheme 2.3). 2 types of resins were suitable for our purposes: the Wang (p-benzyloxybenzyl alcohol) and the 2-chlorotritylchloride resins. Loading of an acid onto Wang resin typically involves pre-activation of the acid by carbodiimide reagents, e.g. diisopropylcarbodimide (DIC) to form a symmetrical anhydride followed by addition of the anhydride dissolved in DCM and a catalytic amount of DMAP to the alcoholfunctionalized resin to form an ester linkage. This procedure was first tried but loading onto Wang resin could not be carried out due to unsuccessful formation of the symmetrical anhydride of Fmoc-Asp-SG1. Instead, the N-acylurea formed between DIC and Fmoc-Asp-SG1 was found. This side reaction took place possibly because of the low nucleophilicity of the carboxylate anion which is hindered from attack by the bulky SG1 (Figure 2.5). 23 N N O O N FmocHN O O C N O N H O O N FmocHN N N H O O FmocHN O O O N O O O O-acylisourea N-acylurea Figure 2.5. Formation of the undesired N-acylurea from Fmoc-Asp-SG1 and DIC. The carboxylic acid is first deprotonated by DIC to form the carboxylate anion as the active nucleophile which adds to DIC. In the absence of a second nucleophile, an intramolecular N O acyl shift takes place to form the N-acylurea from the O-acylisourea. Following this observation, we tried loading Fmoc-Asp-SG1 onto the 2chlorotritylchloride resin with the use of a weak base, N,N-diisopropylethylamine (DIEA). The reaction was rapid and loading was successful with just 2 h of reaction time. While converting Fmoc-Asp-SG1 to an acid chloride and coupling onto Wang resin was another viable option, we chose the 2-chlorotritylchloride resin for subsequent peptide synthesis as it did not require further manipulation of Fmoc-AspSG1 (Scheme 2.3). Standard peptide synthesis on solid phase was carried out using the well-established Fmoc chemistry. Deprotection of the side chain protecting groups and cleavage from solid support with TFA and TIS as a radical scavenger afforded Ac-DEVD-SG1. To characterize the tetrapeptide product, LC-MS analysis was carried out. The desired product was obtained in good purity. (Figure 2.6) 24 O HN O O O Cl OH N FmocHN O N Cl O O FmocHN O a, b O O O c O O O FmocHN d O N H OtBu O H N O O e O O O N O AcHN O HO O N H O H N O O O f O OH O g O O N N H O O N H O O H N FmocHN O OtBu O AcHN O N N H OtBu OH O N H O N O O O Ac-DEVD-SG1 Scheme 2.3. Derivatization of SG1 and solid phase synthesis of caspase-3/7 probe, AcDEVD-SG1. Reagents and conditions: (a) Fmoc-Asp(OtBu)-OH, HBTU, HOBT, DIEA, 2 h; (b) 20% TFA/DCM, 5 h; (c) DIEA, DCM, 2h; (d) i: 20% piperidine/DMF, 30 min; ii: Fmoc-Val-OH, HBTU, HOBt, DIEA, DMF, 3 h; (e) i: 20% piperidine/DMF, 30 min; Fmoc-Glu(OtBu)-OH, HBTU, HOBt, DIEA, DMF, 3 h; (f) i: 20% piperidine/DMF, 30 min; ii: Ac-Asp(OtBu)-OH, HBTU, HOBt, DIEA, DMF; (e) TFA/TIS (95:5), 2.5 h. m AU(x1,000) 214nm ,4nm (1.00) O OH OH 3.0 O AcHN 2.0 HO 1.0 H N N H O O O N N H O O O Ac-DEVD-SG1 O Exact Mass: 815.3 0.0 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in Inten.(x1,000,000) 5.0 316.103 4.0 3.0 816.224 2.0 1.0 0.0 250 500 750 1000 Figure 2.6. LC-MS profile of Ac-DEVD-SG1. 25 1250 1500 1750 m /z We next sought to test the feasibility of using peptide conjugates of SG2 for reporting protease activities on the microarray. We picked 10 peptide sequences from commercially available p-nitroanilide or coumarin-based substrates targeting various proteases and replaced the p-nitroaniline chromophore or coumarin fluorophores with SG2. While the synthesis of Ac-DEVD-SG1 was straightforward, the same strategy could not be applied to all peptide sequences as it required a functional group on the first amino acid that could be attached to the resin, which precludes amino acids with alkyl side chains. We thus used another approach for the synthesis of these 10 different peptide sequences containing SG2. The alcohol functional group could be oxidized to an aldehyde which could serve as both an anchor to the resin and for attachment to functionalized glass slides for microarray-based experiments. To render the fluorophore compatible with Fmoc-based chemistry used in solid-phase peptide synthesis, the amino end of SG2 was protected with Fmoc, giving 2-8 and the hydroxyl-containing linker at the other end was oxidized to an aldehyde with DessMartin periodinane to give 2-9 (Scheme 2.4). HN O O 4 FmocN OH O O 4 Fmoc-Cl, NaHCO3, OH FmocN O O 4 O DMP, DCM THF/H2O, 0οC - rt SG2 2-8 2-9 Scheme 2.4. Synthesis of Fmoc-SG2-CHO (2-9) for peptide synthesis We adopted a solid-phase synthetic strategy that had been used for the preparation of peptide aldehyde libraries on threonine-functionalized resin [39]. As shown in Scheme 5, this functional resin was synthesized from aminomethyl polystyrene resin using standard Fmoc chemistry for peptide couplings. This was followed by deprotection of the Fmoc group with 20% piperidine in DMF and the 26 tert-butyl ether group with 95% TFA/TIS. The resulting ammonium salt was neutralized by stirring the resin in 10% DIEA/DCM to give a 1,2-amino alcohol. Fmoc-SG2-CHO was then loaded onto the resin via acid-catalyzed oxazolidine formation. Following Boc protection of the secondary amine to prevent acylation and hydrolytic cleavage of the oxazolidine ring during peptide synthesis, Fmoc deprotection of the resin-bound SG2 was carried out and standard solid phase peptide synthesis ensued. The N-terminus of the peptide was capped with an acetyl group using acetic anhydride, followed by deprotection of the amino acid side chains and the Boc group on the oxazolidine with dry TFA. For peptides containing Arg, 1% H2O was included as a scavenger for the released Pbf group and the deprotection was prolonged to 2 h to ensure complete deprotection. The SG2-peptide conjugates were then released from the solid support by acid-catalyzed hydrolysis of the oxazolidine ring, yielding the desired aldehyde-functionalized peptides in sufficient purity to be 27 O H N a H2N H2N b FmocHN O FmocN O H N O d 3 O O H N N H f N H OtBu O H N O FmocN e 3 O Boc N O N AcHN [AA]n O 3 N n[AA] O 3 AcHN N n[AA] O O N H N H O O O N H H N O O H N O O N H H N H N O O H N O O O Boc N O O h O O HN AcHN Boc N O O O g N H HO 3 Peptide coupling H2N c H N O O O Scheme 2.5. Solid phase synthesis of aldehyde-functionalized SG2-peptide conjugates. Reagents and conditions: (a) i: Fmoc-Gly-OH, HBTU, HOBt, DIEA, DMF, 2 h; ii: 20% piperidine/DMF, 30 min. (b) Fmoc-Thr(OtBu)-OH, HBTU, HOBt, DIEA, DMF, 2 h. (c) i: 20% piperidine/DMF, 30 min; ii: TFA/TIS (95:5) 1 h; iii: 10% DIEA/DCM. (d) Fmoc-SG2-CHO, MeOH/DCM/DMF/AcOH 6:2:1:1. (e) Boc2O, DIEA, DCM, 3 h. (f) 20% piperidine/DMF, 30 min. (g) TFA/TIS (95:5) or TFA/TIS/H2O (95:4:1), 45 min – 2 h. (h) DCM/MeOH/AcOH/H2O (12:5:2:1) used without purification for subsequent enzymatic screening. The crude products with the exception of P9 showed a single major peak in LC-MS profiles, thus demonstrating the compatibility of SG2 with standard Fmoc chemistry (Figure 2.7). 28 Ac-FG-SG2-CHO. IT-TOF-MS: m/z [M+1]+ calcd: 632.268, found 632.212. m AU(x100) 4.0 214nm ,4nm (1.00) 3.0 2.0 1.0 0.0 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in Inten.(x1,000,000) 632.212 4.0 3.0 2.0 1.0 0.0 250 500 750 1000 1250 1500 1750 m /z Ac-EY-SG2-CHO. IT-TOF-MS: m/z [M+1]+ calcd: 720.284, found 720.227. m AU(x1,000) 214nm ,4nm (1.00) 1.50 1.25 1.00 0.75 0.50 0.25 0.00 -0.25 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in Inten.(x1,000,000) 720.227 1.5 1.0 0.5 0.0 250 500 750 1000 1250 1500 1750 m /z Ac-FRR-SG2-CHO. IT-TOF-MS: m/z [M/2+1]+ calcd: 444.225, found 444.180. m AU(x1,000) 214nm ,4nm (1.00) 1.50 1.25 1.00 0.75 0.50 0.25 0.00 -0.25 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in Inten.(x1,000,000) 5.0 444.180 2.5 386.128 502.227 0.0 250 500 750 1000 29 1250 1500 1750 m /z Ac-AAF-SG2-CHO. IT-TOF-MS: m/z [M+1]+ calcd: 717.321, found 717.246. m AU(x1,000) 3.0 214nm ,4nm (1.00) 2.5 2.0 1.5 1.0 0.5 0.0 -0.5 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in Inten.(x1,000,000) 5.0 4.0 717.246 3.0 2.0 1.0 386.132 0.0 250 500 750 1000 1250 1500 1750 m /z Ac-AAL-SG2-CHO. IT-TOF-MS: m/z [M+1]+ calcd: 683.337, found 683.306. m AU(x1,000) 1.50 214nm ,4nm (1.00) 1.25 1.00 0.75 0.50 0.25 0.00 -0.25 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in Inten.(x100,000) 683.306 7.5 5.0 2.5 302.099 0.0 250 500 750 1000 1250 1500 1750 m /z Ac-VPR-SG2-CHO. IT-TOF-MS: m/z [M+1]+ calcd: 780.401, found 780.315. m AU(x1,000) 1.50 214nm ,4nm (1.00) 1.25 1.00 0.75 0.50 0.25 0.00 -0.25 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in Inten.(x1,000,000) 3.0 390.662 2.0 386.131 780.315 1.0 0.0 250 500 750 1000 30 1250 1500 1750 m /z Ac-DEVD-SG2-CHO. IT-TOF-MS: m/z [M+1]+ calcd: 886.343, found 886.242. m AU(x100) 7.5 214nm ,4nm (1.00) 5.0 2.5 0.0 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in Inten.(x1,000,000) 386.130 4.0 3.0 2.0 1.0 886.242 0.0 250 500 750 1000 1250 1500 1750 m /z Ac-YVAD-SG2-CHO. IT-TOF-MS: m/z [M+1]+ calcd: 876.374, found 876.265. m AU(x100) 214nm ,4nm (1.00) 7.5 5.0 2.5 0.0 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in Inten.(x1,000,000) 3.0 386.128 2.0 1.0 876.265 0.0 250 500 750 1000 1250 1500 1750 m /z Ac-ENLYFQ-SG2-CHO. IT-TOF-MS: m/z [M/2+1]+ calcd: 611.769, found 611.711. m AU(x100) 7.5 214nm ,4nm (1.00) 5.0 2.5 0.0 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in Inten.(x1,000,000) 428.142 3.0 2.0 1.0 302.086 611.711 0.0 250 500 750 1000 31 1250 1500 1750 m /z Ac-RPFLLHVY-SG2-CHO. IT-TOF-MS: m/z [M/2+1]+ calcd: 727.380, found 727.271. m AU(x1,000) 214nm ,4nm (1.00) 1.5 1.0 0.5 0.0 0.0 2.5 5.0 Inten.(x10,000) 5.0 7.5 10.0 12.5 15.0 17.5 m in 727.271 4.0 3.0 527.222 2.0 1.0 0.0 250 500 750 1000 1250 1500 1750 m /z Figure 2.7. LC-MS profiles of the 10 SG2-peptide conjugates. LC conditions: 10-100% CH3CN in 20 min. 2.4 Profiling Protease Activity in Microplate and on the Microarray We first evaluated the spectroscopic properties of SG1 to determine its suitability for integration into a fluorescence reporter system. SG1 was found to have excitation maxima at 469 and 473 nm and emission maxima at 517 nm in ethanol (Fig 2.8), similar to fluorescein (494 and 521 nm respectively, in water), and is thus compatible with the 488 nm argon-ion laser used in most microarray scanners and fluorescence microscopes. SG1 has a quantum yield of 0.50 and an extinction coefficient of 28500 M-1cm-1 which is reasonably bright for most applications. With the 11 SG-peptide conjugates in hand, our aim was to evaluate them as synthetic substrates for assaying protease activity on both the conventional microplate platform and the microarray. 32 b) Fluorescence Intensity a) 800 818.9 750 700 650 600 600 550 500 450 400 400 350 300 250 200 200 150 100 50 0.7 390.0 400 420 440 400 460 480 480 500 52 0 NM 540 560 580 560 600 620 640.0 640 Wavelength (nm) Relative Fluorescence (x10 3) c) 30 25 20 15 10 5 0 0 20 40 Tim e (m in) 60 Figure 2.8. a) Protease cleavage of SG-peptide conjugates. Enzymatic hydrolysis of the anilide bond results in a fluorescence increase due to the release of SG. b) Excitation (blue) and emission (red) spectra of SG1. c) Fluorescence increase from cleavage of Ac-DEVDSG1 by caspase-3 (blue) and caspase-7 (red) over the background fluorescence (orange) We next tested Ac-DEVD-SG1 which contains the optimal substrate sequence for the cysteine proteases caspase-3 and -7. Ac-DEVD-SG1 was weakly fluorescent. Upon incubation with caspase-3 and -7, there was a time-dependent increase in fluorescence resulting from the cleavage of the amide bond between the Asp residue and SG1. The fluorescence increase follows typical Michaelis-Menten kinetics, indicating that SG-based peptide conjugates are indeed suitable green light-emitting substitutes of the well-established coumarin-based substrates. Having shown the feasibility of using our tetrapeptide SG-conjugated substrate to report proteolysis, we moved to the microarray platform, where the substrates are immobilized on the glass surface and the protease is applied onto the 33 surface. Proteolysis results in a fluorescence signal on the glass slide, which is O NH2 O SG NH2 PEPTIDE SG SG N O O PEPTIDE PEPTIDE PEPTIDE recorded and quantified by a microarray scanner. SG N SG N O O N O H Protease Figure 2.9. Detecting protease activity on the microarray. To generate our peptide microarray, glass slides had to be appropriately functionalized with hydroxylamines for chemoselective ligation with the SG2-peptide aldehydes. This was accomplished by washing the glass slides in piranha solution (conc. H2SO4/ 30% H2O2 7:3) which hydroxylates the glass surface, forming silanols. The exposed silanols are then reacted with aminopropyltriethoxysilane in the presence of a small amount of water to yield disiloxanes with an amine terminal. The aminefunctionalized slides were then coupled with a phthalimide-protected hydroxylamine linker using standard coupling reagents for amide bond formation. The phthalimide group was then deprotected with hydrazine, giving the hydroxylamine slides (Scheme 2.6.). 34 NH2 NH2 OH HO OH HO OH HO Si Si NH2 O Si Si EtO EtO Si EtO O O O Si O O O Si EtOH, H2O O O HO N 5 O HBTU, DIEA, DMF amine-functionalized slides O O N O 5 O N O O O NH O O 5 NH 5 O 3% hydrazine/DMF NH2 O NH2 5 O NH NH hydroxylaminefunctionalized slides Scheme 2.6. Functionalization of glass slides with alkoxyamines. The actual Si species on the glass surface is probably a mixture of cross-linked disiloxanes and free silanols. We selected peptide sequences that were known substrates for proteases of different classes, and also of different specificity to test the robustness of our platform in screening for protease activity (Table 2.1). Peptide no. Peptide sequence Target Protease Protease Class P1 Ac-FG-SG2-CHO Papain Cysteine P2 Ac-EY-SG2-CHO Pepsin Aspartic P3 Ac-FRR-SG2-CHO Trypsin Serine P4 Ac-AAF-SG2-CHO Chymotrypsin Serine P5 Ac-AAL-SG2-CHO Subtilisin Serine P6 Ac-VPR-SG2-CHO Thrombin Serine P7 Ac-DEVD-SG2-CHO Caspase-3/-7 Cysteine P8 Ac-YVAD-SG2-CHO Caspase-1 Cysteine P9 Ac-ENLYFQ-SG2-CHO TEV Cysteine P10 Ac-RPFHLLVY-SG2-CHO Rennin Aspartic Table 2.1. Peptide sequences synthesized and their target proteases. 35 Cysteine proteases such as the caspases and TEV have stringent substrate specificity, requiring a specific peptide sequence for binding and catalysis to occur, while other proteases such as subtilisin has broader specificity. The fourth class of proteases, the metalloproteases, were not included as they generally require substrate recognition on the prime sites and thus their activity could not be monitored by our designed substrates. We spotted the aldehyde-functionalized peptides onto the hydroxylamine slides to generate the corresponding peptide microarray via oxime bond formation. As a proof-of-concept experiment, the immobilized peptides were treated with four different proteases (caspase-3, caspase-7, α-chymotrypsin and subtilisin). The microarray was subsequently scanned with a microarray scanner equipped with a blue light source. Images obtained immediately revealed discerning “substrate fingerprints” of each enzyme. Of the proteases used, enzymes with broader substrate specificity (subtilisin and α-chymotrypsin) cleaved multiple substrates to different extents, while highly specific proteases (caspase-3 and -7) cleaved only its optimal substrate sequence, Ac-DEVD-SG2. thrombin P2 P4 P6 P8 P10 a) subtilisin α-chymotrypsin trypsin b) P2 Time (min) P1 0 5 ) 15 30 60 P3 P4 P5 P6 Figure 2.10. a) Enzyme “fingerprints” obtained (clockwise from top left: a-chymotrypsin, subtilisin, caspase-3, caspase-7). Peptides were spotted in triplicate vertically. b) Time- 36 dependent kinetic profiles obtained from the peptide microarray. Peptides were spotted in duplicate vertically. We next examined whether the SG-based peptide microarray could be used to obtain quantitative enzyme kinetic data by incubating four selected peptides with four different enzymes in a time-dependent experiment. Fluorescence intensities were quantified and fitted to kinetic curves to obtain kobs values which reflected the substrate preferences of the proteases. Taken together, these experiments indicate the applicability of SG-based substrates in microarray-based protease profiling experiments. a) a-chymotrypsin/P4 a-chymotrypsin/P5 a-chymotrypsin/P6 750 700 500 600 400 500 500 300 400 R2 = 0.98 kobs = 0.18 300 250 200 100 100 0 0 0 0 10 20 30 40 50 60 0 70 10 20 30 40 50 60 Subtilisin/P4 30 R2 = 0.99 kobs = 0.21 250 0 40 50 60 70 0 10 20 30 Time 40 40 50 50 60 0 70 10 20 30 40 50 Time b) Subtilisin/P5 Subtilisin/P4 Trypsin/P6 3000 7000 3000 6000 2500 5000 2000 2000 1000 1000 4000 R2 = 0.99 kobs = 0.11 1500 R2 = 0.98 kobs = 0.16 3000 2000 500 1000 0 0 0 10 20 30 40 Time 50 60 70 0 10 20 30 40 Time 37 70 60 70 R2 = 0.99 kobs = 0.028 Time 3500 60 trypsin/P6 550 500 450 400 350 300 250 200 150 100 50 0 500 30 20 Time 750 20 10 Subtilisin/P5 900 800 700 600 500 400 300 200 100 0 10 0 70 Time Time 0 R2 = 0.98 kobs = 0.37 200 50 60 70 0 0 10 20 30 Time 40 a-chymotrypsin/P5 a-chymotrypsin/P4 a-chymotrypsin/P6 3000 4000 5000 4000 3000 2000 3000 R2 = 0.99 kobs = 0.097 2000 1000 R2 = 0.99 kobs = 0.14 2000 1000 1000 0 0 0 10 20 30 40 50 60 Time 70 0 0 10 20 30 40 50 60 70 Time 0 10 20 30 40 50 Time Figure 2.11. a) Selected kinetic data from microarray enzymatic assays shown in Figure 2.10. b) Microplate assays for the corresponding enzyme/peptide pair carried out as a comparison. 2.5 Imaging Caspase-3 and-7 Activities in Live Cells To demonstrate that our SG-based substrates can be used for live-cell imaging, we tested the ability of Ac-DEVD-SG1 to image apoptosis in live cells. Caspase-3 and -7 are key mediators of this important biological process where improper regulation of caspase activity has detrimental pathological and physiological effects. To image apoptosis, numerous peptide- and protein-based probes including Rhodamine 110 peptide conjugates have been developed from the sensitive detection of caspase activity [40,41]. We thus evaluated Ac-DEVD-SG1 as a fluorogenic probe for reporting caspase-3 and -7 activity in apoptotic HeLa cells. Cells treated with AcDEVD-SG1 developed a strong green fluorescence upon apoptosis induction with staurosporine. In contrast, no significant increase in green fluorescence was observed in non-apoptotic cells even after extended incubation time. Non-apoptotic cells remained viable after treatment with the probe for more than 3 h, indicated that the probe was not cytotoxic. 38 a) RFP GFP DIC b) c) d) Figure 2.12. Detecting caspase activity in live HeLa cells with Ac-DEVD-SG1. Cells were injected with 50 µM of Ac-DEVD-SG1 with tetramethylrhodamine-dextran as marker to identify injected cells in RFP channel. a) Injected cells before apoptosis induction. b) Cells showed a fluorescence increase in the GFP channel after treatment with staurosporine. Scale bar = 15 µm. c) Injected cells 3 h after apoptosis induction. d) Injected cells without treatment with staurosporine 2.6 Conclusions In conclusion, we have designed and synthesized a new green light-emitting fluorophore, Singapore Green (SG), which possesses desirable chemical and 39 fluorescence properties suitable for biomedical applications. We have shown that peptide conjugates of this new fluorophore can be readily synthesized using standard solid-phase peptide chemistry, and conveniently immobilized on a microarray for high-throughput substrate specificitiy profiling of proteases. We further showed that these probes are equally amenable for live-cell imaging of protease activities. To date, positional scanning libraries of coumarin-based peptide substrates are most commonly employed in profiling studies of proteases. These probes are however largely unsuitable for bioimaging purposes as the coumarin fluorophore has an excitation wavelength in the UV region and emission in the blue region where there is significant background fluorescence. As a result, optimal substrates derived from profiling studies cannot be directly be used as imaging agents for protease activity. By introducing SG, a fluorescein analog of ACC or AMC, this dual purpose may now be achieved. We anticipate that this chemically amenable fluorophore and its quenched peptide substrates will become useful tools for further developments in enzyme substrate specificity profiling and live-cell imaging. 40 CHAPTER 3 FLUOROGENIC PROBES FOR DETECTING PROTEASE ACTIVITY AT SUBCELLULAR LOCATIONS 3.1 Targeted Delivery of Molecules into the Cell The central theme in drug delivery research is the delivery of biologically active molecules to their primary site of action to increase therapeutic efficacy. This multi-faceted problem of involves several major issues: i) cell-specific targeting to ensure that therapeutics act on malignant cell types only; ii) transporting molecules into the intracellular space; and iii) delivering them to specific organelles where their intended targets are located. In recent years, researchers have been interested in adapting the methods developed for intracellular delivery to other biologically interesting molecules, as well as understanding the precise mechanisms of cellular uptake and internalization. Notably, a growing trend that has emerged is the shift of focus from enhancing cell permeability to attaining organelle-specific targeting. This stems from the general observation that cellular entry does not equate to access to the intended site of action within the cell for the molecule of interest, thereby largely diminishing the efficacy of various molecular transporters as true delivery vehicles. Academic researchers have henceforth embarked on the task of understanding the transport mechanisms associated with each organelle from a molecular perspective through the use of probes that interrogate organelle function or individual protein function within the organelle of interest. One of the major breakthroughs in cellular delivery came about through the discovery of peptide sequences which can efficiently translocate across the cell 41 membrane, known as cell-penetrating peptides (CPPs). The two most well-known CPPs are the Tat peptide [42] and Penetratin [43], derived respectively from the human immunodeficiency virus (HIV) transcriptional regulator Tat and the Drosophila transcription factor Antennapedia. These short peptide sequences of less than 20 amino acids were found to act as efficient carriers of diverse cargoes such as proteins, peptides and oligonucleotides conjugated to the peptides [44]. Synthetic analogues of these peptides, such as the oligoarginines, comprise the other family of CPPs. Since CPPs are easily synthesized by chemical methods, and may be genetically fused with protein cargoes, they are now routine tools for the general intracellular delivery of biologically active molecules. One of the major problems associated with the use of CPPs, however, has been the vesicular entrapment of the cargoes being trafficked. Endocytosis competes significantly with direct cell penetration in which the cargo is directly delivered into the cytoplasm. Cargo which has been internalized by endocytosis and fails to escape from the endosomes is eventually destroyed in the lysosomes, the cell’s demolition center, which means that the actual amount of cargo that is able to act on its target is less than that administered. Because the precise molecular mechanisms of the competing processes of endocytosis and penetration have not been elucidated, it remains difficult to manipulate them such that the latter process predominates. The true utility of CPPs as delivery vehicles is thus diminished. The abovementioned limitation of using CPPs highlights the question underpinning drug delivery research – how efficiently is the molecule of interest delivered to its intended site of action? One of the recent directions undertaken to address this issue is the design of methods to deliver the molecules of interest directly 42 to the organelle where they can perform their function. This concept of subcellular targeting is perhaps most well-demonstrated in therapeutic strategies used to modulate mitochondrial function. One of these strategies is the use of proapoptotic peptides, which once delivered into the tumor cell, permeabilize the mitochondrial membrane, resulting in the release of cytochrome c and leading to apoptotic death of the tumor cell. The selectivity of these peptides towards the mitochondrial membrane over the plasma membrane is critical for therapeutic use of these peptides. Peptides that cause cell lysis by permeabilization of the plasma membrane will not be effective since they cannot be made to recognize tumor cells by the attachment of tumor cell-specific moieties. Several reports describe the successful applications of such mitochondriatargeting, tumor-specific peptides [45]. In addition to its role in apoptosis, the mitochondrion is also the site where reactive oxygen species reside. Oxidative stress caused by reactive oxygen species has been linked to various diseases [46, 47], and this is found to be reduced through the use of radical scavengers such as TEMPO [48]. To address the problem of poor cell-permeability of these nitroxide scavengers, Wipf and co-workers designed and synthesized TEMPO derivatives conjugated to a membrane-active peptide sequence from the natural antibiotic Gramicidin S [49]. The resulting constructs were cell-permeable, mitochondria-targeting radical scavengers which prevented the increase in intracellular superoxide production induced. An important lesson can be learnt from these studies and numerous other strategies [50] developed to home in on the mitochondria. Researchers have recognized the importance of organelle targeting to achieve the desired therapeutic effect, but this has surprisingly not been utilized in small molecule drug discovery. A recent study by the Simons group [51] may mark a paradigm shift in inhibitor 43 development. They studied the inhibition of β-secretase, an endosomal protease whose aberrant activity is known to propagate Alzheimer’s disease with a known inhibitor and the same inhibitor attached to a sterol moiety which was a membrane anchor. The group showed that the sterol-linked inhibitor was effectively internalized by endocytosis after membrane anchoring, allowing it to inhibit the β-secretase within the endosome. This targeted inhibitor was shown to be markedly more potent than the free inhibitor, giving a definitive example of how understanding protein compartmentalization can lead to the design of inhibitors with better in vivo profiles by the single attachment of a targeting moiety. This principle could pave the way for the design of next-generation inhibitors and therapeutics which can home in on the specific organelle where their target resides, while escaping non-productive internalization pathways. It also calls for the re-evaluation of known inhibitors in the cellular context, from the perspective of addressing the localization of these inhibitors within the cell. Given the prospects of organelle-specific inhibitors or molecules that mediate protein function, enabling strategies which allow the identification and detection of resident proteins in these organelles should become important. In recent years, there have been a number of noteworthy reports on the global analysis of protein subcellular localization by imaging FP-tagged proteins [52]. These systems biology approaches serve to assign individual proteins to their respective organelles or to characterize the proteins within an organelle, but do not report the functional state of these proteins. Activity-based protein labeling and detecting enzyme activity in subcellular locations will thus provide another dimension to studying both the protein of interest in its subcellular microenvironment and organelle function as a whole. 44 There are some recent developments in this aspect. Overkleeft and co-workers developed an activity-based probe targeting cathepsins in antigen-presenting cells by modifying a known cathepsin probe with a mannose cluster to facilitate uptake through receptor-mediated endocytosis [53]. This process brings the probe to its target proteases in the lysosomes, enabling the fluorescent tagging of the proteases. Activity-based probes are well-poised for studies in organellar studies as they have proven to be powerful tools for protein profiling both in vitro and in vivo for different applications [54]. Overkleeft’s targeted probes present a plausible general strategy for developing future probes useful for interrogating the subcellular proteome. The most direct assessment of enzyme activity is the use of enzyme substrates which gives an easily measurable readout after the enzymatic reaction. They have long been used to assay enzyme activity for inhibitor development, but there have been recent developments in adapting known fluorogenic substrates for imaging. In particular, synthetic peptide substrates have been used routinely for imaging proteases and are thus used in efforts to develop improved substrates. Many of these efforts aim to develop improved substrates by increasing the cell permeability of the substrate for efficient cellular uptake and/or modifying substrates for practical use in in vivo systems [55]. There are however, few examples that report substrates which can image enzymatic activity that is confined within an organelle [56]. There is thus a need to develop fluorogenic substrates which can image the target enzymes in various subcellular locations. Imaging agents, activity-based probes and inhibitors targeted to specific organelles will constitute a multi-pronged approach to advance studies of the proteome at the subcellular level. 45 3.2 Design of Cell-Permeable Protease Substrates Targeting Different Organelles We recently developed a series of fluorogenic peptide substrates that could be used for detecting protease activity on the microarray and in live cells. We designed our fluorophore, SG2, a green light-emitting substitute for the blue-fluorescing ACC fluroophore, to be a versatile fluorophore which can be attached to other functionalities for various applications [22]. The long alkyl linker functionally separates SG2 from other appendages at the other end of the linker and enables synthetic manipulation without the need to re-synthesize the xanthene core structure. In the previous chapter, we synthesized SG2-peptide conjugates by using the linker with an aldehyde moiety as a point of attachment to the solid support and to the microarray glass slides. We also used the SG1-based peptide substrate, Ac-DEVDSG1, in imaging caspase -3/7 activity in apoptotic live cells. We also established that these dye-peptide conjugates can easily be synthesized using well-established Fmoc chemistry for solid-phase peptide synthesis. In addition, having demonstrated the compatibility of our SG-peptide conjugates in live-cell imaging, we looked to delivering substrates targeting various enzymes into different organelles within the cell to image protease activity in intracellular locations. One of the modes of delivering a cargo into a cell is the use of peptide sequences that are known to localize a protein to a particular organelle. Because enzymes can be localized at multiple organelles, or translocated from one organelle to another, we needed a synthetic strategy that would allow us to easily attach different localization sequences to different substrates. 46 We thus conceived a strategy in which the fluorogenic peptide substrates and the localization sequences are synthesized as separate modules and assembled using a ligation reaction. This reaction should be efficient and orthogonal to all the functional groups on the peptides since no protecting groups will be present after peptide synthesis, and should preferably be carried out under mild, water-compatible conditions. The best candidate reaction with the desired characteristics is the Cu(I)catalyzed azide-alkyne cycloaddition, the best known reaction in “click” chemistry [57]. To this end, we attached an alkyne handle to each of the peptide substrates and an azide moiety to each of the localization sequences. The desired fluorogenic peptide substrates targeted to different organelles can then be rapidly synthesized by “click” chemistry. Upon substrate recognition and protease cleavage in the respective organelles, the SG2-conjugated localization peptide will be released, leading to a fluorescence increase which can be detected by fluorescence imaging of the live cells. N N N N3 + N3 "Click" N N N N N N N N N Chemistry Protease N3 AlkyneSG2substrates Azidelocalization peptides N N N Targeted fluorogenic peptide substrates N N N Fluorescent localization peptides Imaging in subcellular organelles Figure 3.1. Overall strategy for imaging protease activity in subcellular organelles. Fluorogenic peptide substrates targeted to different organelles are assembled of individual protease substrates and localization peptides by “click” chemistry. Protease cleavage results in fluorescent localization peptides which can be detected by fluorescence imaging. 47 The primary objective in this work is to demonstrate the feasibility of using short peptide localization sequences to deliver fluorogenic peptide substrates to particular subcellular organelles for the detection of localized protease activity through fluorescence imaging. As a model system, we decided to use the induction of apoptosis to bring about the activation of caspases, which are known to be specific proteases [13]. Central to apoptotic events are the translocation and activation of procaspases upstream apoptotic mediators [58]. This results in further translocation of the caspases and/or the cleavage of substrates localized exclusively in a particular organelle, leading eventually to cell demolition. While the subcellular localization of caspases remains controversial arising from the use of different cell lines and apoptosis inducers, several key observations are consistent in the literature surveyed. The nuclear [59] and mitochondrial [60] localization of pro-caspases and caspases is well-established by subcellular fractionation and western blotting. It is thus imperative that we evaluate the utility of our approach in detecting caspase activity in the mitochondria and nucleus. Expanding on our previous imaging experiments with Ac-DEVD-SG1, we selected a few protease substrates that target proteases which were involved in apoptosis and/or are activated by external stimuli. We selected 3 caspase substrates, including the substrate for caspase-3 and -7 which has often been used for imaging apoptosis, as well as substrates for caspase 2 and caspase 9. We are also interested in other proteases such as the cysteine protease cathepsins which are localized exclusively to the endosomes, and in particular cathepsin B which is known to have a role in apoptosis [61]. The last peptide substrate that we picked is that for µ-calpain and m-calpain, which are activated in the presence of micromolar and millimolar concentrations of calcium respectively, within the cell 48 [62]. In addition, these calcium-dependent proteases are also involved in apoptosis [63]. By selecting caspases, cathepsins and calpains as our target enzymes, we could potentially study these cysteine proteases in synergy with our platform. The list of peptide substrates, their target proteases and the known subcellular localizations of the proteases are summarized in Table 3.1 below. Peptide Target Enzyme(s) SG2-(a) Ac-DEVD-SG2-alkyne Caspase-3/-7 substrate SG2-(b) Ac-VDVAD-SG2-alkyne Caspase-2 substrate SG2-(c) Ac-LEHD-SG2-alkyne Caspase-9 substrate SG2-(d) Ac-FR-SG2-alkyne General cathepsin substrate SG2-(e) Ac-FRR-SG2-alkyne Cathepsin B substrate SG2-(f) Ac-LLVY-SG2-alkyne Calpain I/II substrate Table 3.1 Alkyne-functionalized SG2-based substrates and their target enzymes To deliver these fluorogenic substrates to the desired subcellular compartment, we surveyed the literature for short peptide sequences that have been shown to localize specifically or function as efficient carriers of various cargo types to a particular organelle. A well-established peptide carrier is the Simian virus 40 (SV40) T-antigen nuclear localization signal (NLS) which has been used to deliver proteins and peptide substrates to the nucleus [64]. Another important organelle is the mitochondria which serves to produce the cell’s energy and as a checkpoint regulating apoptosis. In recent years, a plethora of delivery modes and targeting modules specific for the mitochondria has emerged. A systematic study elucidating the molecular requirements in designing synthetic peptides that exhibit efficient cellular uptake and can penetrate the mitochondria was recently carried out by the Kelley group [65]. In 49 the same study, the group identified a cationic, lipophilic peptide containing cyclohexane and arginine residues which were cell-permeable and mitochondriaspecific, which would serve our purpose as a mitochondria-targeting localization sequence. We further used an N-palmitoylated lysine residue to mimic a common post-translational modification that several membrane proteins undergo for trafficking to the plasma membrane [66]. The localization sequences used and their target organelles are summarized in Table 3.2 below. In addition, we included two cellpenetrating peptides (CPPs), also known as protein transduction domains (PTDs) as general modes of delivery into the intracellular space. These short peptide sequences are known to promote the cellular intake of small molecules, quantum dots and even proteins which would otherwise be cell-impermeable. The sequences of the localization peptides selected are summarized in Table 3.2 below. ID Peptide Target Organelle N3-SV40 N3-KKKRKV-NH2 nucleus N3-RrRK N3-RrRK-NH2 nucleus N3- FxrFxK N3-FxrFxK-NH2 mitochondria N3-KK N3-KK(palmitoyl)-NH2 membrane N3-Tat N3-RKKRRQRRR-NH2 general CPP, nucleus N3-R9 N3-RRRRRRRRR-NH2 general CPP Table 3.2 Azide-functionalized localization peptides selected and their target organelles To evaluate whether these localization sequences indeed localize to the correct organelle, we labeled the peptides with SG2 to form SG2-peptide conjugates whose localization can be visualized under the fluorescence microscope. 50 3.3 Chemical Synthesis of Peptide Substrates and Localization Peptides 3.3.1 Chemical Synthesis of Alkyne-Tagged Peptide Substrates The key considerations in designing the synthetic routes to our peptide substrates are i) the placement of the various functional groups (fluorogenic peptide substrate and alkyne handle) and ii) adapting the synthesis on solid phase for facile construction of different peptides. O FmocN O O 4 2-8 FmocN OH O O 4 O FmocN OH O O 4 PDC, DMF (COCl)2, cat. DMF 0oC - rt CH2Cl2 3-1 Cl 3-2 Scheme 3.1 Synthesis of Fmoc-SG2-COOH (3-1) and Fmoc-SG2-COCl (3-2) In our synthetic strategy, we decided to transform SG2 into an Fmoc-protected unnatural amino acid to enable conventional solid-phase peptide synthesis using Fmoc chemistry, which has already been demonstrated to be suitable chemistry for the fluorophore. SG2 was first protected with Fmoc as described in Chapter 2.3.2 and the alcohol moiety was oxidized to the acid Fmoc-SG2-COOH (3-1) using standard oxidation procedures with PDC (Scheme 3.1). The acid group could serve as a point of attachment to the solid support and also to an alkyne moiety for “click” chemistry with azide-conjugated localization sequences. To this end, we started solid-phase synthesis with the reductive amination of the aldehyde-functionalized PL-FMP (polystyrene – 4-formyl-3-methoxyphenoxy resin) with propargyl amine to give a secondary amine. This was followed by the acylation with Fmoc-SG2-COOH which would result in the simultaneous loading of the Fmoc-protected dye onto the solid 51 support and tagging with an alkyne handle. Several conditions were tried to effect the coupling reaction between Fmoc-SG2-COOH and the resin-bound secondary amine. Both overnight reaction using HBTU/HOBt and the stronger coupling reagent PyBrOP failed to load the compound onto the solid support. This was evidenced from the product that was cleaved from the solid support after Fmoc deprotection and coupling of the Fmoc-Asp(OtBu)-OH using HBTU/HOBt. Instead of obtaining the desired Fmoc-Asp-SG2-alkyne, Fmoc-Asp-alkyne was obtained in good purity, indicating that the acylation reaction between Fmoc-SG2-COOH did not take place. Since the same reaction worked well for Fmoc-Asp(OtBu)-OH, it is unlikely that the coupling conditions were not strong enough when Fmoc-SG2-COOH was used. We reasoned that a possible reason why the desired acylation reaction did not occur was the degradation of Fmoc-SG2-COOH via the cyclization of the linker to form a γlactone under the basic conditions employed for the coupling reaction (Figure 3.2). a) O NaBH3CN Fmoc-SG2-COOH DMF/MeOH/AcOH conditions HN OMe FmocN O O N 3 O OMe OMe O b) FmocN O O O O O FmocN O O- FmocHN O O - Figure 3.2. a) Attempted attachment of Fmoc-SG2-COOH by acylation of Fmoc-SG2-COOH and resin-bound secondary amine using various coupling reagents. b) Possible cyclization of linker moiety on Fmoc-SG2-COOH under basic conditions. To prevent this side reaction, Fmoc-SG2-COOH was converted into the corresponding acid chloride using oxalyl chloride with catalytic DMF (Scheme 3.1). The acid chloride was then used directly for coupling onto the secondary amine, 52 which resulted in the successful loading of Fmoc-SG2-COOH onto the solid support. Following Fmoc deprotection of resin-bound SG2, standard solid phase peptide synthesis was carried out employing Fmoc chemistry to furnish the desired peptide substrate sequence (Scheme 3.2). FmocN a HN O HN O O O b N c 3 OMe O OMe O O OMe O N 3 O Peptide coupling H2N [AA]n d N AcHN [AA]n N O N 3 O OMe O O OMe N 3 O e AcHN [AA]n O N O H N O 3 O OMe O Scheme 3.2. Solid-phase synthesis of alkyne-functionalized substrates, Ac-X-SG2-alkyne. a) NaBH3CN, DMF/MeOH/AcOH (8:1.9:0.1), 16 h; b) Fmoc-SG2-COCl (3-2), DIEA, CH2Cl2, 12 h; 20% piperidine/DMF, 30 min; d) Ac2O, DIEA, CH2Cl2, 3 h; e) TFA/H2O/TIS (95:2.5:2.5), 2 h. The N-terminus of the peptide was capped with an acetyl group by reaction with acetic anhydride. Cleavage from the solid support with concomitant deprotection of the side chain protecting groups using TFA/H2O/TIS was carried out to obtain the desired alkyne-tagged peptide substrates which were characterized by LC-MS analysis. In all 6 peptide conjugates, 2 major peaks were found, one corresponding to the desired product (abbreviated Ac-X-SG2-alkyne, X is the peptide sequence) and the other corresponding to the peptide sequence directly conjugated to the alkyne without SG2 (Figure 3.3a). The occurrence of this product indicates that the acylation 53 of the acid chloride of Fmoc-SG2-COOH and the secondary amine was not complete, possibly due to the steric bulk of the dye and the high loading level of the resin (0.9 mmol/g). a) O H N O R N N H n R O H N O 3 O O H N O O Ac-X-SG2-alkyne, X = peptide R R N H n H N O Ac-X-alkyne, X = peptide b) 1. Ac-DEVD-SG2-alkyne m AU(x1,000) 214nm ,4nm (1.00) 3.0 2.0 * 1.0 0.0 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in 1750 m /z 17.5 m in 1750 m /z Inten.(x100,000) 939.3448 5.0 439.1883 596.2136 2.5 157.0301 302.1115 559.1855 0.0 250 500 750 1000 1250 1500 2. Ac-VDVAD-SG2-alkyne m AU(x1,000) 4.0 214nm ,4nm (1.00) 3.0 2.0 * 1.0 0.0 0.0 2.5 5.0 7.5 10.0 12.5 15.0 Inten.(x1,000,000) 439.1821 1.5 1.0 980.3952 0.5 302.1025 490.6986 0.0 250 500 750 1000 54 1250 1500 3. Ac-LEHD-SG2-alkyne m AU(x1,000) 4.0 214nm ,4nm (1.00) 3.0 * 2.0 1.0 0.0 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in 1750 m /z 17.5 m in 1750 m /z 17.5 m in 1750 m /z Inten.(x1,000,000) 5.0 488.1907 2.5 439.1803 302.1010 0.0 250 500 750 1000 1250 1500 4. Ac-FR-SG2-alkyne m AU(x1,000) 4.0 214nm ,4nm (1.00) 3.0 2.0 * 1.0 0.0 0.0 2.5 5.0 7.5 10.0 12.5 15.0 Inten.(x1,000,000) 392.6804 1.5 1.0 0.5 138.0850 0.0 250 500 750 1000 1250 1500 5. Ac-FRR-SG2-alkyne m AU(x1,000) 1.00 214nm ,4nm (1.00) 0.75 0.50 * 0.25 0.00 0.0 2.5 5.0 7.5 10.0 12.5 15.0 Inten.(x1,000,000) 1.0 470.7210 0.5 527.7128 314.1522 0.0 250 500 750 1000 55 1250 1500 6. Ac-LLVY-SG2-alkyne m AU(x1,000) 4.0 214nm ,4nm (1.00) * 3.0 2.0 1.0 0.0 0.0 2.5 5.0 7.5 Inten.(x100,000) 7.5 10.0 12.5 15.0 17.5 m in 1750 m /z 969.4618 806.4063 5.0 2.5 485.2292 699.3995 0.0 250 500 750 1000 1250 1500 Figure 3.3. a) General structures of the 2 products found corresponding to the 2 major peaks in the LC profiles. The major side product resulted from the direct acylation of the first FmocAA-OH onto the resin-bound secondary amines which did not react with Fmoc-SG2-COCl. b) LC-MS profiles of the 6 synthesized Ac-X-SG2-alkyne. LC conditions: 30-100% CH3CN in 15 min. 3.3.2 Chemical Synthesis of Localization Sequences Synthesis of the localization sequences was straightforward, using standard solid-phase peptide synthesis on Rink amide with Fmoc chemistry. After peptide elongation, the N-terminus of each peptide is capped with either 4-azidobutanoic acid to install an azide or acylated with the acid chloride of Fmoc-SG2-COOH as described in Chapter 3.3.1 to obtain a dye-peptide conjugate. This latter series of peptides will be referred to as the “control peptides”. Synthesis of the localization sequence 4 followed a different route due to the need for attachment of the palmitoyl group. The Rink amide resin was first coupled with Fmoc-Lys(Mtt)-OH followed by Fmoc deprotection and Fmoc-Lys(Boc)-OH. Deprotection of the terminal Fmoc group followed by acylation with 4-azidobutanoic acid or Fmoc-SG2-COOH gave the 56 respective N-capped dipeptides. The extremely acid-labile Mtt group was selectively cleaved in the presence of the Boc group with 1% TFA and 5% TIS, exposing a free amine for coupling with palmitic acid. For the azide-functionalized localization peptide, the desired lipid-modified dipeptide was obtained directly by cleavage from the solid support using standard cleavage conditions, while the control peptide 4 required an additional Fmoc deprotection step before cleavage. LC-MS analysis for the azido-peptides showed that the desired peptide was obtained in good purity (single LC-MS peak). The SG2-conjugated control peptides however, showed more impurities especially for the 2 CPPs. During the acylation step with Fmoc-SG2COOH (in both the synthesis of the control peptides and the peptide substrates), it was found that the excess acid chloride could not be washed off easily. A washing cocktail of 1% TFA/DCM was found to be effective in removing most of the excess reagent, but this washing procedure was apparently less effective for the longer peptides, as evidenced by the appearance of several other peaks which showed strong absorbance in the 490 nm channel, which is approximately at the absorbance peak of the fluorophore. a) i) N3-KKKRKV-NH2 (LC conditions: 0-50% ACN in 15 min) m AU 214nm ,4nm (1.00) 1500 1000 500 0 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in 1750 m /z Inten.(x1,000,000) 448.8013 5.0 2.5 299.5372 896.5987 0.0 250 500 750 1000 57 1250 1500 ii ) N3-RrRK-NH2 (LC conditions: 0-50% ACN in 15 min) m AU 214nm ,4nm (1.00) 1000 750 500 250 0 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in 1750 m /z Inten.(x1,000,000) 7.5 363.2291 5.0 2.5 242.4883 420.2232 0.0 250 500 750 1000 1250 1500 iii )N3-FxrFxK-NH2 (LC conditions: 25-100% ACN in 15 min) m AU 214nm ,4nm (1.00) 400 300 200 100 0 -100 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in Inten.(x10,000,000) 1.00 360.2431 0.75 0.50 0.25 719.4798 0.00 250 500 750 1000 1250 1500 1750 m /z iv) N3-KK(palmitoyl)-NH2 (LC conditions: 50-100% ACN in 15 min) m AU(x1,000) 214nm ,4nm (1.00) 3.0 2.0 1.0 0.0 0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 m in 1750 m /z Inten.(x100,000) 623.4997 5.0 2.5 0.0 250 500 750 1000 58 1250 1500 v) N3-RKKRRQRRR-NH2 (LC conditions: 0-50% ACN in 15 min) m AU(x100) 214nm ,4nm (1.00) 5.0 4.0 3.0 2.0 1.0 0.0 2.5 Inten.(x1,000,000) 1.25 5.0 7.5 10.0 12.5 15.0 17.5 m in 1750 m /z 17.5 m in 1750 m /z 483.9669 484.2985 1.00 0.75 0.50 0.25 725.4472 363.2295 0.00 250 500 750 1000 1250 1500 vi) N3-RRRRRRRRR-NH2 (LC conditions: 0-50% ACN in 15 min) m AU(x1,000) 214nm ,4nm (1.00) 1.00 0.75 0.50 0.25 0.00 0.0 2.5 5.0 7.5 10.0 12.5 15.0 Inten.(x100,000) 512.3185 7.5 5.0 664.3030 2.5 384.4913 0.0 250 500 750 1000 1250 1500 b) i) SG-SV40 m AU(x100) 214nm ,4nm (1.00) 5.0 2.5 0.0 0.0 2.5 5.0 7.5 10.0 12.5 m in Inten.(x1,000,000) 390.2346 4.0 584.8486 3.0 2.0 1.0 0.0 200 292.9281 300 400 500 600 59 700 800 900 m /z ii) SG-RrRK m AU(x1,000) 1.0 214nm ,4nm (1.00) 0.5 0.0 0.0 2.5 5.0 7.5 10.0 12.5 m in Inten.(x1,000,000) 4.0 333.1855 499.2766 3.0 2.0 556.2700 1.0 0.0 200 iii) 302.1059 371.1815 300 613.2640 400 500 600 700 800 900 m /z SG-FrFK m AU(x1,000) 214nm ,4nm (1.00) 1.0 0.5 0.0 0.0 2.5 5.0 7.5 10.0 12.5 m in Inten.(x1,000,000) 496.2844 5.0 331.1914 2.5 0.0 200 iv) 302.0978 388.1258 300 400 500 600 700 800 900 m /z SG-Tat m AU(x100) 214nm ,4nm (1.00) 5.0 2.5 0.0 0.0 2.5 5.0 7.5 10.0 12.5 m in Inten.(x100,000) 574.6574 431.2427 5.0 650.6475 688.9819 2.5 459.7349 345.1958 0.0 200 v) 300 400 726.6476 522.9585 500 600 SG-R9 60 700 800 900 m /z m AU(x100) 214nm ,4nm (1.00) 2.0 1.0 0.0 0.0 2.5 5.0 7.5 10.0 12.5 m in Inten.(x100,000) 457.1894 5.0 2.5 0.0 200 792.6532 602.6589 324.9160 300 400 500 600 700 800 900 m /z Figure 3.4. LC-MS profiles of a) azido-localization peptides; b) control peptides 3.4 Bioimaging of Control Peptides As a preliminary examination of whether the localization sequences can act as carrier modules, we synthesized dye-peptide conjugates to enable us to visualize the intracellular locations of these conjugates. Trackers that mark a specific organelle were used to determine if the conjugates have the intended localization. MCF-7 and HeLa cells were used for imaging as these are well-studied cell lines in which the localizations of some of the conjugates have been experimentally determined (with the use of other dyes instead of SG2). The SV40 and RrRK localization sequences were found to be toxic to HeLa cells at a concentration of ~ 10 µM as evidenced by the change in cell morphology after 1 h of incubation with the peptide. The 2 localization sequences were thus not used for further studies. 61 a) MCF7 cells Control peptide (GFP) Organelle Tracker (RFP) FxrFxK KK(palmitoyl) Tat R9 62 Overlay b) HeLa cells Control peptide (GFP) Organelle Tracker (RFP) Overlay FxrFxK KK(palmitoyl) Tat R9 Figure 3.5. Fluorescent images of control peptides and corresponding organelle stains. a) MCF7 cells; b) HeLa cells. The following organelle stains were used for the corresponding peptides: FxrFxK – MitoTracker® Red; KK(palmitoyl) – CellMask TM Orange; Tat and R9 – LysoTracker® Red. Images were taken with a 60× oil immersion objective. 63 Fluorescence images of the control peptides showed that the mitochondria- and membrane-targeting peptides localize as expected in both HeLa and MCF7 cells, but the two CPPs, Tat and R9 show mostly endosomal localization, with the exception of the Tat control peptide in HeLa cells, which shows some nuclear localization. In these initial experiments, we could not achieve nuclear targeting of the dye-peptide conjugate uniformly throughout the cells under observation. It is a well-known problem that these CPPs tend to localize in different organelles of the cell, depending on various factors such as the type of cell line, the cargo and the uptake conditions [67]. Endosomal entrapment of cargo resulting from CPP-mediated cellular uptake is common, resulting in inefficient delivery of the cargo to its intended location. This in part gives us a tool to effectively deliver CPP-conjugated protease substrates (targeting the cathepsins, for example) to the endosomes and the lysosomes if they remained trapped in the vesicles. Nuclear targeting however, may be achieved through optimizing uptake conditions, using other nuclear localization sequences, or more effectively, the use of methods enabling endosomal escape. While these imaging experiments act as controls to ensure that the intended localizations are achieved, they do not necessary imply that subcellular delivery of the peptide remains unchanged when conjugated to another peptide. This is due to the alteration of the overall charges and lipophilicity of the localization peptide, which may affect localization to certain organelles. The mitochondria-penetrating peptide, for example, relies on a balance of cationic charges and lipophilicity for permeating the charged membrane and entry into the hydrophobic intermembrane space. Because the mechanisms of entry into the cell and the mitochondrial membrane are not well understood, the attachment of charged peptides could lead to an unpredictable change 64 in localization. Our studies in this chapter will elucidate whether these relatively unexplored localization peptides (FxrFxK, KK(palmitoyl)) can be used as general targeting modules for peptides. 3.5 Current Work Our initial bioimaging experiments with the localization peptides showed that we could successfully deliver the peptides to the mitochondria, plasma membrane and the endosomes or lysosomes, but not efficiently to the nucleus. We are currently working on optimizing the conditions for the cellular uptake of the 2 general CPPs, as well as the SV40 NLS, which has been used as a delivery vehicle for peptide sequences. An alternative is to enable direct membrane translocation of the localization peptide, instead of through endocytosis which plays a major role in cellular uptake. Futaki’s group has shown that negatively charged, highly hydrophobic molecules can alter the electronic properties of oligoarginines through electrostatic interaction such that the pathway to cellular internatlization is altered [68]. The group subsequently showed that pyrenebutyrate mediated the delivery of a fluorescently-labeled octaarginine peptide directly into the cytosol and nucleus in HeLa cells [69]. This may be a potential solution to achieve direct cytosolic / nuclear delivery of our CPPs. An alternative is to micro-inject the NLS into the cell, which also evades the problem of endosomal entrapment. Further to the problem of the use of these CPPs is the cellular toxicity of these highly charged, cationic peptides still remains an important issue to be addressed. Specifically, we need to determine the concentration at which the peptides become cytotoxic. 65 We have also completed the solid-phase synthesis of the azide-functionalized localization peptides and the SG2-based peptide substrates targeting various cysteine proteases. To further optimize the synthetic procedures described herein, Fmoc-SG2COCl may be used in larger molar ratio (4 equiv instead of 2.5) to drive the acylation reaction with the resin-bound secondary amine to completion. In addition, an additional capping step with acetic anhydride to consume the unreacted secondary amines will prevent the formation of the peptide substrate without conjugation to SG2. This should give a higher yield and purity for each peptide. Currently, with the exception of SG2-LLVY-alkyne which needs further purification, the alkynesubstrates can be used directly for click chemistry with the azido-peptides as they are sufficiently pure for the reaction to be monitored by LC-MS. We are currently optimizing the conditions for the “click” assembly of the different peptide components to furnish the final targeted peptide substrates. Several experiments have also been designed which could give a preliminary evaluation of our approach in live-cell fluorescence imaging of protease activity in subcellular compartments. The proof-of-concept experiments will begin with incubating either HeLa or MCF7 cells with the cathepsin substrates (Ac-FRR-SG2 or Ac-FR-SG2) and observing the fluorescence image. Since the target lysosomal cysteine proteases are constitutively active, we should detect fluorescence outputs localized in the endosomes / lysosomes, which may be detected using the organelle tracker, LysoTracker®. Addition of general cysteine protease inhibitors, such as E-64, should lead to a fluorescence decrease. 66 To study caspase activity, we will incubate HeLa cells with Ac-DEVD-SG2FxrFxK and Ac-DEVD-SG2-Tat followed by apoptosis induction using staurosporine. If caspase activity is present in the corresponding organelles, we should observe a fluorescence increase after induction resulting from caspase cleavage of the substrate. The localization of this fluorescence increase should correlate well with organelle trackers that are known to be highly specific. Again the fluorescence increase should be significantly suppressed by the addition of either pan-caspase inhibitors, such as zVAD-fmk, or inhibitors specific for caspase-3/-7. Similar experiments for the study of calpain activity are also planned. The external stimulus is the calcium ionophore, ionomycin, which activates calpain I and II. Cells loaded with the targeted peptide substrates Ac-LLVY-SG2-Tat or Ac-LLVYSG2-KK(palmitoyl) should display fluorescence increase in the nucleus and plasma membrane respectively after calcium induction. The addition of calpastatin peptide (specific calpain I/II inhibitor) prior to calcium induction will serve as a positive control, similar to the experiments for cathepsins and caspases. We are also looking at the possibilities of multiplexed imaging – detecting protease activity in different organelles in the same cell, and/or imaging different proteases simultaneously. This would be useful in systems where there is significant cross-talk between the different classes of proteases in a biological event. Apoptosis, for example, involves the caspases, as well as the calpains and cathepsins, with each different class contributing to (or in some cases, inhibiting) the apoptotic cascade. It will therefore be useful to develop tools that enable the real-time imaging of different critical proteolytic events. Multiplexing may be accomplished by the using 67 fluorogenic peptides that can be detected at a different fluorescence channel, for example, the blue DAPI channel. A candidate fluorophore would be ACC, since the synthesis of ACC-based peptide substrates are well-established and can readily be adapted to our strategy. The aim of the current work presented herein is to establish a previouslyunexplored platform for the real-time imaging of protease activity localized in distinct organelles in live cells. This platform will be complementary to well-established methods of detecting proteins such as immunofluorescence and subcellular fractionation followed by western blotting. Our approach also carries the advantage of imaging protease activity which is directly correlated to its function, instead of merely registering the presence of the target protease. 68 CHAPTER 4 DISCOVERY AND DEVELOPMENT OF FLUOROGENIC LABELS FOR BIOMOLECULES 4.1 Fluorogenic Labeling of Biomolecules The green fluorescent protein (GFP) has become an invaluable tool in molecular and cell biology since its inception about 15 years ago as a tool for fluorescently tagging proteins of interest in their native environments [70]. Accompanied by parallel advances in imaging technologies, the fluorescent protein toolbox has expanded with the inclusion of variants of various fluorescing wavelengths that span the visible spectrum and with different applications. Genetic fusion of the fluorescent protein (FP) tag to the protein of interest ensures that labeling is specific to the protein under study, but offers no control over when and where visualization of the protein is most desired. This lack of spatiotemporal control for the study of protein dynamics has led to the development of other imaging techniques, such as fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) which involve the eradication of fluorescence in a region of interest by intense laser illumination. With increasing sophistication in imaging technologies, there is growing interest in photoactivatable, photoconvertible and photoswitchable FPs, collectively termed as optical highlighters [71], in which the fluorescence of the FP can be turned on or shifted to another region of the visible spectrum by light. These “highlighters” create a distinct population of proteins for selective visualization and the study of protein dynamics [72]. 69 Despite these advances, the major drawback using FPs as a means of visualizing proteins lies in their size – the modification with a large (~ 27 kDa) protein tag by genetic fusion inevitably has the potential problem of interfering with the endogenous function of the protein. In addition, the optical and physical properties of FPs which are photoresponsive that have been developed so far still fall short of being optimal for routine usage in imaging experiments. There is clearly a need for alternative approaches that could give the experimenter direct control over the labeling event. In recent years, a new dimension to protein labeling has emerged with the advent of using small molecule tags for site-selective protein modification [73]. The seminal work published by Tsien and co-workers in 1998 describes the specific covalent labeling of enhanced cyan fluorescent protein (ECFP) genetically fused with a tetracysteine peptide binding motif employing a biarsenical fluorescein analog as the ligand [74]. Known as the FlAsH (Fluorescein Arsenical Helix binder) ligand, the authors successfully demonstrated the occurrence of FRET between ECFP and FlAsH in live mammalian cells. This work was significant in a number of ways. It was the first example of using a small-molecule ligand to label a protein at a specific site in an in vivo setting. This allowed temporal control of the labeling event, a major advantage over genetically encoded FP tags. In addition, modification of the protein of interest by a short peptide instead of a large FP considerably alleviated the problem of functional interference caused by the tag. Tsien’s group further introduced the redand blue-fluorescing versions of FlAsH, ReAsH and ChoXAsH respectively, and optimized the peptide motif for increased binding affinity to the biarsenical ligands 70 [75, 76]. These findings collectively set the direction for future developments in the field of in vivo protein labeling. A recent trend in the field of protein labeling is the use of fluorogenic dyes which register a dramatic increase in fluorescence upon non-covalent or covalent interaction with the protein of interest. The advantage conferred by these dyes is clear – these molecules serve as self-reporting indicators of the binding or labeling event and the background fluorescence contributed by unbound or unreacted dyes is kept to a minimum. This is particularly important in bioimaging as it leads to a higher signalto-noise ratio which translates to the more sensitive detection of cellular proteins, especially less abundant proteins. This concept was in fact first demonstrated by the FlAsH family of biarsenical ligands which fluoresce strongly only upon binding to the tetracysteine motif but not when it is a bis 1,2-ethanedithiol (EDT) adduct, the form which is used for labeling. However the de novo design of these dyes is not trivial and requires substantial experimentation. Though there are notable examples of novel fluorogenic dyes, the majority of the fluorogenic labels to date rely on the appendage of other functional groups such as fluorophore quenchers to confer latent spectral properties which are activated upon labeling. Fluorogenic labels for biomolecules developed to date may be classified into two categories: i) structurally new fluorogenic dyes and ii) an internally quenched fluorophore where the quenching mechanism is inactivated during the labeling reaction. The fluorescence activation is typically initiated by light or by a highly specific reaction, both of which result in a covalent bond between the dye and the biomolecule. One of the most prominent examples in the former category is the 71 family of “click” coumarins reported independently by the Wang and Fahrni groups. [77] The two groups introduced the concept of fluorogenic “click” reactions in which a weakly fluorescent azido- or alkyne-coumarin is converted into a fluorescent molecule by triazole formation using a Cu(I)- catalyzed azide-alkyne cycloaddition, more commonly known as “click” chemistry. This unique feature, coupled with the bioothorgonal nature of the cycloaddition reaction, has found useful applications in the fluorescence labeling and visualization of glycans [78], newly synthesized proteins [79] and lipids [80]. However, the need for the toxic Cu(I) has limited the use of this fluorogenic bioconjugation method in bioimaging to fixed cells, thereby excluding it from the study of protein dynamics. A potential improvement of the fluorogenic “click” reaction was presented by Lin and co-workers who utilized a photoinduced 1,3-dipolar cycloaddition reaction between an in situ formed nitrile imine from a tetrazole precursor and an alkene, termed as “photoclick chemistry”. The pyrazoline adduct formed was fluorescent, allowing the reaction to be monitored by the fluorescence readout. The authors successfully demonstrated the use of this fluorogenic reaction in functionalizing a protein with a genetically incorporated Oallyl tyrosine residue in live bacterial cells [81]. Another approach to fluorogenic labeling relies on the use of fluorescence quenching systems in which the fluorophore remains spectrally silent until the quenching mechanism is disengaged when labeling occurs. The Bertozzi group first developed a coumarin-phosphine dye whereby the lone pair of electrons quenches the excited state of the coumarin fluorophore [82]. Phosphine oxidation eliminates quenching, leading to a dramatic fluorescence increase. This oxidation reaction was brought about by Staudinger ligation of the coumarin-phosphine to an azide- 72 functionalized protein, such that labeling of the protein was accompanied by a fluorescence increase. However, non-specific auto-oxidation of the phosphine group reduced ligation efficiency and fluorescence output. To overcome this problem, the group recently developed a FRET-based fluorogenic phosphine attached to fluorescein in which a quencher is released upon ligation with an azide, thereby releasing the fluorescence [83]. This design allows for the incorporation of different dye / quencher pairs for multicolor labeling. While applicable to live-cell imaging, the Staudinger ligation has slower reaction kinetics than “click” chemistry, thus limiting its dynamic range of usage. The abovementioned examples utilize a highly specific bioorthogonal chemical reaction for labeling with concomitant fluorescence increase. Another contribution in the development of fluorogenic labeling is the use of enzymatic reactions to label a protein with a fluorogenic probe. Kikuchi and co-workers designed a β-lactam FRET probe that is catalytically trapped by a mutant β-lactamase, and demonstrated the labeling of a membrane-associated protein fused to the enzyme in live cells [84]. In a different approach, the groups of Bogyo [85] and Nagano [86] used quenched and FRET activity-based probes (ABPs) respectively to specifically label and visualize their target enzymes. With the development of these probes, it is likely that future ABPs will be designed for use in bioimaging applications, a direction which is previously unexplored in the field of enzyme profiling by ABPs. A brief survey of fluorogenic labeling methods reveals the need for the development of new fluorogenic reagents and the improvement of current ones to 73 enable a broad range of bioimaging applications. This chapter deals with our approach in our search of fluorogenic labels – fluorescence activation by “click” chemistry. 4.2 Combinatorial Discovery of Fluorophores Small organic dyes and fluorescent probes are well-established labeling agents and sensors in both chemical and biological systems [86]. Despite their wide-ranging applications and popularity in reporter assays and visualization tags, the underlining photophysical properties of some of these dyes in relation to their molecular structures are not yet well understood. Consequently, the rational design of novel fluorophores possessing highly predictable and desirable properties has remained elusive. The lack of well-defined rules to govern fluorophore design has driven combinatorial efforts in fluorophore discovery in recent years [87]. These efforts have been aided by the use of novel synthetic methodologies to construct novel fluorophore cores [88]. However, fluorophore discovery is often not the end purpose in these cases but rather a serendipitous finding in the process of developing synthetic methodologies. A more directed, systematic approach to combinatorial fluorophore synthesis is the modification of core structures from known fluorophores to furnish analogs of the parental molecules. This has thus far led to the discovery of new fluorescent molecules with a range of spectroscopic properties. One notable method for the rapid discovery of fluorophores is the use of the Cu(I)- catalyzed azide-alkyne cycloaddition, the representative reaction in “click” chemistry [89], to assemble a variety of structurally-related fluorophores. The Wang and Fahrni groups independently introduced the concept of fluorogenic “click” 74 reactions in which a weakly fluorescent azido- or alkyne-coumarin is converted into a fluorescent molecule by triazole formation using “click” chemistry [77]. At present, only the coumarins [77], carbostyrils [90], anthracenes [91], naphthalimides [92] and pyridyloxazole [93] mimics comprise the family of “click” fluorophores in which “click” chemistry has been used as a fluorogenic reaction and/or for diversification to generate analogs of the parental fluorophore. Type of Fluorophore Parent Fluorophore Modified “Click” Fluorophores COOH X O O O Coumarins O N R1 N N N N R2 O N O OH O X = OH, NH2 O N Carbostyrils O N N N R1 O N N N N R1 O R N H Pyridyloxazoles R2 R2 R1 R2 R1 R2 O O N N N N N R3 N N N N N N N N R1 Anthracenes R2 R1 R3 R3 n R2 n = 0, 1 O Naphthalimides R O NH2 N N N O O N N O N R N N N O Figure 4.1. Fluorophore types which have been synthesized using “click” chemistry One major drawback of the current “click” fluorophores is that all of them are UVexcited dyes, making them undesirable choices for bioimaging applications where 75 cells or tissues are used. The key aim in the current work is twofold. We wish to extend the “click” chemistry-mediated discovery of fluorescent dyes to previously unexplored fluorophore scaffolds, especially those with excitation wavelengths in the visible range. In addition, we hope to find new scaffolds that can be used as fluorogenic labeling reagents for bioimaging applications. 4.3 Design of Xanthone- and Xanthene-based “Click” Fluorophores Recently, our group introduced a new fluorophore, Singapore Green [22], a structural hybrid of Tokyo Green (a fluorescein analog) [37] and Rhodamine 110 with similar emission and excitation properties to both (Figure 4.2). We reasoned that replacement of the oxygen electron donor at the 6’ position with an alkyne in both Singapore Green and Tokyo Green will significantly decrease the fluorescence output of their xanthene core. We further extended this design to Rhodamine B by similarly substituting the diethylamino group at the 6’-position with an alkyne, as well as replacing the carboxylic acid moiety in Rhodamine B with a methyl group at the 2position to lock the xanthene core in the conjugated quinol-iminium form. We anticipate that the formation of a triazole ring at this position using “click” chemistry will result in a fluorescence change in these xanthene-alkynes through an extended πconjugated system, and that this change can be tuned by the use of azides with different electronic properties. In the interest of extending the emission range of our “click” fluorophores from blue to the yellow region, we also used the blue-light emitting xanthones which are synthetic precursors of our xanthenes (Figure 4.2). 76 Known fluorophores O 1 O 7 3 4 O 5 6 OH X 3,6-dihydroxy xanthone 3' Y New "click" fluorophores O 8 2 HO Alkyne pro-fluorophores 4' O 5' 6' O X X = OMe, NH2, NEt2 Y 1' N N R1 O "Click" assembly in microplate 7' 2' O R N3 8' N N Y O N R N R 2 Singapore Green: Y = NH, R1 = OMe Tokyo Green: Y = O, R1 = OH Y = O, NH, NEt2+Cl- Figure 4.2. Design of xanthone- and xanthene-based “click” fluorophores from known fluorophores. Similar to the design of our xanthene-alkynes, we replaced the heteroatom at the 6-position with an alkyne to yield the xanthone-alkyne for “click” modification. We noted that while there are several reports on the synthesis and spectroscopic characterizations of rosamine dyes from 3,6-disubstituted xanthones [93, 94], to the best of our knowledge there are no detailed studies on xanthone-based fluorophores and their fluorescence properties. 4.4 Chemical Synthesis of Xanthone- and Xanthene-based “Click” Fluorophores 4.4.1 Chemical Synthesis of Xanthone- and Xanthene-Alkynes and Azides The general synthetic strategy for the alkynes A, B, D and E involves the desymmetrization of the common starting material 3,6-dihydroxyxanthone 4-1 to give the appropriate substituent at the 6-position, leaving the other phenolic group for conversion into a triflate which serves as the substrate for Sonogashira coupling with trimethylsilylacetylene. Deprotection of the TMS group affords alkynes A and B, while Grignard addition to the xanthone followed by removal of the protecting groups gave alkynes D and E. 77 O NaOH X MeOH/H2O O R O TMS PdCl2(PPh3)2, CuI O OTf NEt3, DMF R 4-2a: R = OMe 4-2b: R = NEt2 O A: X = OMe (90%) B: X = NEt2 (92%) O 4-3a: R = OMe (60%) 4-3b: R = NEt2 (81%) TMS 1. Y O MgBr THF, 50 οC 2. deprotection D: Y = O E: Y = NEt2+Cl- Scheme 4.1. General synthetic strategy towards alkynes A, B, D and E. To synthesize 4-2a, 4-1 underwent monomethylation in the presence of 1 equiv of K2CO3 which acts as a base to deprotonate the phenol followed by conversion into a triflate. 4-2b was synthesized by direct substitution of the ditriflate formed from 4-1 (Scheme 2.2) with diethylamine. As reported in recent literature [27], the presence of an electron-withdrawing carbonyl group in the para position activates the triflate group towards nucleophilic aromatic substitution, making this reaction feasible under conditions that are milder than normally required for similar reactions in other substrates. The reactivity of the xanthone unit is considerably reduced after monosubstitution as the electron-donating diethylamino moiety deactivates the ring. A prolonged reaction time (16 h) however led to significant formation of the O Me2SO 4, K2CO3, O HO O 4-1 DMF 43% O Tf2O, pyridine O O OH OH CH2Cl2 O 84% 4-1i CH2Cl2 91% OTf O O Tf2O, pyridine O 4-2a Et2NH, DMSO, TfO O 4-1ii OTf 90 οC 43% Et2N O 4-2b OTf Scheme 4.2. Synthesis of 4-2a and 4-2b from 1 disubstituted product 4-2bi. Attempts to install the alkyne moiety by direct nucelophilic substitution of the second triflate by sodium acetylide were unsuccessful, giving compound 4-2bii instead. The triflate group was removed during the reaction, 78 probably due nucleophilic attack at the sulfur atom instead of the sp2 aromatic carbon. In a separate experiment, the reaction between 4-1ii and NaN3 in DMSO at 70οC similarly led to the removal of one triflate group. In the reaction with diethylamine, only a trace amount of 4-2bii was obtained (Figure 4.3). O a) Et2N O 4-2b O Nu- O O S O CF3 Et2N O- O O O b) O Et2N Et2N O O 2bii O OH HO O OTf TfO O 4-2b OH O OTf 4-1ii Et2N c) O O N3 O O O O OTf O Et2NH, DMSO, TfO O 4-1ii OTf 90 οC, 16 h Et2N + O OTf 4-2b + Et2N O 4-2bi NEt2 Et2N O OH 4-2bii Figure 4.3. Undesired products obtained during the nucleophilic aromatic substitution of 2b and 1ii with different nucleophiles. a) Mechanism of nucleophilic attack leading to a formal hydrolysis of the triflate. b) Formation of undesired phenols with sodium acetylide and NaN3 as nucleophiles. c) Various products observed from the prolonged reaction of 1ii and diethylamine. A possible explanation for this pattern of reactivity lies in the nucleophilicity of the attacking species; the acetylide and azide anions which are charged and unhindered, are strong nucleophiles compared to diethylamine which is neutral and relatively more hindered. These nucleophiles can attack the more hindered sulfur atom in the triflate group to displace the phenoxide anion as the leaving group. Consequently alkynes 4-3a and 4-3b had to be synthesized by Pd(0)-catalyzed Sonogashira coupling in the presence of CuI. 79 The synthesis of alkynes C and F followed a strategy similar to that employed for alkynes A, B, D and E. Starting from 3-nitro-6-hydroxyxanthone 4-4, it was converted to triflate 4-5 which underwent Sonogashira coupling to give the TMSprotected nitroxanthone 4-6 (Scheme 4.3). The nitro group was reduced under mild conditions to aniline 4-7 with zinc in MeOH/THF buffered at pH 5. Under these conditions, the labile TMS group was preserved. Aniline 4-7 was protected with a trityl protecting group, followed by Grignard addition to give the trityl-protected xanthene 4-9. Subsequent deprotection of the trityl and TMS groups furnished the final alkyne F. Alkyne C was obtained after deprotection of the TMS group in 4-7. O O Tf2O, pyridine O 2N O 4-4 OH CH2Cl2 O2N (87%) O 4-5 OTf NEt3, DMF O (73%, 2 steps) TrtHN TMS 1. O 4-8 O 4-6 TMS TMS HN O , THF, 50 οC 2. TFA/CH2Cl2/H2O 7:2:1 (55%, 2 steps) TMS NaOH MeOH/H2O 4-9 (81%, 2 steps) HN O NaOH, MeOH/H2O 76% O H2N O 2N MgBr CPh3Cl, NEt3, CH2Cl2 O 4-7 Zn, sat. NH4Cl, MeOH/THF 3:1 (65%) O H2N O TMS PdCl2(PPh3)2, CuI O C F Scheme 4.3. Synthesis of alkynes C and F. The azides used in the study were selected from a panel of azides previously synthesized and reported by our group. To investigate the influence of the electronic properties on the azides on the fluorescence properties of the fluorophores, both electron-rich and electron-deficient aromatic azides were used, including halogenated and heterocyclic azides. A few structurally different aliphatic azides were used as well 80 to test if peripheral groups that are electronically decoupled from the fluorophore core could contribute a change to fluorescence properties. Aromatic azides N3 N3 N3 N3 N3 N3 Cl O O z1 O N O z3 z2 electron-rich azides Cl z5 N3 N N3 N3 z13 z12 z11 heterocyclic azides O H N NO2 z8 electron-deficient azides others Aliphatic azides z7 N z10 N3 N3 CO2Et F z6 N3 N3 z9 Br O z4 F O HN N3 O O z14 N3 N3 z15 z16 HOOC O O S N H N3 z17 Figure 4.4 Structure of the azides used in this study. Aromatic azides that were not previously by our group were synthesized by a simple diazotization reaction of the precursor aniline followed by substitution by NaN3 as previously reported by our group (Scheme 4.4). NH2 N3 1. 2N HCl, NaNO 2 R 2. NaN3 R Scheme 4.4 Synthesis of aromatic azides from anilines A total of 17 azides were picked for the Cu(I)- catalyzed [3 +2] cycloaddition with the 6 alkynes to give a 102-member library of xanthone- and xanthene-based fluorophores. 81 4.4.2 Microplate-Based Assembly of Fluorophores Using “Click” Chemistry O O X N3 X Y O A-C O z1-z14 + O A-z1 - C-z17 R CuSO4 (2 equiv) Sodium Ascorbate (5 equiv) DMSO/tBuOH/H2O R N3 z15-z17 N R N N N N Y O N R 12 h D-F D-z1 - F-z17 Scheme 4.5. “Click” Assembly of Fluorophores With the 6 alkynes and 17 azides in hand, we assembled the fluorophore library in a 384-deep well block by mixing each of the alkynes with a different azide in DMSO/tBuOH/H2O to give a unique pair of alkyne and azide in each well. The “click” reaction was slow when CuSO4 and sodium ascorbate which generated the active Cu(I) species were used in sub-stoichiometric amounts (0.2 and 0.5 equiv respectively) in several solvent combinations such as tBuOH/H2O and DCE/H2O, such that very little or no product was formed after 24 h. We then tried a combination of CuSO4 (2 equiv) and sodium ascorbate (5 equiv) for the reaction. Under these conditions, the “click” reaction for the xanthene-alkynes D-F proved to be highly efficient, with the alkynes consumed within 12 h to give the products in high purity as monitored by LC-MS. The xanthone-alkynes A-C were considerably less reactive with some incomplete reaction after 12 h. The desired product was obtained for all compounds with the exception of products with azides z13 and z14 which did not give the desired products for all the alkynes and were thus omitted from subsequent studies. LC-MS analysis was carried out on the entire library to ensure that the library is of sufficient quality for in situ fluorescence screening. Selected profiles are shown in below. 82 A-z3 mAU(x100) 3.0 SPD Ch1:254nm O OMe 2.0 O OMe O N N N Exact Mass: 459.14 1.0 OMe 0.0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 min Inten.(x100,000) 2.0 460.00 1.0 919.20 538.15 186.95 221.95 0.0 402.00 315.95 200 300 400 500 559.05 610.35 661.95 600 700 761.10807.70 859.00 800 900 957.25 m/z A-z6 mAU(x100) 1.5 SPD Ch1:254nm O * F 1.0 O O 0.5 Exact Mass: 405.09 F N N N 0.0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 750.85 811.05 800 874.75 900 min Inten.(x100,000) 5.0 405.95 2.5 447.00 484.00 0.0 183.90 262.85 200 524.40 500 364.10 400 300 600 641.20687.95 700 952.05 m/z B-z7 mAU(x100) SPD Ch1:254nm O CO2Et 5.0 Et2N 2.5 O N Exact Mass: 482.2 N N ** * 0.0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 min Inten.(x100,000) 5.0 483.10 2.5 161.90 262.05 0.0 200 300 524.15 352.05 408.75455.15 400 500 600 83 687.30 700 774.80 800 902.05 900 996.30 965.35 m/z B-z16 mAU(x100) 1.5 SPD Ch1:254nm O O ** 1.0 Et2N NH O N Exact Mass: 507.23 N N 0.5 0.0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 min Inten.(x100,000) 1.5 508.10 1.0 160.00 0.5 208.15 0.0 303.10 200 380.15 300 452.10 400 981.80 550.00 602.30 663.15 500 600 700 783.95 839.90 800 902.85 900 m/z C-z1 mAU(x1,000) SPD Ch1:254nm O 1.0 H2 N 0.5 O Exact Mass: 384.12 OMe N N N 0.0 0.0 1.0 2.0 Inten.(x100,000) 3.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 min 384.95 2.0 426.05 1.0 769.10 356.95 171.05 463.10507.55 559.55 611.45 500 600 313.05 0.0 200 300 400 736.30 700 869.20 900 800 978.15 m/z C-z9 mAU(x100) 3.0 SPD Ch1:254nm ** O Br 2.0 H2 N O N Exact Mass: 446.04 N N 1.0 0.0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 min Inten.(x10,000) 5.0 160.95 446.95 2.5 206.90 274.05 0.0 200 300 378.55 400 489.65 526.95 571.00 500 600 84 638.90 711.80 759.75 808.70 700 800 895.65 947.50 900 m/z D-z2 mAU(x100) SPD Ch1:254nm 5.0 O N N N O 2.5 Exact Mass: 457.18 0.0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 min Inten.(x1,000,000) 3.0 458.10 2.0 1.0 0.0 182.95 200 415.20460.10 536.30 400 500 319.25 300 600 675.10 700 996.40 915.25 900 m/z 789.95 800 D-z5 mAU(x100) SPD Ch1:254nm 5.0 Cl N N O O N Cl 2.5 ** Exact Mass: 497.07 0.0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 min Inten.(x1,000,000) 2.0 498.00 1.0 0.0 182.95 240.65 200 325.00 300 392.00 400 503.05 577.70 600 500 660.00 762.85 700 826.75 800 994.95 902.70948.65 900 m/z D-z16 mAU(x100) SPD Ch1:254nm 5.0 O O N N N O N H 2.5 Exact Mass: 526.2 0.0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 min Inten.(x1,000,000) ** 1.5 527.20 1.0 0.5 175.05 0.0 223.95 200 324.90 383.10 300 400 499.15 500 605.20 655.85 749.70 600 700 800 85 857.05 986.90 932.55 900 m/z E-z1 mAU(x100) SPD Ch1:254nm 10.0 N N Et2N 7.5 O N OMe 5.0 Exact Mass: 515.24 ** 2.5 0.0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 min Inten.(x1,000,000) 515.15 1.0 0.5 186.05 0.0 306.10 300 200 396.20 400 487.15 500 585.15 635.15 600 738.95 700 886.80 941.80 900 m/z 800 E-z11 mAU(x100) SPD Ch1:254nm 10.0 7.5 Et2N O N N N 5.0 Exact Mass: 573.26 ** 2.5 0.0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 min Inten.(x1,000,000) 3.0 573.20 2.0 1.0 160.00 205.25 259.10 339.90 391.90 446.10 501.15 200 300 400 500 0.0 606.40 600 693.30738.20 700 838.00 800 981.95 933.00 900 m/z F-z9 mAU(x100) SPD Ch1:254nm 5.0 HN Br N N N O 2.5 Exact Mass: 520.09 0.0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 min Inten.(x1,000,000) 521.05 2.0 1.0 0.0 160.05 218.85264.10 335.25 200 300 414.05 400 495.00 500 986.65 563.20 610.15 600 86 700 781.90 800 880.80 900 m/z F-z10 mAU(x100) SPD Ch1:254nm 5.0 Br N N HN N ** O 2.5 Exact Mass: 520.09 0.0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 min Inten.(x1,000,000) 3.0 479.00 2.0 1.0 0.0 161.90 227.15 200 300 392.00 451.10 361.15 537.30 400 500 600 700.10 700 856.85 800 954.75 900 m/z Figure 4.5. LC-MS profiles of selected “click” fluorophores. LC conditions: 30-100% (alkynes A-C) or 20-100% (alkynes D-F) ACN in 10 min . * - unconsumed alkyne; ** - no MS found for peak, assumed to be excess azide 4.5 Spectroscopic Analysis of the “Click” Fluorophore Library For preliminary evaluation of the fluorescence properties of the library, the crude reaction mixture was diluted to 400 µM in DMSO, which was further diluted to 20 µM for fluorescence screening. Since the quantum yield of fluorophores can be significantly influenced by solvent effects, we chose four different solvents – H2O (aqueous, with DMSO as cosolvent to prevent precipitation), DMSO (polar aprotic), EtOH (polar protic) and DCE (apolar aprotic) to record the excitation and emission spectra of all the fluorophores in the library. The excitation and emission maxima for each click fluorophore are summarized in the table below. 87 Max λex Max λem Max λex Max λem A 326 429 B 389 471 C 362 487 A-z1 362 495 B-z1 389 465 C-z1 365 472 A-z2 368 489 B-z2 386 462 C-z2 362 481 A-z3 365 522 B-z3 386 453 C-z3 356 469 A-z4 356 435 B-z4 386 453 C-z4 365 481 A-z5 347 438 B-z5 389 459 C-z5 362 478 A-z6 347 447 B-z6 383 468 C-z6 362 478 A-z7 332 444 B-z7 386 468 C-z7 356 475 A-z8 - - B-z8 - - C-z8 - - A-z9 359 480 B-z9 386 462 C-z9 365 478 A-z10 359 480 B-z10 386 462 C-z10 362 478 A-z11 338 474 B-z11 386 453 C-z11 359 475 A-z12 350 423 B-z12 386 465 C-z12 365 460 A-z13 NA NA B-z13 NA NA C-z13 NA NA A-z14 NA NA B-z14 NA NA C-z14 NA NA A-z15 374 444 B-z15 392 453 C-z15 362 454 A-z16 341 423 B-z16 383 453 C-z16 359 454 A-z17 335 423 B-z17 383 450 B-z17 362 454 D 466 538, 574 E 511 589 F 483 547 D-z1 469 538, 577 E-z1 514 580 F-z1 483 550 D-z2 469 547, 577 E-z2 511 577 F-z2 486 547 D-z3 - - E-z3 505 580 F-z3 495 541 D-z4 - - E-z4 505 589 F-z4 486 547 D-z5 469 535, 580 E-z5 511 586 F-z5 486 544 D-z6 466 544, 583 E-z6 511 586 F-z6 483 547 D-z7 - - E-z7 517 589 F-z7 483 547 D-z8 - - E-z8 520 583 F-z8 483 550 D-z9 - - E-z9 514 577 F-z9 486 550 D-z10 469 547, 577 E-z10 511 577 F-z10 486 547 D-z11 469 547, 577 E-z11 517 580 F-z11 486 544 D-z12 - - E-z12 514 586 F-z12 486 544 D-z13 NA NA E-z13 NA NA F-z13 NA NA D-z14 NA NA E-z14 NA NA F-z14 NA NA D-z15 466 538, 577 E-z15 514 574 F-z15 483 547 Solvent DCE 40% DMSO /H2O Solvent DCE DCE Max λex Solvent DMSO DCE Max λem D-z16 469 541, 577 E-z16 514 583 F-z16 483 544 E17 472 538, 571 E-z17 514 580 F-z17 486 544 Table 4.1. Wavelengths of maximum excitation (λex) and emission (λem) for each “click” fluorophore. NA = not applicable. Desired MS were not found for fluorophores marked ‘NA’. Compounds marked “–” had very low fluorescence intensities and were not analyzed The wavelengths of excitation and emission were largely dictated by the type of alkyne, but there were some variations within each alkyne series which were probably contributed by the different azides. Fluorescence screening of the xanthenes reveals another interesting feature – the alkynes used were structurally similar, but they differed greatly in terms of fluorescence emission. At the same concentration, fluorophores derived from alkynes D and E were considerably less bright than those from alkyne F. The reason for this is yet unclear, but it shows that minor changes in 88 the structure while preserving the same core framework can have significant effects on fluorophore properties. From the library, several fluorophores were selected and their fluorescence spectra are shown below. DMSO (excitation at 350 nm ) DCE (excitation at 390 nm ) A-z5 A-z6 16000 B -z3 B -z8 A-z9 4000 B -z10 A-z16 B -z15 12000 RFU A B 8000 2000 4000 0 450 500 Wavelength (nm ) 0 550 450 500 DMSO (excitation at 360 nm ) 40% DMSO (excitation at 460 nm ) C-z4 25000 D-z2 C-z7 D-z6 C-z8 20000 550 Wavelength (nm ) C-z17 C D-z16 4000 D-z17 D RFU RFU 15000 10000 2000 5000 0 400 0 520 450 500 Wavelength (nm ) DCE (excitation at 510 nm ) 3000 570 620 Wavelength (nm ) DCE (excitation at 490 nm ) E-z2 E-z8 F-z5 E-z11 F-z7 E F-z12 30000 F-z17 F 2000 RFU RFU 20000 1000 0 540 10000 0 520 590 Wavelength (nm ) 89 570 Wavelength (nm ) 620 B-z15 16000 C-z17 40%DM SO DM SO 25000 DCE DM SO DCE 12000 40%DM SO Et OH 20000 Et OH RFU 15000 8000 10000 4000 5000 0 400 0 450 500 550 Wavelength (nm ) 450 500 Wavelength (nm ) F-z9 F-z12 40%DM SO 40%DM SO DM SO DM SO DCE Et OH 20000 DCE 30000 Et OH RFU RFU 20000 10000 10000 0 520 570 Wavelength (nm ) 0 520 620 570 Wavelength (nm ) 620 Figure 4.6. Emission spectra in different solvents of selected fluorophores from microplatebased fluorescence screening To quantify the fluorescence change after triazole formation, fluorescence values from each “click” product were expressed as a ratio of the fluorescence value of control (iii) which represents the fluorescence of the corresponding alkyne in each alkyne series. The final output values (ranging from -50 to 50) used for computing the values for the heat map were generated by this formula: Final output values = 10 × log2(RFUx/RFUiii) 90 S o lve nt λex λem Alkyne 370 480 A 350 480 A 390 489 40% DM S O 360 454 / H 2O 470 538 B 510 E F 370 480 A 350 480 A 390 489 B DM S O 360 454 C 470 538 D 579 E 490 547 F 370 480 A 350 480 A 390 489 B 360 454 C 470 538 D 510 EtOH 579 E 490 547 F 370 480 A 350 480 A 390 489 B 360 454 C 470 538 D 510 z3 z4 z5 z6 z7 z8 z9 z10 z11 z12 z13 z14 z15 z16 z17 i ii iii D 490 547 DC E z2 C 579 510 z1 579 E 490 547 F Fluorescence change (-) Fluorescence change (+) Figure 4.7. Heat map showing fluorescence intensities of each ‘click’ product relative to its corresponding alkyne building block. Control (i) = alkyne + CuSO4, control (ii) = alkyne + sodium ascorbate, control (iii) = alkyne only. ‘Hit’ fluorophores selected for scale-up and purification are highlighted in black boxes. Red bars indicate a fluorescence increase and Blue bars indicate a fluorescence decrease after the “click” reaction, compared to the corresponding alkyne (white) The results are summarized in the form of a heat map displaying the fluorescence intensities of each fluorophore relative to its alkyne precursor (that is, the xanthone or xanthene core prior to click chemistry) (Figure 4.5). 91 Several observations can be made from the heat map. As anticipated, the majority of the “click” products registered an increase in fluorescence intensity (red squares) over the parent alkyne, but a considerable number also led to a fluorescence decrease (blue squares). In particular, a morpholino substituent in the azide (z4, column 4) led to a consistent diminution of fluorescence throughout the alkyne series, possibly due to photoinduced-electron-transfer (PET) quenching from the nitrogen atom, and a strongly electron-withdrawing nitro substituent (z8, column 8) also effectively quenched the fluorescence in alkynes series A-D. Notably, the “click” reaction between aliphatic azides (column 15-17) and the xanthene-alkynes was fluorogenic, implying that triazole formation is itself sufficient for fluorescence activation. This feature is particularly important when searching for green lightemitting “click” fluorophores for use in labeling azide-modified biomolecules for bioimaging purposes. To examine in detail the fluorescence properties of our “click” fluorophores, we picked a few “hits” from each of the 6 alkyne series for scale-up, purification and characterizations. These “hits” were selected on the basis of the following points: (i) they showed significantly higher fluorescence intensity than their alkyne precursors; (ii) the “hits” give a good representation of aromatic and aliphatic azides of different properties; (iii) their fluorescence intensities showed some solvent sensitivity (F-z12 and F-z17). The spectroscopic properties of 4 of the brightest “hits” and their corresponding alkyne precursors were further evaluated. 92 O O N N HN Et2N N O N H2N O HN O N N N O HOOC S O O B-z15 N N N O N N N NH O H N S O O F-z12 C-z17 F-z17 HOOC Figure 4.8. Structures of the fluorophores used for quantitative fluorescence analysis. Fluorophore λmax abs (nm) εmax λem max (nm) Φf Brightness (εmax × Φf) B B-z15 C C-z17 F F-z12 F-z17 366 381 362 358 476 480 479 13,500 21,900 12,500 24,200 11,700 32,800 27,700 461 445 482 456 537 540 537 0.57 0.94 0.43 0.62 0.11 0.34 0.4 7,700 20,600 5,380 15,000 1,290 11,200 11,100 Table 4.2. Summary of spectroscopic properties of “hit” fluorophores and alkynes It was found that triazole formation led to an increase in both molar absorptivity and quantum yield, leading to an overall increase in brightness (ε × Φ f) of 2-3-fold for B-z15 and C-z17 in DMSO, and ~10-fold for F-z12 in DCE and F-z17 in EtOH (Table 4.2). F-z12 and F-z17 also displayed different solvent sensitivity despite being derivatives of the same alkyne, with F-z12 fluorescing most brightly in DCE, while in other solvents it is considerably less bright. F-z17 is the brightest in EtOH and has varying intensities in other solvents (Figure 4.8). There is however, no direct trend between the fluorescence intensities and solvent dielectric constants, suggesting that the observed solvent effects are specific to the molecular structure of the fluorophore. Because these effects shown in both our microplate screening and detailed analysis are not easily predicted, the combinatorial approach to various analogs is an advantage in searching for the desired fluorophore properties as demonstrated in this report. 93 a) A (DCE) 200 A-z6 (DCE) 200 Emission Emission Excitat ion Excitation 150 RFU RFU 150 100 100 50 50 0 300 350 400 450 Wavelength (nm ) 0 300 500 λex = 333 nm, λem = 397 nm 350 400 450 Wavelength (nm ) 500 λex = 340 nm, λem = 396 nm A-z9 (DCE) B (DCE) 400 Excitation Emission Emission Excitat ion 150 RFU RFU 300 100 50 200 100 0 300 350 400 450 Wavelength (nm ) 0 300 500 λex = 388 nm, λem = 461 nm λex = 347 nm, λem = 398 nm B-z15 (DCE) 500 400 500 Wavelength (nm ) Excitation C (DMSO) 300 Emission Excitation Emission 400 RFU RFU 200 300 200 100 100 0 300 0 300 400 500 Wavelength (nm ) λex = 385 nm, λem = 445 nm 400 500 Wavelength (nm ) λex = 376 nm, λem = 482 nm 94 C-z17 (DMSO) D (EtOH) Excitation 800 Emission 600 50 RFU RFU Excitation Emission 400 200 0 300 0 350 400 500 Wavelength (nm ) λex = 360 nm, λem = 456 nm D-z2 (EtOH) 450 550 Wavelength (nm ) λex = 473 nm, λem = 537 nm E (DCE) Excitation Emission 500 650 Emission Excit ation 300 RFU RFU 400 200 100 0 350 450 550 Wavelength (nm ) 0 400 650 λex = 472 nm, λem = 533 nm 500 600 Wavelength (nm ) λex = 504 nm, λem = 569 nm E-z2 (DCE) F (EtOH) 100 Emission Excit ation Emission RFU RFU Excit at ion 0 400 0 350 500 600 Wavelength (nm ) λex = 516 nm, λem = 577 nm 450 550 Wavelength (nm ) λex = 478 nm, λem = 537 nm 95 650 F-z12 (EtOH) 800 Emission F-z17 (EtOH) 800 Excit at ion Emission Excit at ion 600 RFU RFU 600 400 400 200 200 0 350 450 550 Wavelength (nm ) 0 350 650 λex = 481 nm, λem = 540 nm 450 550 Wavelength (nm ) 650 λex = 478 nm, λem = 538 nm b) 800 600 600 RFU RFU F-z12 800 400 400 200 200 0 490 F-z17 540 590 Wavelength (nm ) 0 490 640 540 590 Wavelength (nm ) 640 c) F-z12 F-z17 H2O H2O 800 EtOH 400 EtOH EA EA toluene toluene RFU DMSO DCE 200 DCE 400 DMSO RFU 600 200 0 490 540 590 Wavelength (nm ) 0 490 640 540 590 Wavelength (nm ) 640 Figure 4.9. a) Excitation and emission spectra of ‘hit’ fluorophores and their corresponding alkynes. b) Emission spectra of F-z12 (in DCE, green line) and F-z17 (in EtOH, red line) 96 compared against the alkyne precursor F (in each corresponding solvent; black line). Black arrows indicate the increase in fluorescence after the “click” reaction. c) Emission spectra of F-z12 and F-z17 in various solvents. 4.6 Conclusions In conclusion, we have successfully designed and synthesized 2 new classes of “click” fluorophores based on the xanthone and xanthene scaffolds. The rapid assembly of these fluorophores enabled by “click” chemistry gives easy access to xanthene analogs, which are traditionally difficult to synthesize. We have also identified two fluorophores, F-z12 and F-z17, in which triazole-formation resulted in a significant fluorescence increase. While these fluorophores can potentially be used as green light-emitting substitutes for the existing “click” fluorophores in bioconjugation and bioimaging applications, further optimization of the azide moiety will be necessary to achieve a large fluorescence increase in aqueous solutions. 97 CHAPTER 5 EXPERIMENTAL SECTION 5.1 General Information All chemicals were purchased from commercial vendors and used without further purification. Tetrahydrofuran (THF) was distilled over sodium benzophenone and used immediately. DMF for Sonogashira coupling was dried over CaH2 and distilled under reduced pressure. HPLC grade solvents are used for all other solvents. All reactions requiring anhydrous conditions were carried out under an argon or nitrogen atmosphere using oven-dried glassware. Reaction progress was monitored by TLC on precoated silica plates (Merck 60 F254, 0.25 µm) and spots were visualized by UV, basic KMnO4 or iodine. Flash column chromatography was carried out using Merck 60 F254 0.040-0.063 µm silica gel. 1H and 13C NMR spectra were recorded on Bruker Avance ACF300 spectrometer. Chemical shifts are reported in parts per million relative to internal standard tetramethylsilane (Si(CH3)4 = 0.00 ppm) or residual solvent peaks (CHCl3 = 7.26 ppm, MeOH = 3.31 ppm, DMSO = 2.50 ppm). 13 C- NMR spectra are reported parts per million relative to solvent signal (CHCl3 = 77.0 ppm, MeOH = 49.0 ppm, DMSO = 39.5 ppm). Analytical LC profiles and mass spectra were recorded on a Shimadzu LC-ESI-MS system or a Shimadzu LC-IT-TOFMS system. Reverse-phase Phenomenex Luna 5µm C18(2) 100 Å 50 X 3.0 mm column or Phenomenex Luna 5µm C18(2) 100 Å 150 X 3.0 mm (for peptides). 0.1% formic acid/H2O and 0.1% formic acid/acetonitrile were used as eluents for the LCESI-MS system and 0.1% TFA/H2O and 0.1% TFA/acetonitrile for the LC-IT-TOFMS. The flow rate for both was 0.6 ml/min. 98 5.2 Solution-phase Synthesis of Fluorophores, Linkers and Azides 5.2.1 Synthesis of SG1, SG2 and Related Derivatives O H 2N O OTBS 3-amino-6-(tert-butyldimethylsilyloxy)-9H-xanthen-9-one (2ii) To a solution of 2i (1.14 g, 5 mmol) in dry DMF (50 ml) was added imidazole followed by TBS-Cl (2.26 g, 15 mmol). The reaction was stirred at room temperature for 5 h. The reaction mixture was concentrated in vacuo to half the volume and then poured into H2O (100 ml) and the white precipitate formed was taken into EA (80 ml). The aqueous layer was further washed with EA (80 ml) and the combined organic phase was washed with H2O (2 × 100 ml), brine, dried over Na2SO4 and concentrated in vacuo. The crude product was purified by silica gel chromatography (20-50% EA/hexane) to afford the pure product as a white solid (1.35 g, 79%). 1H-NMR (500 MHz, CDCl3) δ 8.17 (d, J = 8.9 Hz, 1H), 8.11 (d, J = 8.2 Hz), 6.81 (dd, J = 8.2, 1.9 Hz, 1H), 6.80 (apparent s, 1H), 6.63 (dd, J = 8.8, 2.5 Hz, 1H), 6.55 (d, J = 1.9 Hz, 1H), 4.29 (s, 2H), 1.01 (s, 9H), 0.28 (s, 6H). ESI-MS: m/z [M+1]+ calcd: 342.1, found 342.2. O TrtHN O OTBS 3-(tert-butyldimethylsilyloxy)-6-(tritylamino)-9H-xanthen-9-one (2iii) To a solution of 2ii (1.35 g, 3.96 mmol) and pyridine (1.92 ml, 23.7 mmol) in DMF (30 ml) was added CPh3Cl (3.32 g, 11.9 mmol) portionwise at room temperature and stirred for 4 h. The reaction was diluted with H2O (100 ml) and the white precipitate 99 formed was taken into EA (70 ml). The aqueous layer was extracted with EA (70 ml) and the combined organic phase was washed with H2O (2 × 100 ml), brine, dried over Na2SO4 and concentrated in vacuo. The crude product was purified by silica gel chromatography (10-25% EA/hexane) to afford the pure product as a white solid (1.92 g, 83%). 1H-NMR (500 MHz, CDCl3) δ 8.10 (d, J = 8.9 Hz, 1H), 7.93 (d, J = 8.9 Hz, 1H), 7.36 – 7.25 (m, 15H), 6.76 (dd, J = 8.8, 1.9 Hz, 1H), 6.67 (d, J = 2.5 Hz, 1H), 6.53 (dd, J = 8.9, 2.5 Hz, 1H), 0.98 (s, 9H), 0.24 9s, 6H). ESI-MS: m/z [M+1]+ calcd: 584.3, found 584.1. TrtHN O O 9-o-tolyl-6-(tritylamino)-3H-xanthen-3-one (2iv) To a solution of o-tolylmagnesium bromide (2.0 M in Et2O, 2.5 ml, 5.0 mmol) in freshly distilled THF (40 ml) was added dropwise 2iii (0.58 g, 1.0 mmol) dissolved in THF (20 ml) and heated at 60°C under N2 atmosphere for 16 hrs. Another portion of the Grignard reagent (2 eq) was added dropwise and the reaction was stirred for another 16 hrs. The reaction was then quenched with the addition of H2O (THF/H2O, 1:1). The mixture was neutralized to pH 6 with the slow addition of 1N HCl at 0°C, followed by extraction with ether twice, then EtOAc. The combined organic layers were washed with H2O, brine, dried over Na2SO4 and concentrated in vacuo. The crude compound was purified by silica gel chromatography (100% DCM – 10% MeOH/DCM) to give a red solid (0.20 g, 36%). 1H-NMR (300 MHz, DMSO-d6) δ 8.33 (s, 1H), 7.46 – 7.16 (m, 19H), 6.70 (br s, 1H), 6.69 (d, J = 9.5 Hz, 1H), 6.57 (apparent d, J = 9.0 Hz, 1H), 6.28 (dd, J = 9.7, 1.5 Hz, 1H), 6.04 (d, J = 1.5 Hz, 1H). 1.96 (s, 3H). ESI-MS: m/z [M+1]+ calcd: 544.2, found 544.0. 100 AcHN OH 3-acetamidophenol 3-acetamidophenol was synthesized from 3-aminophenol according to a modified published procedure. To a rapidly stirred suspension of ZnO (16.3 g, 0.2 mol), acetic anhydride (56.7 ml, 0.6 mol) and DCM (400 ml) in a 1-litre conical flask was added 3-aminophenol (43.6 g, 0.4 mol) portionwise over 5 min. (Caution: Heat from the reaction causes the DCM to boil, but it is not necessary to cool the reaction mixture). After stirring for 10 min by which time the boiling of the DCM would have subsided, the suspension was filtered and washed rapidly with DCM (100 ml). The solid was collected and washed with EA repeatedly until all the product is extracted into EA. EA was then removed in vacuo and the crude product recrystallized in hot EA to give pinkish white crystals as the pure product. A second crop of crystals may be obtained by recrystallization in EA/hexane. Yield = 80% (48.3 g). 1H-NMR values were in accordance with reported literature values. O O2 N O NH2 3-amino-6-nitro-9H-xanthen-9-one (2-1) Compounds 2-1 and 2-2 were synthesized according to published procedures1 with some modifications. To a solution of 3-acetamidophenol (18.1 g, 120 mmol) in DMF was added K2CO3 (16.6 g, 120 mmol) and with vigorous stirring. 2-chloro-4nitrobenzoic acid (12.1 g, 60 mmol) and Cu powder (0.4 g, 6 mmol) was then added and the reaction was heated to 130°C for 16 hrs. The reaction was cooled to room temperature and poured into 5N HCl (50 ml) and ice (total volume of 500 ml). The pH was adjusted to 1 by the further addition of 5N HCl if necessary. The mixture was 101 stirred at 0°C until a pale brown solid is formed. The solid was then filtered, washed with cold H2O and air-dried. This solid was added portionwise to conc. H2SO4 (100 ml) and heated at 80°C for 1 hr. After cooling to room temperature, the reaction mixture was poured slowly onto ice and stirred until a reddish-brown solid is formed. This solid was filtered and stirred in 20% w/v Na2CO3 solution (200 ml) until the gas evolution ceased. The crude product was then filtered and washed with cold H2O. The product was then suspended in hot acetonitrile (250 ml) and stirred until a shiny yellowish-brown solid was observed. The suspension was then removed from the heat and filtered while hot. The product thus obtained was washed with cold acetonitrile and dried. The combined washings were concentrated in vacuo and the crude product was suspended in acetonitrile or acetone and sonicated. The red solid that settled to the bottom of the flask was removed by decanting. Repeating this process removed most of the side product (red solid). The remaining solid suspended in the solution was then recrystallized in acetone or acetonitrile to give a yellow solid (3.07 g, 20% over 2 steps). Note: the 3-acetamidophenol obtained from Sigma-Aldrich typically gave lower yields and product was tainted with a brown substance that was not easily removed. 1H-NMR (300 MHz, DMSO) δ 8.37 (d, J = 2.1 Hz, 1H), 8.32 (d, J = 8.7 Hz, 1H), 8.15 (dd, J = 8.7, 2.13 Hz, 1H), 7.88 (d, J = 8.7 Hz, 1H), 6.74 (br s, 2H), 6.72 (dd, J = 8.7, 2.0 Hz, 1H), 6.56 (d, J = 2.0 Hz, 1H). ESI-MS: m/z [M-1]- calcd: 255.0, found 255.3. O O2 N O OH 3-hydroxy-6-nitro-9H-xanthen-9-one (2-2) 102 To a solution of 2-1 (3.0 g, 11.7 mmol) in conc. H2SO4 (30 ml) and water (15 ml) at 0°C was added dropwise a solution of NaNO2 (2.43 g, 35.1 mmol) in H2O (10 ml). The reaction was stirred at 0°C for 1hr, then poured into boiling H2O (100 ml). The mixture was stirred at 90°C until an orange-brown solid was formed. The suspension was then cooled to room temperature and filtered. The solid was washed with cold H2O and dried in vacuo to yield the product (2.80 g, 93%) which was pure enough for the next reaction. 1H-NMR (300 MHz, DMSO-d6) δ 8.22 (s, 1H), 8.21 (d, J = 8.6 Hz, 1H), 8.06 (dd, J = 8.7, 1.8 Hz, 1H), 7.91 (d, J = 8.9 Hz, 1H), 6.84, (dd, J = 8.8, 1.89 Hz, 1H), 6.74 (d, J = 1.8 Hz, 1H). 13 C-NMR (75 MHz, DMSO-d6) δ 173.5, 164.7, 157.7, 154.9, 150.4, 128.1, 127.8, 125.0, 118.0, 114.8, 113.8, 113.7, 102.1. ESI-MS: m/z [M-1]- calcd: 256.0, found 256.3. General procedure for the synthesis of 2-3a and 2-3b: To a solution of 2-2 (3.0 g, 11.7 mmol) in DMF at room temperature under N2 atmosphere was added anhydrous K2CO3 (3.23 g, 23.4 mmol) and then dimethylsulfate (for 2-3a, 2.11 ml, 23.4 mmol) or tert-butyldimethylsilyl 5-iodopentyl ether 2-12 (for 2-3b, 4.99 g, 15.2 mmol; see section 5.2.3 for synthesis). The reaction was monitored by TLC to completion. The reaction mixture was then filtered and DMF was removed in vacuo. The solid residue was taken into DCM and washed with NaHCO3, water, brine and dried over Na2SO4. After the removal of the solvent, the crude solid was purified by silica gel chromatography. O O 2N O O 3-methoxy-6-nitro-9H-xanthen-9-one (2-3a) 103 White solid. Yield = 70%. 1H-NMR (300 MHz, CDCl3) δ 8.45 (d, J = 8.7 Hz, 1H), 8.34 (d, J = 2.1 Hz, 1H), 8.25 (d, J = 9.0 Hz, 1H), 8.16 (dd, J = 8.7, 2.1 Hz, 1H), 7.01 (dd, J = 8.9, 2.3 Hz, 1H), 6.93 (d, J = 2.3 Hz, 1H). 13 C-NMR (75 MHz, CDCl3) δ 174.8, 165.9, 158.4, 155.6, 151.0, 128.5, 128.5, 125.8, 118.0, 115.7, 114.3, 113.9, 100.4, 56.0 O O 2N O O OTBS 3-(5-(tert-butyldimethylsilyloxy)pentyloxy)-6-nitro-9H-xanthen-9-one (2-3b) White solid. Yield = 82%. 1H-NMR (300 MHz, CDCl3) δ 8.40 (d, J = 8.7 Hz, 1H), 8.23 (d, J = 2.1 Hz, 1H), 8.16 (d, J = 8.9 Hz, 1H), 8.09 (dd, J = 8.7, 2.1 Hz, 1H), 6.93 (dd, J = 8.7, 2.3 Hz, 1H), 6.83 (d, J = 2.1 Hz, 1H), 4.08 (t, J = 6.4 Hz, 2H), 3.65 (t, J = 5.9 Hz, 2H), 1.91 – 1.83 (m, 2H), 1.66 – 1.53 (m, 4H), 0.89 (s, 9H), 0.05 (s, 6H). 13 C-NMR (75 MHz, CDCl3) δ 174.5, 165.3, 158.2, 155.5, 150.8, 128.4, 128.3, 125.7, 117.9, 115.3, 114.6, 113.8, 100.7, 68.9, 62.8, 32.4, 28.7, 25.9, 22.3, 18.3, -5.3. ESIMS: m/z [M+1]+ calcd: 458.2, found 458.1 O H 2N O O 3-amino-6-methoxy-9H-xanthen-9-one (2-4a) To a suspension of 2-3a (1.2 g, 4.42 mmol) in EtOH (30 ml) was added SnCl2.2H2O (2.89 g, 12.8 mmol), then refluxed overnight. The reaction was cooled to room temperature and EtOH removed in vacuo. Saturated NaHCO3 (30 ml) was then added to the solid, stirred for 5 min and filtered. Acetone was added to dissolve the product. The remaining tin salts were removed by filtration, and washed with acetone. The 104 pure product (1.07 g, 90%) was obtained after removal of acetone in vacuo. 1H-NMR (500 MHz, DMSO) δ 7.99 (d, J = 8.9 Hz, 1H), 7.82 (d, J = 8.9 Hz, 1H), 7.01 (d, J = 1.9 Hz, 1H), 6.94 (dd, J = 8.8, 2.5 Hz, 1H), 6.65 (dd, J = 8.5, 1.9 Hz, 1H), 6.42 (s, 2H), 3.88 (s, 3H). 13C-NMR (125 MHz, DMSO) δ 173.3, 163.9, 158.0, 157.2, 155.3, 127.3, 127.2, 115.2, 112.5, 112.2, 110.7, 100.4, 97.6, 55.9. ESI-MS: m/z [M-1]- calcd: 240.1, found 240.3. O H2N O O OTBS 3-amino-6-(5-(tert-butyldimethylsilyloxy)pentyloxy)-9H-xanthen-9-one (2-4b) To a solution of 2-3b (4.5 g, 9.46 mmol) in EtOAc (60 ml) was added 10% Pd/C (0.45 g) and stirred under hydrogen atmosphere at room temperature overnight. After the completion of the reaction, the mixture was filtered through celite and concentrated to give the pure product (3.72 g, 92%) as a yellow solid. 1H-NMR (300 MHz, CDCl3) δ 8.18 (d, J = 8.8 Hz, 1H), 8.09 (d, J = 8.5 Hz), 6.86 (dd, J = 8.8, 2.3 Hz, 1H), 6.78 (d, J = 2.3 Hz, 1H), 6.62 (dd, J = 8.5, 2.0 Hz, 1H), 6.54 (d, J = 2.0 Hz, 1H), 4.05 (t, J = 6.4 Hz, 2H), 3.70 – 3.63 (m, 2H), 1.87 – 1.80 (m, 2H), 1.62 – 1.53 (m, 4H), 0.90 (s, 9H), -0.01 (s, 6H). 13C-NMR (75 MHz, CDCl3) δ 175.3, 163.9, 158.3, 157.8, 152.4, 128.4, 128.0, 115.8, 113.8, 112.9, 112.3, 100.64, 99.9, 68.5, 62.9, 32.4, 28.8, 26.0, 22.3, 18.3, -5.3. ESI-MS: m/z [M+1]+ calcd: 428.2, found 428.2. General procedure for the synthesis of 2-5a and 2-5b: To a solution of 2-4a or 2-4b in DCM at room temperature was added triethylamine followed by trityl chloride portionwise. The reaction was stirred for 1 hr at room temperature, then H2O was added. The two layers were separated and the aqueous 105 layer extracted with DCM. The combined DCM layers were washed with water, brine and dried over Na2SO4. After the removal of the solvent in vacuo, the crude product was purified by silica gel chromatography (10-20% EA/hexane) as a white solid. O TrtHN O O 3-methoxy-6-(tritylamino)-9H-xanthen-9-one (2-5a) Yield = 88%. 1H-NMR (300 MHz, CDCl3) δ 8.14 (d, J = 8.9 Hz, 1H), 7.93 (d, J = 8.7 Hz, 1H), 7.37-7.26 (m, 15H), 6.83 (apparent d, J = 8.9 Hz, 1H), 6.69 (d, J = 2.3 Hz, 1H), 6.08 (d, J = 2.1 Hz, 1H), 5.67 (s, 1H), 3.85 (s, 3H). 13C-NMR (75 MHz, CDCl3) δ 175.2, 164.2, 157.8, 157.5, 151.8, 144.3, 129.1, 128.2, 127.9, 127.3, 126.9, 116.0, 113.4, 112.4, 101.3, 100.1, 71.8, 55.6. ESI-MS: m/z [2M+Na]+ calcd: 989.4, found 989.0. O TrtHN O O OTBS 3-(5-(tert-butyldimethylsilyloxy)pentyloxy)-6-(tritylamino)-9H-xanthen-9-one (2-5b) Yield = 97%. 1H-NMR (300 MHz, CDCl3) δ 8.12 (d, J = 8.9 Hz, 1H), 7.93 (d, J = 8.7 Hz, 1H), 7.37 – 7.25 (m, 15H), 6.82 (dd, J = 8.9, 2.3 Hz, 1H), 6.67 (d, J = 2.2 Hz, 1H), 6.53 (apparent d, J = 8.8, 1H), 6.08 (d, J = 2.3 Hz, 1H), 4.00 (t, J = 6.5 Hz, 2H), 3.63 (t, J = 6.1 Hz, 2H), 1.87 – 1.77 (m, 2H), 1.59 – 1.50 (m, 4H), 0.89 (s, 9H), 0.05 (s, 6H). 13C-NMR (75 MHz, CDCl3) δ 175.2, 163.7, 157.8, 157.5, 144.3, 129.0, 128.2, 127.8, 127.3, 126.9, 115.7, 113.8, 113.0, 112.8, 101.3, 100.5, 71.8, 68.4, 62.9, 32.4, 28.8, 25.9, 22.3, 18.3, -5.3. ESI-MS: m/z [2M+Na]+ calcd: 1361.7, found 1361.3. 106 General procedure for the synthesis of SG1 and SG2: To a solution of o-tolylmagnesium bromide (2.0 M in Et2O, 5 eq) in freshly distilled THF (0.5 M) was added dropwise 2-5a or 2-5b dissolved in THF (0.1 M) and heated at 60°C under N2 atmosphere for 16 hrs. Another portion of the Grignard reagent (2 eq) was added dropwise and the reaction was stirred for another 16 hrs. The reaction was then quenched with the addition of H2O (THF/H2O, 1:1). The mixture was neutralized to pH 6 with the slow addition of 1N HCl at 0°C, followed by extraction with ether twice, then EtOAc. The combined organic layers were washed with H2O, brine, dried over Na2SO4 and concentrated in vacuo. The crude compound was purified by silica gel chromatography (100% DCM – 10% MeOH/DCM). This product was then dissolved in DCM and H2O followed by addition of trifluoroacetic acid (DCM:TFA:H2O, 7:2:1) at room temperature. The reaction was stirred for 30 min, then H2O was added to the reaction and the two layers were separated. The DCM layer was washed with sat. NaHCO3, water, brine and dried over Na2SO4. After the removal of the solvent, the crude product was purified on a short silica gel column. An analytically pure sample was obtained by recrystallization in CHCl3/hexane. HN O O 6-methoxy-9-o-tolyl-3H-xanthen-3-imine (SG1) Bright red solid. Yield = 68% from 2-5a. 1H-NMR (500 MHz, MeOD) δ 7.89 (s, 1H), 7.59 (t, J = 7.55 Hz, 1H), 7.53 (d, J = 7.55 Hz, 1H), 7.48 (t, J = 7.55 Hz, 1H), 7.46 (d, J = 2.5 Hz, 1H), 7.35 (d, J = 8.80 Hz, 1H), 7.32 (d, J = 9.45 Hz, 1H), 7.29 (d, J = 7.55 Hz, 1H), 7.17 (dd, J = 8.85, 2.5, 1.9 Hz, 1H), 7.06 (dd, J = 9.45, 1.85 Hz, 1H), 6.98 (d, J = 2.55 Hz, 1H), 4.08 (s, 3H), 2.06 (s, 3H). 13C-NMR (125 MHz, MeOD) δ 107 169.7, 164.3, 161.7, 161.5, 158.5, 137.2, 134.4, 132.8, 132.3, 132.0, 131.6, 130.1, 127.4, 121.3, 118.3, 118.3 116.7, 101.6, 98.6, 57.5, 19.7. IT-TOF-MS: m/z [M+1]+ calcd for for C21H18NO2+: 316.1332, found 316.1053. HN OH O O 5-(6-imino-9-o-tolyl-6H-xanthen-3-yloxy)pentan-1-ol (SG2) Red solid. Yield = 56% from 2-5b. 1H-NMR (500 MHz, MeOD) δ 7.89 (s, 1H), 7.59 (dt, J = 7.55, 1.25 Hz, 1H), 7.53 (d, J = 7.55 Hz, 1H), 7.48 (t, J = 7.58 Hz, 1H), 7.43 (d, J = 2.5 Hz, 1H), 7.33 (d, J = 9.45 Hz, 1H), 7.31 (d, J = 9.45 Hz, 1H), 7.28 (d, J = 7.55 Hz, 1H), 7.15 (dd, J = 8.85, 2.50 Hz, 1H), 7.05 (dd, J = 9.45, 1.90 Hz, 1H), 6.97 (d, J = 1.90 Hz, 1H), 4.29 (t, J = 6.60 Hz, 2H), 3.60 (t, J = 6.30 Hz, 2H), 2.05 (s, 3H), 1.94 – 1.89 (m, 2H), 1.65 – 1.57 (m, 4H). 13 C-NMR (125 MHz, MeOD) δ 169.1, 164.2, 161.7, 161.5, 158.5, 137.2, 134.3, 132.8, 132.3, 132.0, 131.6, 130.1, 127.4, 121.3, 118.6, 118.2, 116.6, 102.0, 98.6, 71.1, 62.7, 33.2, 29.7, 23.4, 19.7. IT-TOF-MS: m/z [M+1]+ calcd for C25H26NO3+ : 388.1907, found 388.1323. O OH N FmocHN O O O Fmoc-Asp-SG1 (2-7) To a solution of Fmoc-Asp(OtBu)-OH (323 mg, 0.78 mmol), HBTU (297 mg, 0.784 mmol) and HOBt (106 mg, 0.784 mmol) in DMF (10 ml) was added DIEA (0.315 ml, 1.81 mmol) and stirred for 5 min at room temperature under nitrogen atmosphere. 26a (190 mg, 0.603 mmol) was then added and the reaction stirred for 1 hr. The 108 reaction was quenched by pouring the mixture into water. After extracting 3 times with EA, the combined EA layers were washed with water, brine, dried over Na2SO4 and concentrated. The crude product was purified by silica gel chromatography to give a yellowish white solid. This solid was then dissolved in DCM and trifluoroacetic acid was added. The reaction was stirred for 5 hrs at room temperature, after which the solvent and TFA was removed in vacuo. The residue was taken into DCM and washed twice with water, brine and dried over Na2SO4. Evaporation of the solvent gave 2-7 (0.267 g, 68% over 2 steps) which was sufficiently pure (as shown by HPLC analysis) to be used for peptide synthesis. IT-TOF-MS: m/z [M+1]+ calcd for C40H32N2O7+: 653.2282, found: 653.1644. FmocN O O OH Fmoc-SG2-ol (2-8) To a mixture of 2-6b (1.00 g, 2.58 mmol) and NaHCO3 (0.87 g, 10.3 mmol) in 1:1 THF/H2O (40 ml) at 0°C was added Fmoc-Cl. The temperature was gradually raised to room temperature and stirred for 4 hrs. H2O was added and the aqueous layer extracted with ether (3X). The combined ether layers were washed with brine, dried over Na2SO4, filtered and concentrated. The pure product was obtained as an orangered foam solid (1.02 g, 65%) after column purification (100% DCM – 5% MeOH/DCM). IT-TOF-MS: m/z [M+1]+ calcd: 610.252, found: 610.194. 109 FmocN O O O Fmoc-SG2-CHO (2-9) To a solution of 2-8 (1.00 g, 1.64 mmol) in DCM (15 ml) was added Dess-Martin periodinane (1.25 g, 2.95 mmol) at 0°C. After 15 min, the ice bath was removed and stirring was continued at room temperature for 2 hrs. Saturated NaHCO3 was added to the reaction and stirred until the two layers became clear. The DCM layer was then separated, washed with H2O, brine, dried over Na2SO4, filtered and concentrated. The product was purified on a short silica gel column to afford an orange-red foam solid (0.92 g, 92%). IT-TOF-MS: m/z [M+1]+ calcd for C40H34NO5+: 608.2431, found: 608.1861. FmocN O OH O O Fmoc-SG2-COOH Fmoc-SG2-ol (2-8) (1.83 g, 3 mmol) was dissolved in DMF (25 ml) and cooled to 0οC under a nitrogen atmosphere. PDC (4.51 g, 12 mmol) was added portionwise over 5 min to the rapidly stirred solution. The temperature was slowly raised to room temperature and stirred for 16 h. The reaction mixture was poured into an ice/water mixture (100 ml) and stirred for 15 min. The orange precipitate formed was filtered and washed with cold water (5 ×). The resulting solid was dried briefly in vacuo, and then lyophilized to remove the remaining traces of water (1.59 g, 85%). This solid was used directly for subsequent reactions. 110 FmocN O O O Cl Fmoc-SG2-COCl Fmoc-SG2-COOH (0.94 g, 1.5 mmol) was dissolved in freshly distilled DCM (20 ml) under a nitrogen atmosphere and cooled to 0οC. A catalytic amount of DMF was added and then oxalyl chloride (0.26 ml, 3 mmol) was added dropwise over 3 min. The reaction was then slowly raised to RT and stirred for another 3 h. The solvent and oxalyl chloride was removed in vacuo to afford a yellow solid (0.89 g, 92%) which was used without further purification. 5.2.2 Synthesis of Alkynes A – F General procedure for the conversion of phenols to triflates To a suspension of the phenol (1 mmol) in dry DCM (15 ml) was added dry pyridine (0.40 ml, 5 mmol) and cooled to 0°C. Trifluoromethanesulfonic anhydride (0.25 ml, 1.5 mmol) was then added dropwise at 0°C. The reaction was gradually raised to room temperature and stirred for another 2 hrs. H2O (10 ml) was then added to quench the reaction. After stirring for 5 min, the two phases were separated and the aqueous layer extracted with DCM (15 ml). The combined organic phase was washed with 1N HCl (2 × 10 ml), water (10 ml), dried over Na2SO4 and concentrated in vacuo. The pure product was isolated by silica gel chromatography using EtOAc/hexane as the eluent. General procedure for Sonogashira coupling 111 Dry DMF was degassed by bubbling argon gas for 30-45 min. The triflate (1 mmol), PdCl2(PPh3)2 (70.2 mg, 0.1 mmol) and CuI (38.1 mg, 0.2 mmol) were dissolved in degassed DMF (15 ml) under an argon atmosphere Triethylamine (1.39 ml, 10 mmol) was then added followed by ethynyltrimethylsilane (0.28 ml, 2 mmol). The reaction was stirred for 2 hrs and poured into water (40 ml) and extracted with ethyl acetate (3 × 20 ml). The combined organic phase was washed with brine, dried over Na2SO4 and concentrated. The crude product was purified by silica gel chromatography using EtOAc/hexane as the eluent. General procedure for the deprotection of the trimethylsilyl (TMS) group for alkynes A-C The TMS-protected alkynes (1 mmol) were dissolved in MeOH/THF 1:2 (30 ml) and 1 N NaOH (10 ml) was added at room temperature. The reaction was stirred for 30 min. Ether was added to the reaction mixture and stirred for 15 min, then the two phases were separated. The aqueous phase was extracted with diethyl ether or EtOAc (2 × 20 ml). The combined organic phase was washed with brine, dried over Na2SO4 and concentrated in vacuo. The pure product was obtained by purification over a short silica gel column using EtOAc/hexane as the eluent. O O O OH 3-hydroxy-6-methoxy-9H-xanthen-9-one1 (4-1i) To a solution of 3,6-dihydroxy-xanthen-9-one (1.14 g, 5 mmol) in DMF (80 ml) was added anhydrous powdered K2CO3 (0.90 g, 6.5 mmol) and stirred for 2 hrs at room temperature until the K2CO3 dissolved completely. Dimethylsulfate (0.48 ml, 5 mmol) 112 in DMF (10 ml) was then added dropwise at room temperature over 20 min. The reaction was stirred for an additional 2 hrs. The solid formed was filtered off and then the DMF was removed in vacuo. H2O (30 ml) was added to the residue and extracted with EtOAc (4 × 50 ml). The combined organic phase was washed with water (50 ml), brine, dried over Na2SO4 and concentrated. The product was isolated by silica gel chromatography (30-50% EtOAc/hexane) to give 4-1i as an off-white solid (0.52 g, 43%). The remaining starting material was recovered in approximately 30% yield. 1HNMR (300 MHz, DMSO) δ 8.04 (d, J = 8.9 Hz, 1H), 8.00 (d, J = 8.7 Hz, 1H), 7.09 (d, J = 2.3 Hz, 1H), 7.00 (dd, J = 8.9, 2.5 Hz, 1H), 6.89 (dd, J = 8.7, 2.3 Hz), 6.84 (d, J = 2.3 Hz, 1H), 3.91 (s, 3H). ESI-MS: m/z [M+1]+ calcd: 243.1, found 243.0. O O O OTf 6-methoxy-9-oxo-9H-xanthen-3-yl trifluoromethanesulfonate (4-2a) 4-2a was obtained from 4-1i as a white solid following the general procedure for the conversion of phenols to triflates. Yield = 84%. 1H-NMR (300 MHz, CDCl3) δ 8.42 (d, J = 8.9 Hz, 1H), 8.24 (d, J = 8.9 Hz, 1H), 7.42 (d, J = 2.3 Hz, 1H), 7.27 (dd, J = 8.9, 2.3 Hz, 1H), 6.98 (dd, J = 8.9, 2.3 Hz, 1H), 6.89 (d, J = 2.5 Hz, 1H), 3.95 (s, 3H). 13 C-NMR (75 MHz, CDCl3) δ 174.8, 165.5, 158.1, 156.5, 152.6, 129.3, 128.4, 121.8, 117.1, 115.6, 113.9, 111.0, 100.4, 55.9. ESI-MS: m/z [M+1]+ calcd: 375.0, found 374.9. O O O TMS 3-methoxy-6-(2-(trimethylsilyl)ethynyl)-9H-xanthen-9-one (4-3a) 113 4-3a was obtained from 4-2a as a white solid following the general procedure for Sonogashira coupling. Yield = 60%. 1H-NMR (300 MHz, CDCl3) δ 8.24 (d, J = 8.1 Hz, 1H), 8.23 (d, J = 8.9 Hz, 1H), 7.53 (d, J = 1.3 Hz, 1H), 7.41 (dd, J = 8.2, 1.5 Hz, 1H), 6.94 (dd, J = 8.9, 2.3 Hz, 1H), 6.87 (d, J = 2.3 Hz, 1H), 3.94 (s, 3H), 0.29 (s, 9H). 13 C-NMR (75 MHz, CDCl3) δ 175.6, 165.2, 158.1, 155.8, 129.1, 128.3, 127.3, 126.6, 121.6, 121.6, 120.9, 115.9, 113.4, 103.4, 100.3, 98.9, 55.8, -0.22. ESI-MS: m/z [M+1]+ calcd: 323.1, found 323.0. O O O Alkyne A Alkyne A was obtained from 4-3a as a white solid following the general procedure for the deprotection of TMS-protected alkynes A-C. Yield = 90%. 1H-NMR (300 MHz, CDCl3) δ 8.26 (d, J = 8.9 Hz, 1H), 8.24 (d, J = 9.7 Hz, 1H), 7.57 (d, J = 1.3 Hz, 1H), 7.44 (d, J = 8.2, 1.3 Hz, 1H), 6.95 (dd, J = 8.9, 2.5 Hz, 1H), 6.88 (d, J = 2.3 Hz, 1H), 3.94 (s, 3H), 3.31 (s, 1H). 13 C-NMR (75 MHz, CDCl3) δ 175.6, 165.3, 158.1, 155.7, 128.3, 128.0, 127.4, 126.7, 121.3, 115.8, 113.6, 100.2, 82.3, 80.9, 55.9. ESIMS: m/z [M+1]+ calcd: 251.1, found 251.0. O O Alkyne D A solution of 4-3a (0.129 g, 0.4 mmol) in dry THF (10 ml) was added to otolylmagnesium bromide (2.0 M in Et2O, 1.0 ml, 2 mmol) in dry THF (10 ml) under an argon atmosphere at room temperature. The reaction was then heated to 50°C and 114 stirred for 5 hrs, and quenched by the dropwise addition of H2O until gas evolution ceased. The reaction was then neutralized with 2 N HCl and diluted with the addition of diethyl ether (20 ml) and stirred for 5 min. The 2 phases were separated and the aqueous phase extracted with Et2O (2 ×10 ml). The combined organic phase was washed with H2O (10 ml), brine, dried over Na2SO4 and concentrated to give the intermediate tertiary alcohol which was used immediately without further purification. The alcohol was dissolved in dry DCM (20 ml) and cooled to -78° in a dry iceacetone bath. BBr3 (0.15 ml, 1.6 mmol) was added dropwise to the solution. The reaction was gradually raised to room temperature and stirred for 16 hrs. The reaction was poured slowly into ice-water (~ 10 ml) with vigorous stirring and stirred for another 15 min. The 2 phases were separated and the aqueous phase was extracted with DCM (2 × 10 ml). The combined organic phase was washed with brine, dried over Na2SO4 and concentrated. The pure product D was obtained by silica gel chromatography (20-50% EtOAc/hexane) as a bright orange solid (27.3 mg, 22% over 2 steps). 1H-NMR (300 MHz, DMSO) δ 7.69 (d, J = 1.5 Hz, 1H), 7.53 – 7.36 (m, 5H), 6.95 (d, J = 8.2 Hz, 1H), 6.89 (d, J = 9.7 Hz, 1H), 6.59 (dd, J = 9.8, 2.0 Hz, 1H), 6.26 (d, J = 2.0 Hz, 1H), 4.60 (s, 1H), 2.03 (s, 3H). 13 C-NMR (75 MHz, DMSO-d6) δ 184.4, 157.8, 151.5, 147.1, 135.7, 131.6, 130.8, 130.5, 129.6, 129.1, 128.0, 127.9, 126.4, 126.2, 120.6, 120.5, 119.5, 105.1, 85.1, 82.3. (Note: 1 C was not resolved). HRMS calcd for [C22H15O2]+ : 311.1067, found 311.1056. O TfO O OTf 3,6-Di-OTf-xanthone (4-1ii) Compound 4-1ii was synthesized following a literature procedure3. 115 1 H-NMR (300 MHz, CDCl3) 8.45 (d, J = 8.9 Hz, 2H), 7.49 (d, J = 2.3 Hz, 2H), 7.36 (dd, J = 8.9, 2.3 Hz, 2H). O Et2N O OTf 6-(diethylamino)-9-oxo-9H-xanthen-3-yl trifluoromethanesulfonate (4-2b) 4-2b was synthesized from 4-1ii following a reported procedure3 with modifications. Diethylamine (1.06 ml, 20 mmol) was added to a solution of 4-1ii (0.985 g, 2 mmol) in DMSO (15 ml) at room temperature. The reaction was stirred at 80°C for 4 hrs, followed by removal of diethylamine in vacuo. The solution was then poured into icewater (15 ml) and the solid residue was taken up into diethyl ether. The aqueous layer was extracted with diethyl ether (2 × 15 ml). The combined organic phase was washed twice with H2O, brine, dried over Na2SO4 and concentrated. The pure product was isolated by silica gel chromatography (20-30% EtOAc/hexane) as a yellow solid (0.357 g, 43%). 1H-NMR (300 MHz, CDCl3) δ 8.37 (d, J = 8.9 Hz, 1H), 8.10 (d, J = 9.2 Hz, 1H), 7.33 (d, J = 2.3 Hz, 1H), 7.22 (dd, J = 8.9, 2.3 Hz, 1H), 6.72 (dd, J = 9.0, 2.5 Hz, 1H), 6.47 (d, J = 2.3 Hz), 3.48 (q, J = 7.1 Hz, 4H), 1.29 (t, J = 7.1 Hz, 6 H). 13 C-NMR (75 MHz, CDCl3) δ 173.9, 158.8, 156.4, 153.2, 152.1, 129.0, 128.3, 122.2, 116.4, 110.9, 110.6, 110.0, 96.1, 44.9, 12.4. 19 F-NMR (282 MHz, CDCl3) δ 3.3 (s) ESI-MS: m/z [M+1]+ calcd: 416.1, found 416.0. O Et2N O TMS 3-(diethylamino)-6-(2-(trimethylsilyl)ethynyl)-9H-xanthen-9-one (4-3b) 116 4-3b was obtained from 4-3a as a yellow solid following the general procedure for Sonogashira coupling. Yield = 81%. 1H-NMR (300 MHz, CDCl3) δ 8.21 (d, J = 8.0 Hz, 1H), 8.11 (d, J = 9.2 Hz, 1H), 7.46 (d, J = 1.5 Hz, 1H), 7.37 (dd, J = 8.1, 1.5 Hz, 1H), 6.69 (d, J = 9.0, 2.5 Hz, 1H), 6.45 (d, J = 2.5 Hz, 1H), 3.47 (q, J = 7.1 Hz, 4H), 1.25 (t, J = 7.1 Hz, 6H), 0.28 (s, 9H). 13 C-NMR (75 MHz, CDCl3) δ 174.9, 158.7, 155.6, 152.9, 128.2, 128.1, 126.8, 126.4, 122.1, 120.6, 111.3, 109.6, 103.8, 98.0, 96.2, 44.8, 12.5, -0.2. ESI-MS: m/z [M+1]+ calcd: 364.2, found 364.1. O Et2N O Alkyne B Alkyne B was obtained from 4-3b as a yellow solid following the general procedure for deprotection of TMS-protected alkynes A-C. Yield = 92%. 1H-NMR (300 MHz, CDCl3) δ 8.23 (d, J = 8.0 Hz, 1H), 8.12 (d, J = 9.2 Hz, 1H), 7.50 (d, J = 1.1 Hz, 1H), 7.40 (dd, J = 8.0, 1.5 Hz, 1H), 6.70 (dd, J = 9.2, 2.5 Hz, 1H), 6.47 (d, J = 2.5 Hz, 1H), 3.48 (q, J = 7.1 Hz, 4H), 3.27 (s, 1H), 1.26 (t, J = 7.1 Hz, 6H). 13C-NMR (75 MHz, CDCl3) δ 174.8, 158.7, 155.6, 153.0, 128.2, 127.1, 126.9, 126.6, 122.5, 121.0, 111.3, 109.7, 96.2, 82.6, 80.2, 44.9, 12.5. HRMS calcd for [C19H18O2N1]+ : 292.1332, found 292.1330. Et2N Cl- TMS O N-ethyl-N-(9-o-tolyl-6-(2-(trimethylsilyl)ethynyl)-3H-xanthen-3ylidene)ethanaminium chloride (4-3ii) 117 A solution of 4-3b (0.145 g, 0.4 mmol) in dry THF (10 ml) was added to otolylmagnesium bromide (2.0 M in Et2O, 1.0 ml, 2 mmol) in dry THF (10 ml) under an argon atmosphere at room temperature. The reaction was then heated to 50°C and stirred for 12 hrs, and quenched by the dropwise addition of H2O until gas evolution ceased. The reaction was then acidified with 2 N HCl to pH = 1 and diluted with the addition of DCM (20 ml) and stirred for 5 min. The 2 phases were then separated and the aqueous layer saturated with brine (20 ml) and extracted with DCM (4 × 20 ml). The combined organic phase was dried over Na2SO4, filtered and concentrated. This crude product was used for the next step without further purification. Et2N Cl- O Alkyne E Compound 4-3ii was stirred in a solution of 1:1 1N NaOH/MeOH (20 ml) at room temperature for 1 hr. The reaction was acidified to pH = 1 and MeOH was removed in vacuo. The solution was then saturated with brine (10 ml) and extracted with DCM (4 × 20 ml). The combined organic phase was dried over Na2SO4, filtered and concentrated. The pure product E was isolated as a dark purple solid (0.112 g, 70% over 2 steps) by silica gel chromatography (100% DCM – 10% MeOH/DCM). 1HNMR (300 MHz, DMSO) δ 7.99 (d, J = 1.3 Hz, 1H), 7.64 – 7.48 (m, 5H), 7.35 – 7.21 (m, 4H), 4.90 (s, 1H), 3.93 – 3.81 (m, 4H), 2.03 (s, 3H), 1.33 – 1.23 (m, 6H). 13 C- NMR (75 MHz, DMSO) δ 159.0, 158.5, 156.4, 153.1, 135.8, 132.5, 130.8, 130.7, 130.3, 129.3-129.1 (5 C), 126.2, 121.1, 120.1, 119.8, 96.8, 87.7, 82.1, 46.9, 46.8, 19.2, 13.3, 12.3. HRMS calcd for [C26H24O1N1]+ : 366.1852, found 366.1862. 118 O O2 N O OTf 6-nitro-9-oxo-9H-xanthen-3-yl trifluoromethanesulfonate (4-5) 4-5 was obtained from 3-hydroxy-6-nitro-9H-xanthen-9-one4 4-4 as an off-white solid following the general procedure for the conversion of phenols to the corresponding triflates. Yield = 87%. 1H-NMR (300 MHz, CDCl3) δ 8.53 (d, J = 8.7 Hz, 1H), 8.47 (d, J = 8.9 Hz, 1H), 8.41 (d, J = 2.1 Hz, 1H), 8.24 (dd, J = 8.7, 2.0 Hz, 1H), 7.55 (d, J = 2.3 Hz, 1H), 7.38 (dd, J = 8.7, 2.3 Hz, 1H). 13C-NMR (75 MHz, CDCl3) δ 174.6, 156.8, 155.7, 153.6, 151.6, 129.7, 128.9, 125.3, 121.4, 120.8, 118.9, 118.4, 116.6, 114.3, 111.6. 19F-NMR (282 MHz, CDCl3) δ 3.5 (s). ESI-MS: not found. O O2N O SiMe3 3-nitro-6-(2-(trimethylsilyl)ethynyl)-9H-xanthen-9-one (4-6) 4-6 was obtained from 4-5 as a white solid following the general procedure for Sonogashira coupling. Yield = 65%. 1H-NMR (300 MHz, CDCl3) δ 8.49 (d, J = 8.7 Hz, 1H), 8.37 (d, J = 2.0 Hz, 1H), 8.26 (d, J = 8.2 Hz, 1H), 8.18 (dd, J = 8.7, 2.1 Hz, 1H), 7.63 (d, J = 1.3 Hz, 1H), 7.49 (d, J = 8.2, 1.3 Hz, 1H), 0.30 (s, 9H). 13C-NMR (75 MHz, CDCl3) δ 175.3, 156.0, 155.7, 151.4, 131.0, 128.7, 128.4, 126.8, 125.6, 121.3, 118.3, 114.31, 102.8, 100.8, 77.2, -0.29. ESI-MS: not found. O H2N O TMS 3-amino-6-(2-(trimethylsilyl)ethynyl)-9H-xanthen-9-one (4-7) 119 To a solution of 4-6 (0.169 g, 0.5 mmol) in 3:1 MeOH/THF (20 ml) was added saturated NH4Cl solution (2 ml) followed by Zn powder (0.327 g, 5 mmol) and stirred for 30 min. The solids were then removed by filtration and washed with EtOAc (30 ml). The solution was concentrated in vacuo to approximately half the volume and H2O (20 ml) was added. The 2 phases were separated and the aqueous phase was extracted with EtOAc (2 × 10 ml). The combined organic phase was washed with H2O, brine, dried over Na2SO4, filtered and concentrated. This crude product was then used without purification for the next step. O H2N O Alkyne C Alkyne C was obtained as a yellow solid (95 mg, 81% over 2 steps) from the intermediate aniline 4-7 following the general procedure for the deprotection of the TMS group for alkynes A-C. 1H-NMR (300 MHz, DMSO) δ 8.07 (d, J = 8.2 Hz, 1H), 7.84 (d, J = 8.7 Hz, 1H), 7.65 (d, J = 1.3 Hz, 1H), 7.44 (dd, J = 8.1, 1.3 Hz, 1H), 6.66 (dd, J = 8.7 Hz, 1H), 6.59 (s, 2H), 6.51 (d, J = 2.0 Hz, 1H), 4.55 (s, 1H). 13C-NMR (75 MHz, DMSO) δ 173.1, 158.0, 156.0, 155.0, 127.6, 126.9, 126.8, 126.1, 121.6, 120.7, 112.7, 110.7, 97.4, 84.3, 82.3. O TrtHN O SiMe3 3-(2-(trimethylsilyl)ethynyl)-6-(tritylamino)-9H-xanthen-9-one (4-8) To a solution of 4-7 (0.154 g, 0.5 mmol) in DCM (20 ml) was added triethylamine (0.208 ml, 1.5 mmol) and trityl chloride (0.209 g, 0.75 mmol) at room temperature. 120 The reaction was stirred for 3 hrs. H2O (10 ml) was added to the reaction and stirred for 10 min, followed by phase separation. The aqueous layer was extracted once with DCM (10 ml). The combined organic phase was washed with H2O (10 ml), brine, dried over Na2SO4 and concentrated. The pure product 4-8 was isolated by silica gel chromatography (10-20% EtOAc/hexane) as a white solid (0.20 g, 73% over 2 steps) 1 H-NMR (300 MHz, CDCl3) δ 8.13 (d, J = 8.0 Hz, 1H), 7.92 (d, J = 8.9 Hz, 1H), 7.36 – 7.15 (m, 18H), 6.54 (dd, J = 8.7, 2.1 Hz), 6.09 (d, J = 2.1 Hz, 1H), 5.78 (s, 1H), 0.25 (s, 9H). 13C-NMR (75 MHz, CDCl3) δ 175.2, 157.5, 155.5, 152.4, 144.2, 129.0, 128.3, 128.3, 127.4, 127.0, 126.7, 126.3, 121.8, 120.8, 114.2, 113.1, 103.7, 101.2, 98.2, 71.8, -0.23. ESI-MS: m/z [M+1]+ calcd: 550.2, found 550.4. TMS HN O 9-o-tolyl-6-(2-(trimethylsilyl)ethynyl)-3H-xanthen-3-imine (4-9) A solution of 4-8 (0.192 g, 0.35 mmol) in dry THF (10 ml) was added to otolylmagnesium bromide (2.0 M in Et2O, 0.875 ml, 1.75 mmol) in dry THF (10 ml) under an argon atmosphere at room temperature. The reaction was then heated to 50°C and stirred for 20 hrs, and quenched by the dropwise addition of H2O until gas evolution ceased. The solution was then acidified by 2N HCl to pH = 2. DCM was added and stirred for 5 min. The 2 phases were separated, the aqueous phase saturated with brine (10 ml) and extracted with DCM (4 × 10 ml). The combined organic phase was dried over Na2SO4, filtered and concentrated. This crude product was redissolved in DCM (7 ml). H2O (1 ml) and TFA (2 ml) were then added at room temperature. The reaction was stirred for 1 hr and quenched by the addition of H2O (5 ml). The 2 121 phases were separated and the aqueous phase saturated with brine (5 ml) and extracted with DCM (4 × 10 ml). The combined organic phase was washed once with NaHCO3 (10 ml), dried over Na2SO4, filtered and concentrated. The crude product was purified by a short silica gel column to yield 4-9 as an orange-red solid (73 mg, 55% over 2 steps). 1H-NMR (300 MHz, DMSO) δ 9.84 (br s, 1H), 9.72 (br s, 1H), 8.01 (d, J = 1.3 Hz, 1H), 7.63 – 7.49 (m, 4H), 7.32 (apparent d, 1H), 7.28 (d, J = 9.5 Hz, 1H), 7.22 (d, J = 8.4 Hz, 1H), 7.20 (dd, J = 9.5, 1.8 Hz, 1H), 6.97 (d, J = 1.8 Hz, 1H), 2.01 (s, 3H), 0.28 (s, 9H). 13 C-NMR (75 MHz, DMSO) δ 163.6, 159.4, 156.6, 152.9, 135.7, 133.6, 130.8, 130.7, 130.3, 129.4, 129.1, 126.2, 122.5, 120.8, 120.2, 120.0, 103.2, 101.9, 97.6, 19.2, -0.47. ESI-MS: m/z [M+1]+ calcd: 382.2, found 381.2. HN O Alkyne F 4-9 (73 mg, 0.19 mmol) was stirred in 1:1 1N NaOH/MeOH (8 ml) for 1 hr, then the pH of the solution was adjusted to pH = 2. DCM was added (20 ml) and the 2 phases were separated. The aqueous layer was saturated with brine (5 ml), and extracted with DCM (4 × 10 ml). The combined organic phase was dried over Na2SO4 and concentrated in vacuo. The crude product was purified by a short silica gel column to yield F as an orange-red solid (45 mg, 76%). 1H-NMR (300 MHz, MeOD) δ 7.96 (d, J = 1.3 Hz, 1H), 7.63 – 7.47 (m, 4H), 7.39 (d, J = 2.8 Hz, 1H), 7.35 (d, J = 1.8 Hz, 1H), 7.30 (apparent d, J = 7.4 Hz, 1H), 7.14 (dd, J = 9.4, 2.1 Hz, 1H), 6.99 (d, J = 2.0 Hz, 1H), 4.17 (s, 1H), 2.07 (s, 1H). 13 C-NMR (75 MHz, MeOD) δ 165.7, 161.9, 160.1, 154.9, 137.5, 135.0, 132.3, 132.2, 132.1, 131.8, 130.7, 130.6, 130.3, 127.5, 123.2, 122 122.3, 121.9, 121.6, 98.9, 86.3, 82.7, 19.7. HRMS calcd for [C22H16O1N1]+ : 310.1226, found 310.1239. 5.2.3 Synthesis of Linkers pTscl, NEt3, DCM TBS-Cl, imidazole, DMF HO OH overnight, 53% HO OTBS 5 hrs, 78% 2-10 NaI, acetone TsO OTBS 2-11 overnight, 90% I OTBS 2-12 Scheme 5.1. Synthesis of linker 2-12 used in the preparation of SG2 5-(tert-butyldimethylsilyloxy)pentan-1-ol2 (2-10) To a well-stirred solution of imidazole (6.80 g, 100.0 mmol) and 1,5-pentanediol (26.2 ml, 250 mmol) in dry DMF (200 ml) under nitrogen atmosphere was added a slurry of TBDMS-Cl (7.54 g, 50.0 mmol) in DMF (30 ml) dropwise over 30 min. The resulting mixture was stirred at room temperature overnight. DMF was removed in vacuo and H2O was added to the colorless oil residue and extracted twice with ether. The combined ether layers were washed with water, brine and dried over Na2SO4. After the removal of the solvent in vacuo, the crude product was purified by silica gel column chromatography (100% hexane – 20% EA/hexane) to give a slightly yellow oil (5.78 g, 53%) as the desired product. 1H-NMR (300 MHz, CDCl3) δ 3.65 – 3.59 (m, 4H), 1.63 – 1.50 (m, 5H), 1.43 – 1.36 (m, 2H), 0.88 (s, 9H), 0.039 (s, 6H); ESI-MS: m/z [M+1]+ calcd: 219.2, found 219.0. 5-(tert-butyldimethylsilyloxy)pentan-1-ol (2-11) To a solution of 2-10 (5.00 g, 22.9 mmol) and triethylamine (9.51 ml, 68.8 mmol) in DCM (100 ml) at 0°C was added p-toluenesulfonyl chloride (5.68 g, 29.8 mmol) in 123 small portions. The reaction was gradually raised to room temperature and stirred overnight. H2O was added and stirred for 5 min. The two layers were separated. The DCM layer was washed twice with saturated NaHCO3, water, brine and dried over Na2SO4. After the removal of the solvent in vacuo, the crude product was purified by silica gel chromatography to give a pale yellow oil (5.58 g, 78%). 1 H-NMR (300 MHz, CDCl3) δ 7.78 (d, J = 8.22 Hz, 2H), 7.33 (d, J = 8.22 Hz, 2H), 4.02 (t, J = 6.59 Hz, 2H), 3.55 (t, J = 6.17 Hz, 2H), 2.44 (s, 3H), 1.70 – 1.61 (m, 2H), 1.48 – 1.32 (m, 4H), 0.93 (s, 9H), 0.089 (s, 6H); ESI-MS: m/z [M+1]+ calcd: 373.2, found 373.0. tert-butyldimethylsilyl 5-iodopentyl ether3 (2-12) 2-11 (5.5 g, 16.0 mmol) was added to a suspension of NaI (12.0 g, 80.1 mmol) in dry acetone (200 ml) at room temperature. The reaction mixture was stirred overnight. The white solid formed was filtered, and the filtrate concentrated. The oil residue was taken into ether and washed with saturated Na2SO3, H2O and brine, and dried over Na2SO4. The pure product (4.72 g, 90%) was obtained after concentration in vacuo. 1 H-NMR (300 MHz, CDCl3) δ 3.61 (t, J = 6.17 Hz, 2H), 3.19 (J = 6.99 Hz, 2H), 1.89 – 1.80 (m, 2H), 1.57 – 1.41 (m, 4H), 0.89 (s, 9H), 0.05 (s, 6H). 5.2.4 Synthesis of Azides Aromatic azides Aromatic azides except z4, z8 and z12 were previously prepared and reported by our group.5 Azides z4, z8 and z12 were prepared from the corresponding anilines following the protocol from the same reference. 124 N3 N O 4-(4-azidophenyl)morpholine (z4) 1 H-NMR (300 MHz, CDCl3) δ 6.97 – 6.88 (m, 4H), 3.87 – 3.84 (m, 2H), 3.13 – 3.10 (m, 2H). 13C-NMR (75 MHz, CDCl3) δ 148.8, 131.7, 129.2, 119.8, 117.1, 115.7, 66.8, 49.6, 49.4. N3 O2N 1-azido-4-nitrobenzene (z8) 1 H-NMR (300 MHz, CDCl3) δ 8.23 (dt, J = 9.2, 3.0, 2.2 Hz, 2H), 7.13 (dt, J = 9.1, 2.9, 2.2 Hz, 2H). 13C-NMR (75 MHz, CDCl3) δ146.8, 144.6, 125.5, 119.3. N3 Br 4-azido-2-bromo-1-methylbenzene (z9) 1 H-NMR (300 MHz, CDCl3) δ 7.21 – 7.18 (m, 2H), 8.87 (dd, J = 8.2, 2.3 Hz, 1H), 2.36 (s, 3H). 13 C-NMR (75 MHz, CDCl3) δ138.8, 134.4, 131.4, 125.4, 122.7, 117.9, 22.2. N N3 3-azidoquinoline (z12) 125 1 H-NMR (300 MHz, CDCl3) δ 8.61 (d, J = 2.6 Hz, 1H), 8.08 (d, J = 8.6 Hz, 1H), 7.77 – 7.73 (m, 2H), 7.69 – 7.63 (m, 1H), 7.56 (dt, J = 8.4, 1.2 Hz, 1H). 13C-NMR (75 MHz, CDCl3) δ 145.8, 143.8, 129.4, 128.8, 128.1, 127.7, 126.8, 122.6. N3 N 8-azidoquinoline (z13) 1 H-NMR (300 MHz, CDCl3) δ 8.90 (dd, J = 4.3, 1.7 Hz, 1H), 8.14 (dd, J = 8.4, 1.6 Hz, 1H), 7.57 (dd, J = 8.2, 1.3 Hz, 1H), 7.50 (d, J = 7.4 Hz, 1H), 7.44 (dd, J = 8.4, 13 C-NMR (75 MHz, CDCl3) δ 149.4, 4.3 Hz, 1H), 7.36 (dd, J = 7.4, 1.3 Hz, 1H). 137.2, 136.1, 129.3, 126.5, 124.1, 121.9, 118.2. Aliphatic azides Aliphatic azides z14, z16 and z17 were previously prepared and reported by our group.6 Synthesis of azide z15 is as below: O O pTsCl, KOH, OH Et2O O O 80% NaN3, DMF OTs ο 70 C 95% O O z15 N3 Scheme 5.2. Synthesis of azide z15. O O OTs (2,3-dihydrobenzo[b][1,4]dioxin-2-yl)methyl 4-methylbenzenesulfonate To a solution of 2-hydroxymethyl-1,4-benzodioxane (0.166g, 1 mmol) in DCM (10 ml) was added triethylamine (0.42 ml, 3 mmol) and cooled to 0°C. p-Toluenesulfonyl chloride (0.286 g, 1.5 mmol) was then added portionwise. The reaction was gradually 126 raised to room temperature and stirred overnight. Water (10 ml) was then added and stirred for 5 min. The two phases were separated and the aqueous phase extracted with DCM (10 ml). The combined organic phase was washed with saturated NaHCO3 solution, brine, dried over Na2SO4 and concentrated. The crude product was purified by silica gel chromatography to yield the product as a white solid (0.256 g, 80%). 1HNMR (300 MHz, CDCl3) δ 7.80 (d, J = 8.3 Hz, 2H), 7.35 (d, J = 8.4 Hz, 2H), 6.87 – 6.76 (m, 4H), 4.42 – 4.35 (m, 1.08), 4.28 – 4.16 (m, 3H), 4.03 (dd, J = 11.6, 6.3 Hz, 1H), 1.64 (s, 3H). 13 C-NMR (75 MHz, CDCl3) δ 145.2, 142.7, 142.2, 132.3, 130.0, 128.0, 121.8, 121.7, 117.3, 117.2, 70.2, 67.1, 64.3, 21.6. O O N3 Azide z15 To a solution of the tosylate (0.256 g, 0.8 mmol) in DMF (10 ml) was added NaN3 (0.156 g, 2.4 mmol). The suspension was stirred at 65°C for 5 hrs and filtered. DMF was removed in vacuo and the residue was taken into ether and washed with water (2 × 5 ml). The organic phase was washed with brine, dried over Na2SO4, filtered and concentrated to give a colorless oil as the pure product z15 (0.145 g, 95%). 1H-NMR (300 MHz, CDCl3) δ 6.94 - 6.84 (m, 4H), 4.38 – 4.31 (m, 1H), 4.25 (dd, J = 11.3, 2.4 Hz, 1H), 4.06 (dd, J = 11.3, 6.7 Hz, 1H), 3.58 (dd, J = 13.1, 6.0 Hz, 1H), 3.49 (dd, J = 13.1, 5.2 Hz, 1H). 13C-NMR (75 MHz, CDCl3) δ 142.8, 142.4, 121.9, 121.9, 117.4, 117.2, 71.9, 65.2, 50.6. 127 5.3 Solid-Phase Synthesis of Peptides and SG-Peptide Conjugates 5.3.1 General Information All peptide synthesis described herein are carried out using standard Fmoc chemistry. HBTU, HOBt, and Fmoc-protected amino acids were purchased from GL Biochem (Shanghai, China). Fmoc-protecting amino acids with side chain protecting groups are listed here: Fmoc-Asp(OtBu)-OH, Fmoc-Glu(OtBu)-OH, Fmoc-His(Trt)-OH, FmocLys(Boc)-OH, Fmoc-Asn(Trt)-OH, Fmoc-Gln(Trt)-OH, Fmoc-Arg(Pbf)-OH, FmocThr(tBu)-OH, Fmoc-Tyr(tBu)-OH. MicroKansTM or MacroKansTM were used to contain the resin for the synthesis of individual peptide sequences. 5.3.2 General Procedures General procedure for Fmoc deprotection The Fmoc-protected amino-functionalized resin or peptide chain was treated with 20% piperidine/DMF for 45 min at room temperature. The resin was washed with DMF (3×), DCM (3×) and DMF (3×). General procedure for coupling of Fmoc-amino acids onto resin Fmoc-amino acid (4 equiv), HBTU (4 equiv) and HOBt (4 equiv) were dissolved in DMF (0.05 – 0.1 M) and DIEA (8 equiv) was added and agitated for 5 min. This preactivated Fmoc-amino acid solution was added to the resin and shaken for 3 h at room temperature. The resin was filtered and washed with DMF (3×), DCM (3×) and DMF (3×). 128 General procedure for the N-terminal capping of resin-bound peptides Following Fmoc deprotection of the last amino acid in the peptide sequence, the resin was washed in DMF (4×) and DCM (4×). The resin was resuspended in DCM, and DIEA (20 equiv) and acetic anhydride (10 equiv) were added sequentially. The resin was shaken for 3 h, then filtered and washed with DCM (4×) and dried thoroughly in vacuo. 5.3.3 Synthesis of Ac-DEVD-SG1 Loading of compound 2-7 onto 2-chlorotritylchloride resin Compound 2-7 (33 mg, 0.05 mmol) was dissolved in dry DCM (1.5 ml) and DIEA (35 µL, 0.2 mmol) was added. This solution was then added to 2-chlorotrityl chloride resin (50 mg, 0.025 mmol, loading ~0.5 mmol/g) previously swollen in dry DCM and shaken for 2 hrs at room temperature. The resin was filtered and washed thoroughly with DCM (5×) until the filtrate became colorless. The resin was then end-capped with MeOH for 1 hr. After filtration the resin was washed with DMF (3×), DCM (3×) and DMF (3×). Peptide synthesis Deprotection of resin-bound Fmoc-Asp-SG1 was carried out using the general procedure for Fmoc deprotection. The peptide chain was elongated using the general procedure for coupling amino acids onto the resin. Ac-Asp(OtBu)-OH was used instead of Fmoc-Asp(OtBu)-OH for the terminal residue. 129 Cleavage of peptide from resin After coupling Ac-Asp(OtBu)-OH onto the solid support, the resin was washed with DMF (3×), DCM (3×), MeOH (3×) and dried thoroughly under vacuum. A solution of TFA/TIS (95:5, 2 ml) was added to the resin at room temperature and shaken for 2.5 hrs. The resin was filtered off and washed with DCM (2×). The combined DCM and cleavage solutions were concentrated to ~0.3 ml, then cold diethyl ether (3 ml) was added to precipitate the peptide. The peptide was then collected by centrifugation and washed with cold diethyl ether. This washing process was repeated for another time. The peptide thus obtained was dried in vacuo. 5.3.4 Synthesis of SG2-Peptide Conjugates Synthesis of threonyl-glycyl resin (TG-resin) Fmoc-Gly-OH was loaded onto aminomethyl polystyrene resin (500 mg, loading 0.30.8 mmol) previously swollen in DMF as described in the general procedure for coupling amino acids onto solid support. After the coupling reaction, the resin was filtered and washed with DMF (3×), DCM (3×) and DMF (3×). Fmoc deprotection was carried out also as described above. The resin was washed with DMF (3×), DCM (3×) and DMF (3×). The procedure was repeated with Fmoc-Thr(OtBu)-OH to give the O-protected TG-resin. Deprotection of the tert-butyl group was carried out with TFA/TIS (95:5) for 1 hr. The resin was subsequently washed with DCM (5×), then 10% DIEA/DCM for 15 min, then with DMF (3×), THF (3×) and DCM (3×) and dried in vacuo to give the final TG-resin. Loading Fmoc-SG2-CHO onto TG-resin 130 Fmoc-SG2-CHO (3 eq) was dissolved in MeOH/DCM/DMF/AcOH (6:2:1:1, 3 ml) and added to the TG-resin (350 mg) and shaken for 5 hrs at room temperature. The resin was filtered and washed briefly with DCM (2×), then with DMF (3×), THF (3×), DCM (3×). Boc protection of secondary amine in oxazolidine moiety Boc2O (5 eq) was dissolved in DCM and added to the loaded resin and DIEA (2.5 eq) was added. The resin was shaken for 3 hrs, filtered and washed with DMF (3×), THF (3×), DCM (3×). The resin was then dried and swollen in DMF for 30 min before prior to coupling the first Fmoc amino acid. Peptide synthesis Fmoc amino acids were coupled sequentially onto the resin-bound SG2 using the general procedures for coupling and Fmoc deprotection. 30 mg of resin was used for each peptide. Side-chain deprotection and cleavage of peptides from resin The side chains and Boc-protected oxazolidines were deprotected by shaking the resin in TFA/TIS (95:5) for 45 min. For peptides containing the Arg(Pbf) residues, the deprotection cocktail used was TFA/TIS/H2O (95:4:1) and deprotection was carried out for 2 hrs. The resin was filtered and washed with DCM (5×). The peptides were released from the solid support by adding a mixture of DCM/MeOH/AcOH/H2O (12:5:2:1) to the resin and shaking for 15 min. This release procedure was repeated for another 2 times. The combined cleavage solutions containing individual peptides were concentrated and dried completely in vacuo for short and hydrophobic peptides 131 (sequences: FG, EY, AAF and AAL). For the rest of the peptides, the solutions were concentrated to ~ 300 µL and precipitated with 3 ml of cold ether. The peptides were then collected by centrifugation and dried in vacuo. 5.3.5 Synthesis of Alkyne-Functionalized SG2-Based Substrates Reductive amination of PL-FMP resin with propargyl amine 500 mg of the resin (0.45 mmol, loading = 0.9 mmol/g) was swollen in DMF for 1 hr. The resin was then filtered and resuspended in DMF/MeOH/AcOH (80:19:1, 8 ml). Propargyl amine was added (0.31 ml, 4.5 mmol), followed by NaBH3CN (301 mg, 4.5 mmol). The suspension was agitated gently by magnetic stirring and heated at 55°C overnight. The resin was then filtered and washed with DMF (3×), MeOH (3×) and DCM (3×). Acylation with Fmoc-SG2-COCl The resin (500 mg, 0.45 mmol) was swollen in dry DCM, then filtered and resuspended in freshly distilled DCM. DIEA (0.78 ml, 4.5 mmol) was then added and the suspension was cooled to 0°C. Fmoc-SG2-COCl in dry DCM was then added dropwise with gentle stirring. The reaction was raised to RT and stirred overnight. The resin was filtered and washed with DCM (5×) followed by DMF (5×). The resin appears deep red in color. Peptide synthesis After Fmoc deprotection, peptide synthesis and N-terminal capping were carried out using the general procedures described in 5.3.2. 132 Cleavage Prior to the final cleavage step, the resin was washed with 1% TFA/DCM for 0.5 hrs, then washed again with DCM (3×). The resin was then dried thoroughly in vacuo. Deprotection of the side chain protecting groups and cleavage from solid support was carried out with TFA/TIS/H2O (95:2.5:2.5). The solution was then concentrated to ~ 300 µL and precipitated with 3 ml of cold ether. The peptides were then collected by centrifugation and dried in vacuo. 5.3.6 Synthesis of Azido-Peptides and Control Peptides Loading Fmoc-AA-OH onto Rink Amide resin Rink amide resin (0.65 mmol, 1.3 g, loading = 0.5 mmol/g) was swollen in DMF (10 ml) for 30 min. The resin was filtered and the Fmoc group was removed by shaking the resin 20% piperidine/DMF for 30 min, followed by washing with DMF (3×) and repeating the deprotection procedure for another 30 min. The resin was then filtered and washed with DMF (3×), DCM (3×) and DMF (3×). Peptide synthesis Sequential coupling of Fmoc-protected AAs onto the resin and Fmoc deprotection was carried out using the general procedures described in 5.3.2. When FmocArg(Pbf)-OH is used as the AA, the coupling reaction was allowed to proceed for 14h to ensure complete coupling. Double coupling was not necessary for the nonaarginine peptide. 133 N-terminal capping with 4-azidobutanoic acid After Fmoc deprotection of the final AA in the peptide (40 mg of resin), a preactivated solution of 4-azidobutanoic acid (4 equiv), HBTU (4 equiv), HOBt (4 equiv) and DIEA (8 equiv) in DMF (1 ml) was added to the resin and shaken for 3 h. The resin was filtered and washed with DMF (4×), DCM (4×) and dried in vacuo. N-terminal capping with Fmoc-SG2-COCl After Fmoc deprotection of the final AA in the peptide (40 mg of resin), the resin was washed with DMF (3×) and DCM (3×). The resin was then swollen in dry DCM for 30 min, filtered and re-suspended in dry DCM and cooled to 0°C. DIEA (10 equiv) was then added. A solution of Fmoc-SG2-COCl in dry DCM was added dropwise to the resin and shaken at RT for 3 h. The resin was filtered and washed with DCM (5×), DMF (3×) and DCM (3×). Prior to the final cleavage step, the resin was washed with 1% TFA/DCM for 0.5 hrs, then washed again with DCM (3×). The resin was then dried thoroughly in vacuo. Cleavage Deprotection of the side chain protecting groups and cleavage from solid support was carried out with TFA/TIS/H2O (95:2.5:2.5). The solution was then concentrated to ~ 300 µL and precipitated with 3 ml of cold ether. The peptides were then collected by centrifugation and dried in vacuo. 134 5.4 Synthesis of Fluorophores Using “Click” Chemistry 5.4.1 Microplate-Based Assembly of “Click” Fluorophore Library 20 mM stock solutions of the alkynes and azides were prepared in DMSO. 40 mM CuSO4 (50 mg in 5 ml) solution and 100 mM sodium ascorbate (99 mg in 5 ml) solution were prepared in H2O. tBuOH/H2O was first added to a 384-deep well microplate, followed by the alkyne (4 µL) and azide (10 µL). CuSO4 and sodium ascorbate solutions were mixed separately prior to the reaction and the mixture (8 µL) was added immediately to the solution of alkyne and azide in the microplate. The plate was then sealed with a silicone cap-mat and shaken at room temperature. The volumes of each building block and reagent are summarized in the table below. The final concentrations of alkynes in the reaction are 2 mM for alkynes A-C and 1.33 mM for alkynes D-F. Stock concentration (mM) Vol. used (µ µL) Molar ratio Alkyne 20 4 1 Azide 20 10 2.5 CuSO4 40 4 2 Sodium ascorbate 100 4 5 3.75:1 tBuOH/H2O (for alkynes A-C) - 38 - 1:1 tBuOH/H2O (for alkynes D-F) - 18 - Table 5.1. Reagent concentrations and volumes used per ‘click’ reaction 135 For alkynes A-C: After shaking for 12 hrs, the solvent was removed in vacuo in a GeneVac HT-4X Series II parallel evaporation system. The ‘click’ product was re-suspended in 200 µL of DMSO (concentration ~ 400 µM) and used directly for LC-MS analysis. For alkynes D-F: After shaking for 12 hrs, 20 µL of each reaction solution was diluted to 100 µL in DMSO (concentration ~ 400 µM) and used directly for LC-MS analysis. 5.4.2 Scale-up Synthesis of “Hit” Fluorophores Reaction set-up The alkyne (0.03 mmol) and azide (0.075 mmol) were dissolved in DMSO/tBuOH/H2O in a 15 ml centrifuge tube. CuSO4·5H2O (15 mg, 0.06 mmol) and sodium ascorbate (30 mg, 0.15 mmol) were dissolved in H2O (0.5 ml) and then added to the solution of the alkyne and azide. The tube was then shaken for 3 days for alkynes A-C, and 16 hrs for alkynes D-F. Reaction progress was monitored by LCMS. Due to difference in solubility of the alkynes, the amount and proportion of solvent for different alkynes varies. The table below summarizes the volumes of each solvent used. The final concentrations for the reaction are 8.6 mM for alkynes A-D and 10 mM for alkynes E-F. Vol. of tBuOH (µ µL) Total vol. of H2O (µ µL) Vol. of DMSO (µ µL) Alkynes A-D 1500 1000 1000 Alkynes E, F 1500 1500 0 Table 5.2. Volumes of solvents used for scale-up ‘click’ chemistry 136 Work-up and purification i) “Click” products from alkynes A – C: product was precipitated by the addition of H2O (10 ml) and collected by centrifugation. The solid residue was dissolved in chloroform or DCM (10 ml), washed with H2O (2 × 5 ml) and dried over Na2SO4. The crude product was purified by silica gel chromatography (A-z9 and B-z15) or preparative HPLC (C-z17). A-z6 was purified by washing the crude product with acetonitrile twice. ii) “Click” products from alkynes D – F: DCM (10 ml) was added to the reaction mixture and washed with H2O (5 ml). The aqueous layer was saturated with brine and extracted with DCM (10 ml). The combined organic phase was dried over Na2SO4, filtered and concentrated. The crude product was purified by silica gel chromatography (D-z2) or preparative HPLC (E-z2, F-z12 and F-z17). O O O A-z6 F N N N F 1 H-NMR (300 MHz, CDCl3) δ 8.41 (d, J = 8.2 Hz, 1H), 8.41 (d, J = 2.6 Hz, 1H), 8.27 (d, J = 8.9 Hz, 1H), 8.11 (d, J = 1.3 Hz, 1H), 8.07 – 8.00 (m, 1H), 7.84 (dd, J = 8.2, 1.5 Hz, 1H), 7.16 – 7.11 (m, 2H), 6.97 (dd, J = 8.9, 2.3 Hz, 1H), 6.93 (d, J = 2.3 Hz, 1H), 3.96 (s, 3H). ESI-MS: m/z [M+1]+ calcd: 406.1, found 406.0. O O O A-z9 N N N Br 137 1 H-NMR (300 MHz, CDCl3) δ 8.40 (d, J = 8.2 Hz, 1H), 8.31 (s, 1H), 8.27 (d, J = 8.9 Hz, 1H), 8.11 (d, J = 1.3 Hz, 1H), 8.02 (d, J = 2.1 Hz, 1H), 7.82 (dd, J = 8.4, 1.5 Hz, 1H), 7.68 (dd, J = 8.2, 2.5 Hz, 1H), 7.43 (d, J = 8.0 Hz, 1H), 6.95 (dd, J = 8.7, 2.3 Hz, 1H), 6.93 (d, J = 2.3 Hz, 1H), 3.96 (s, 3H), 2.50 (s, 3H). ESI-MS: m/z [M+1]+ calcd: 462.0, found 463.9. O N O N N B-z15 1 N O O H-NMR (300 MHz, CDCl3) δ 8.34 (d, J = 8.2 Hz, 1H), 8.13 (d, J = 9.1 Hz, 1H), 8.01 (s, 1H), 7.95 (d, J = 1.3 Hz, 1H), 7.71 (dd, J = 8.2, 1.5 Hz, 1H), 6.95 – 6.88 (m, 4H), 6.70 (dd, J = 9.0, 2.5 Hz, 1H), 6.50 (d, J = 2.5 Hz, 1H), 4.81 – 4.70 (m, 2H), 4.69 – 4.66 (m, 1H), 4.38 (dd, J = 11.7, 2.1 Hz, 1H), 3.96 (dd, J = 11.7, 5.8 Hz, 1H), 3.48 (q, J = 7.1 Hz, 4H), 1.26 (t, J = 7.1 Hz, 1H). ESI-MS: m/z [M+1]+ calcd: 483.2, found 483.1. O H2N O C-z17 1 N N N NH S O O COOH H-NMR (300 MHz, CDCl3) δ 8.74 (s, 1H), 8.29 (br s, 1H), 8.15 (d, J = 8.2 Hz, 1H), 8.15 - 8.10 (m, 2H), 7.98 (apparent d, J = 7.6 Hz, 1H), 7.94 (apparent s, 1H), 7.87 – 7.81 (m, 2H), 7.72 – 7.67 (m, 1H), 4.49 (t, J = 5.6 Hz, 1H). Note: triplet CH2 peak at ~ 3.33 obscured by H2O peak. ESI-MS: m/z [M+1]+ calcd: 506.1, found 506.0. 138 O N N N O D-z2 1 H-NMR (300 MHz, CDCl3) δ 8.33 (s, 1H), 7.98 (d, J = 1.5 Hz, 1H), 7.78 (dd, J = 8.4, 1.7 Hz, 1H), 7.59 (apparent s, 1H), 7.50 – 7.38 (m, 4H), 7.29 (d, J = 8.2 Hz, 1H), 7.21 (d, J = 7.6 Hz, 1H). 7.12 (d, J = 8.4 Hz, 1H), 6.98 (d, J = 9.7 Hz, 1H). 6.58 (dd, J = 9.7, 2.0 Hz, 1H), 6.48 (d, J = 1.8 Hz, 1H), 2.37 (s, 3H), 2.34 (s, 3H), 2.11 (s, 3H). HRMS calcd for [C30H24O2N3]+ : 458.1863, found 458.1871. N Cl- O N N N E-z2 1 H-NMR (300 MHz, CDCl3) δ 9.63 (s, 1H), 8.36 (s, 1H), 8.12 (dd, J = 8.5, 1.6 Hz, 1H), 7.77 (d, J = 2.1 Hz, 1H), 7.69 – 7.51 (m, 5H), 7.47 (d, J = 2.3 Hz, 1H), 7.43 (d, J= 8.6 Hz, 1H), 7.43 – 7.38 (m, 2H), 7.31 (d, J = 2.1 Hz, 1H), 7.24 (d, J = 9.9 Hz, 1H), 3.88 (q, J = 6.7 Hz, 2H), 3.82 (q, J = 7.2 Hz, 2H), 2.36 (s, 3H), 2.32 (s, 3H), 1.33 (t, J = 7.0 Hz, 3H), 1.26 (t, J = 7.2 Hz, 3H). HRMS calcd for [C34H33O1N4]+ : 513.2649, found 513.2645. HN O N N N N F-z12 1 H-NMR (300 MHz, CDCl3) δ 9.90 (s, 1H), 9.57 (br s, 1H), 9.51 (br s, 1H), 9.01 (d, J = 2.3 Hz, 1H), 8.46 (s, 1H), 8.21 – 8.14 (m, 3H), 7.93 – 7.87 (m, 1H), 7.78 (apparent t, J = 7.71 Hz, 1H), 7.64 – 7.51 (m, 4 H), 7.45 (d, J = 8.6 Hz, 1H), 7.38 (d, J = 7.4 Hz, 139 1H), 7.30 (d, J = 9.5 Hz, 1H), 7.19 (dd, J = 9.4, 1.7 Hz, 1H), 7.00 (d, J = 1.8 Hz, 1H), 2.08 (s, 3H). HRMS calcd for [C32H22O1N5]+ : 480.1819, found 480.1835. N N HN O N O NH S O COOH F-z17 1 H-NMR (300 MHz, CDCl3) δ 9.47 (br s, 1H), 9.41 (br s, 1H), 8.88 (s, 1H), 8.33 (s, 1H), 8.26 (s, 1H), 8.18 – 7.94 (m, 3H), 7.70 – 7.49 (m, 3H), 7.36 (d, 8.4 Hz, 1H), 7.28 (d, J = 2.3 Hz, 1H), 7.18 (apparent d, J = 9.5 Hz, 1H), 7.11 (s, 1H), 6.99 (br s, 1H), 6.94 (s, 1H), 4.52 (t, 5.6 Hz, 1H), 3.5 (m, 2H), 2.07 (m, 3H). HRMS calcd for [C31H26O5N5S1]+ : 580.1649, found 580.1662. 5.5 Spectroscopic Analysis 5.5.1 General Information Fluorescence measurements were recorded on a Perkin Elmer LS55 fluorescence spectrometer. Absorbance data were recorded on a Shimadzu UV-2450 spectrometer. Samples were prepared as 10 mM stock solutions in DMSO and diluted in the appropriate solvents for analysis. 5.5.2 Determination of Molar Extinction Coefficients and Quantum Yields UV absorption and integrated fluorescence emission for SG1 was determined in EtOH with sample concentrations ranging from 0.4 to 1.6 µM, with excitation 140 wavelength at 493 nm. Fluorescein (Φst = 0.95) in 0.1 N NaOH was used as standard, with excitation at 496 nm. UV absorption and integrated fluorescence emission was determined in DMSO for A, Az-6, Az-9, B, B-z15, C and C-z17, and in EtOH for D, D-z2, E, E-z2, F, F-z12 and F-z17. Rhodamine 6G (Φst = 0.95, excitation wavelength = 488 nm) in H2O and Coumarin 1 (Φst = 0.73, excitation wavelength = 360 nm) were used as standards for alkynes D-F and alkynes A-C respectively. Relative fluorescence quantum yield was determined using the following equation: Φx = Φst(mx/mst)(ηx2/ηst2) Where Φx, Φst = quantum yield of SG1 and fluorescein respectively mx, mst is the slope of best linear fit from the plot of integrated fluorescence intensity against absorbance (at 493 nm for SG1, 496 nm for fluorescein) η = refractive index of solvent used (η = 1.333 for H2O, η = 1.334 for 0.1 N NaOH, η = 1.361 for EtOH at 25°C, η = 1.479 for DMSO at 20°C) 5.6 Microplate-Based Fluorescence Assays 5.6.1 General Information Serine proteases trypsin (Cat# 93610, Fluka), α-chymotrypsin (Cat# C-4129, Sigma), β-chymotrypsin (Cat# C-4629, Sigma) and subtilisin (Cat# 85968, Fluka) and thrombin Cat# (T-3399, Sigma) were purchased from commercial sources. Cysteine proteases caspase-3 and caspase-7 were recombinantly expressed as previously describedi. 141 5.6.2 Enzymatic Assays with SG-Peptide Conjugates Cleavage of the peptide substrates by selected enzymes were monitored in black flatbottom polypropylene 384-well plates (Nunc, USA) using 25 µL volume for each reaction. 20 µΜ of substrate and 0.125 U of each enzyme were used for each assay. Enzymatic cleavage of the substrates was monitored by fluorescence increase (excitation and emission wavelengths at 485 nm and 520 nm respectively) with a SynergyTM 2 multi-mode microplate reader (Biotek Instruments). 5.6.3 Fluorescence Analysis of “Click” Fluorophore Library The 400 µM DMSO solution of the crude product from the click chemistry reaction used for LC-MS analysis was further diluted to 20 µM in 4 different solvents (40% DMSO/H2O, DMSO, EtOH and DCE) for preliminary examination of fluorescence properties in microplate format. Fluorescence spectra of each reaction product were obtained in black flat-bottom polypropylene 384-well plates (Nunc, USA) using a volume of 40 µL for each click product with a SynergyTM 2 multi-mode microplate reader (Biotek Instruments). The solvent used for each fluorophore is the solvent in which the fluorescence intensity was the highest. 142 5.7 Microarray Experiments Preparation of slides Amine-functionalized glass slides were prepared as previously described6. The Nphthalimide-protected alkoxyamine linker 15 (85 mM) was pre-activated with HBTU (128 mM) and DIEA (170 mM) in DMF for 5 min and the slides were incubated with this solution for 2-3 hrs. The slides were then rinsed with DMF, then ethanol and dried under a stream of nitrogen. Deprotection of the phthalimide group was then carried out by incubating the slides with 3% hydrazine/DMF for 3 hrs. Slides were then rinsed with ethanol and air-dried. These alkoxyamine-functionalized slides may be stored at room temperature until the time of usage. Microarray preparation and reactions Peptides were diluted to a final concentration of 0.25 mM in 16 µL 1:1 DMSO/spotting buffer (250 mM NaOAc, 300 mM NaCl, pH 5) and distributed in a 384-well plate. The peptides were spotted onto the alkoxyamine-functionalized slides with an ESI SMA arrayer (Ontario, Canada) with the printhead installed with Stealth SMP15B Microspotting pins (Telechem, U.S.A.). Spots generated were of approximately 350 µm diameter and were printed with a spot to spot spacing of 750 µm. The pins were rinsed in between samples using two cycles of wash (for 10 s) and sonication (for 10 s) in reservoirs containing 70% ethanol followed by drying under reduced pressure (for 10 s). The slides were allowed to stand for 3 h on the printer platform, and rinsed with ethanol to remove the excess peptides followed by airdrying. 143 Stock solutions of the proteases were prepared in appropriate buffers according to the table below. Enzymatic reactions were initiated by applying the enzyme solution onto the slides. To halt the reaction, slides were dipped in water for 2 min, rinsed with ethanol and air-dried. Protease Concentration Buffer Papain 0.01U/µL Tris.HCl, pH 8 Chymopapain 0.01U/µL PBS, pH 7.4 α-Chymotrypsin 0.01U/µL PBS, pH 7.4 β-Chymotrypsin 0.01U/µL PBS, pH 7.4 Trypsin 0.01U/µL PBS, pH 7.4 Thrombin 0.01U/µL PBS, pH 7.4 Subtilisin 0.01U/µL PBS, pH 7.4 Caspase-3 20 nM Caspase-3/7 assay buffer Caspase-7 20 nM Caspase-3/7 assay buffer Table 5.3. Concentrations and buffers for proteases used in microarray experiments. Caspase-3/7 assay buffer consists of 20 mM PIPES, 100 mM NaCl, 10 mM DTT, 1 mM EDTA, 0.1% w/v CHAPS, 25% w/v sucrose, pH 7.2. Spotting pattern for fingerprint and kinetic experiments A) B) P2 P4 P6 P8 P10 P2 P4 P6 P8 P10 P2 P4 P6 P8 P10 P1 P3 P5 P7 P9 P1 P3 P5 P7 P9 P1 P3 P5 P7 P9 P3 P4 P5 P6 P3 P4 P5 P6 Figure S1. Spotting pattern used in each sub-grid in A) fingerprint experiments and B) kinetic analysis on the microarray, as shown in Figure 3 in the maintext. 144 Kinetic analysis of peptide cleavage from microarray and microplate experiments Time-dependent experiments on the microarray was carried out with various enzymes and peptides using an enzyme concentration of 0.01 U/µL (total volume = 20 µL for each sub-grid in Figure S1). Fluorescence data was taken at various time points and quantified. To obtain kinetic data, the data was fitted to the following equation: ∆RF = RFinit × [1-exp(-kobs × time)] Where kobs = (kcat/Km)*E, representing the observed kcat/Km under a given enzyme concentration E, and ∆RF is the change in fluorescence intensity (RF – RFinit). For microplate-based kinetic experiments, 0.125 U of the enzyme was incubated with 20 µM of peptide and the fluorescence intensity was taken at various time points. The kinetic curves from the microarray- and microplate-based experiments are shown below. 5.8 Bioimaging 5.8.1 General Information Images were captured using an Olympus IX71 inverted microscope, equipped with a 60X oil objective (NA 1.4, WD 0.13 mm) and CoolSNAP HQ CCD camera (Roper Scientific, Tucson, AZ, USA). Images were processed with MetaMorph software (version 7.1.2.; Molecular Devices, PA, USA). The filter sets used for the different fluorophores were as follows: SG1/SG2: Ex 460– 480HQ, dichroic DM485, 145 Em 495–540HQ; TMR-dextran: Ex BP535–555HQ, dichroic DM565, Em 570– 625HQ. 5.8.2 Apoptosis imaging in live cells with Ac-DEVD-SG1 The human carcinoma epithelial carcinoma cell line HeLa was cultured in growth media (DMEM) supplemented with 10% fetal bovine serum, penicillin and streptomycin). Cells were maintained in a humidified atmosphere of 5% CO2 at 37 °C. Cells were seeded on glass-bottom dishes (Mattek, USA) and grown to 70% confluence. Before microinjection, the growth media was replaced with phenol-red free media (Invitrogen). TMR-dextran (Invitrogen, D-1819) was co-injected with 50 µM of Ac-DEVD-SG1 into the cells as a marker. All injections were performed using Eppendorf InjectMan® NI 2 (injection pressure = 25 hPa, compensation pressure = 13 hPa, injection time = 0.2s). To induce apoptosis, cells were incubated with 1.0 µM of staurosporine for 1 hr and imaged. 5.8.3 Evaluating the subcellular locations of the localization peptides† The human carcinoma epithelial carcinoma cell line HeLa and the human breast adenocarcinoma cell line MCF7 were cultured in growth media (DMEM) supplemented with 10% fetal bovine serum, penicillin and streptomycin). Cells were maintained in a humidified atmosphere of 5% CO2 at 37 °C. Cells were seeded on glass-bottom 24-well plates (Mattek, USA) and grown to 80% confluence. Prior to incubation with the peptides, the growth media was removed and † Work done in collaboration with Candy Lu H. S. 146 replaced with serum-free phenol red-free media. The peptides were added to individual wells to a final concentration of ~10 µM (1% DMSO). The cells were then incubated with the peptides for 1 h and rinsed briefly with PBS, and resuspended in serum-free media. The corresponding organelle trackers were added to each well at a concentration of 2 µ/ml and incubated for 15 min before rinsing and reconstitution in serum-free phenol red-free media. 147 CHAPTER 7 REFERENCES 1. Strebhardt, K.; Ullrich, A. Nat. Rev. Cancer 2008, 8, 473-480. 2. Drews, J. Science 2000, 287, 1960-1964. 3. Giepmans, B. N. G.; Adams, S. R.; Ellisman, M. H.; Tsien, R. Y. Science 2006, 312, 217-224. 4. Lakowicz, J. R. Principles of fluorescence spectroscopy; 2nd ed.; Springer US, 2006. 5. 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Letter Three Letter Amino Acid A Ala Alanine C Cys Cysteine D Asp Aspartic acid E Glu Glutamic acid F Phe Phenylalanine G Gly Glycine H His Histidine I Ile Isoleucine K Lys Lysine L Leu Leucine M Met Methionine N Asn Asparagine P Pro Proline Q Gln Glutamine R Arg Arginine S Ser Serine T Thr Threonine V Val Valine W Trp Tryptophan Y Tyr Tyrosine r D-Arg D-Arginine Fx - Cyclohexylalanine xiv LIST OF PUBLICATIONS... a result of employing organic dyes in peptide- or small-molecule based substrates Continued research is certainly necessary in pushing the frontiers of enzyme assays and bioimaging to include enzymes and applications that have not yet been accessible 14 CHAPTER 2 DEVELOPING NEW FLUOROGENIC SUBSTRATES FOR DETECTING PROTEASE ACTIVITY 2.1 Fluorogenic Protease Substrates for Detecting Protease Activity... enzymatic activity In a similar vein, Xiong and co-workers incorporated a radionuclide in a precedent fluorogenic caspase probe for both fluorescence and nuclear imaging in preliminary in vivo imaging experiments [21] Given that substrates used in high throughput screening and those that are suited for bioimaging differ in the type of fluorophore, our group probed the possibility of developing fluorogenic... emerged as indispensable tools in the profiling and visualization of protease activities both in vitro and in vivo [31] Two types of synthetic fluorogenic peptides are widely used in high-throughput screening of protease inhibitors: i) extended FRET-based peptide substrates containing fluorophore and a dark quencher and ii) fluorogenic peptide substrates containing a C-terminally capped coumarin derivative... Uttamchandani, M.; Li, J.; Sun, H.; Yao, S Q Activity-based profiling: new developments and directions in protein fingerprinting Chembiochem 2008, 9, 667-675 5 Srinivasan, R.; Li, J.; Ng, S L.; Kalesh, K A.; Yao, S Q Methods of using click chemistry in the discovery of enzyme inhibitors Nat Protocols 2007, 2, 26652664 6 Lee, W L.; Li, J.; Uttamchandani, M.; Sun, H.; Yao, S Q Inhibitor fingerprinting of... as their suitability for both quantitative analyses for real-time monitoring of enzyme kinetics and for 1 visual tracking of enzymatic activity The proven utility of these assays has driven active research in designing and/ or modifying fluorescent proteins, inorganic nanoparticles and small molecule organic fluorophores for use in these assays Enzyme assays with fluorescence-based detection methods are... diverse array of serine proteases The screening platform thus established by these groups has since become a reliable tool 8 for probing protease substrate preferences and generating a “fingerprint” for each protease under study, thereby allowing the differentiation of closely-related enzyme The Ellman group then took a step in the direction of assay miniaturization for highthroughput screening with large... preferences for individual peptides on a miniaturized fluorogenic assay This had the potential of generating a proteolytic fingerprint of each protease rapidly, using minimal amounts of enzymes and substrates, in a single experiment At the same time, our group independently prepared a complementary microarray platform for the detection of proteases and other hydrolytic enzymes, such as alkaline phosphatases,... MMP activity in tumors The work by Weissleder and co-workers is considered an important advance in clinical molecular imaging and set the stage for developing similar imaging strategies and techniques targeting other enzymes In contrast to quenching, fluorescence resonance energy transfer (FRET) is a result of long range dipole-dipole interaction between the donor and acceptor, resulting in the excess... suited for both in vitro assays and live cell imaging We designed and synthesized a new green-emitting fluorophore which could be used as a coumarin substitute in microplate- and microarray-based assays, and also in live-cell imaging of apoptosis [22] This work is the subject of Chapter 2 in this thesis 11 1.2.3 Fluoromorphic probes In contrast to assays for hydrolytic enzymes such as proteases and exoglycosidases, .. .DEVELOPING NEW FLUOROPHORES FOR APPLICATIONS IN PROTEASE DETECTION AND PROTEIN LABELING LI JUNQI A THESIS SUBMITTED FOR THE DEGREE OF MASTER OF SCIENCE DEPARTMENT... Phenylalanine G Gly Glycine H His Histidine I Ile Isoleucine K Lys Lysine L Leu Leucine M Met Methionine N Asn Asparagine P Pro Proline Q Gln Glutamine R Arg Arginine S Ser Serine T Thr Threonine V... serine proteases The screening platform thus established by these groups has since become a reliable tool for probing protease substrate preferences and generating a “fingerprint” for each protease