Arts and Sciences, Irving K. Barber School of (Okanagan)
DSpace
UBCO
Slattery, Wyatt Thomas
Master of Science - MSc
University of British Columbia
Integrins are transmembrane receptors that mediate cellular adhesion to the extracellular matrix and initiate signaling cascades that may be relevant to synaptic elimination events occurring in neurodegenerative disease. To study the role of integrins in synaptic elimination, a novel ligand-directed labeling molecule has been created, comprised of the Pacific Blue™ (PB) fluorophore and the high-affinity integrin inhibitor, cilengitide. It was hypothesized that the function of integrin αVβ5 could be studied in live cells by using this probe to track subcellular integrin localization through fluorescence microscopy. This study evaluated the pharmacodynamic properties of the PB-probe, finding that its fluorescence excitation and emission properties are similar to PB and highly photostable over the time period of at least 2 h. Fluorescence enhancement and anisotropy experiments revealed that a derivative of the probe binds integrin αVβ5 with a dissociation constant of 0.11 ± 0.07 μM. Binding was competitively inhibited by cilengitide, suggesting that both molecules compete for the integrin binding domain with similar affinities. Application to BV-2 murine microglia in vitro confirmed that the PB-probe is a robust and rapidly applicable labeling method for the fluorescence detection of endogenous integrins. These experiments demonstrated that PB-probe labeling is specific, amenable to a variety of sample preparation methods, and compatible with simultaneous immunocytochemistry techniques. Separately, the PB-probe was found to inhibit integrin-mediated signaling cascades with equal efficacy as cilengitide. The results of this study confirm that the PB-probe retains both similar fluorescent properties as PB and the integrin-binding properties of cilengitide. These findings establish the PB-probe as an effective means of tracking endogenous integrin αVβ5 in live-cell systems, representing a critical first step in understanding the role of integrin signaling in synaptic elimination.
Evaluation of a ligand-directed labeling molecule targeting integrin alphaVbeta5 by Wyatt Thomas Slattery A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF Master of Science in The College of Graduate Studies (Biochemistry and Molecular Biology) THE UNIVERSITY OF BRITISH COLUMBIA (Okanagan) December 2018 © Wyatt Thomas Slattery, 2018 ii The following individuals certify that they have read, and recommend to the College of Graduate Studies for acceptance, a thesis/dissertation entitled: Evaluation of a ligand-directed labeling molecule targeting integrin alphaVbeta5 Submitted by Wyatt Thomas Slattery in partial fulfillment of the requirements of the degree of Master of Science. Dr. Frederic Menard, Irving K. Barber School of Arts and Sciences Supervisor Dr. Andis Klegeris, Irving K. Barber School of Arts and Sciences Supervisory Committee Member Dr. Kirsten Wolthers, Irving K. Barber School of Arts and Sciences Supervisory Committee Member Dr. Michael Deyholos, Irving K. Barber School of Arts and Sciences University Examiner iii Abstract Integrins are transmembrane receptors that mediate cellular adhesion to the extracellular matrix and initiate signaling cascades that may be relevant to synaptic elimination events occurring in neurodegenerative disease. To study the role of integrins in synaptic elimination, a novel ligand-directed labeling molecule has been created, comprised of the Pacific BlueÔ (PB) fluorophore and the high-affinity integrin inhibitor, cilengitide. It was hypothesized that the function of integrin αVβ5 could be studied in live cells by using this probe to track subcellular integrin localization through fluorescence microscopy. This study evaluated the pharmacodynamic properties of the PB-probe, finding that its fluorescence excitation and emission properties are similar to PB and highly photostable over the time period of at least 2 h. Fluorescence enhancement and anisotropy experiments revealed that a derivative of the probe binds integrin αVβ5 with a dissociation constant of 0.11 ± 0.07 µM. Binding was competitively inhibited by cilengitide, suggesting that both molecules compete for the integrin binding domain with similar affinities. Application to BV-2 murine microglia in vitro confirmed that the PB-probe is a robust and rapidly applicable labeling method for the fluorescence detection of endogenous integrins. These experiments demonstrated that PB-probe labeling is specific, amenable to a variety of sample preparation methods, and compatible with simultaneous immunocytochemistry techniques. Separately, the PB-probe was found to inhibit integrin-mediated signaling cascades with equal efficacy as cilengitide. The results of this study confirm that the PB-probe retains both similar fluorescent properties as PB and the integrin-binding properties of cilengitide. These findings establish the PB-probe as an effective means of tracking endogenous integrin aVb5 in live-cell systems, representing a critical first step in understanding the role of integrin signaling in synaptic elimination. iv Lay Summary Disease processes underlying dementia all share in common the targeted death of neurons. In the brain, neuron cells form connective networks that are the basis of thought, memory, and behavior. There is evidence that neuron connections are eliminated during disease and may contribute to neuron death, leading to cognitive decline. This process is suggested to be mediated by a protein called integrin. This thesis describes the first application of a new molecule designed to track integrins in the brain. It has been found that this molecule binds effectively to integrin and is easily detectable with common microscopy techniques. As a rapidly applicable tool to study integrins in live-cell systems, this molecule improves on existing labeling methods and will enable future investigation of the role of integrins in dementia. v Preface All organic syntheses presented in this thesis were performed by Dr. Vojtech Kapras. Chapter 4 is based on work conducted in collaboration with Dr. Vojtech Kapras using instruments from the Algar Research Group laboratory at the University of British Columbia Vancouver. I assisted with sample preparation and experimental design. Chapter 5 includes data collected by Dr. Pui Ying Lam of the Peterson Lab at the University of Utah. I provided instruction for the application of the Pacific BlueÔ-probe. Select data from Chapters 4-6 are included in an original research article that is nearing submission to Cell: Chemical Biology. The manuscript was contributed to equally by Dr. Vojtech Kapras and I. An excerpt of this manuscript, written by Dr. Vojtech Kapras, is included as Appendix A. vi Table of Contents Abstract ……………………………………………………………………………………………………………………… iii Lay Summary ……………………………………………………………………………………………………………… iv Preface ………………………………………………………………………………………………………………………. v Table of Contents ………………………………………………………………………………………………………. vi List of Tables ……………………………………………………………………………………………………………… ix List of Figures ……………………………………………………………………………………………………………. x List of Abbreviations …………………………………………………………………………………………………. xiii Acknowledgements …………………………………………………………………………………………………… xv Dedication ………………………………………………………………………………………………………………… xvi 1 Introduction …………………………………………………………………………………………………… 1 1.1 The central nervous system …………………………………………………………………… 1 1.1.1 Glia and synaptic elimination ……………………………………………………… 1 1.2 Integrin heterodimer complexes ……………………………………………………………. 3 1.2.1 Integrins function in signaling events ………………………………………….. 4 1.2.2 Integrins signaling in phagocytosis ……………………………………………… 6 1.3 Methods to monitor proteins in cells ……………………………………………………… 8 1.3.1 Recombinant protein expression ………………………………………………… 9 1.3.2 Endogenous protein labeling ……………………………………………………… 10 1.3.3 Small-molecule labeling techniques …………………………………………… 10 1.4 Cilengitide as a small-molecule integrin inhibitor …………………………………… 11 1.5 Objective of this study ……………………………………………………………………………. 12 1.6 Thesis overview ……………………………………………………………………………………… 14 2 Materials and methods …………………………………………………………………………………… 16 2.1 Reagents ………………………………………………………………………………………………… 16 2.2 Cell culture ……………………………………………………………………………………………… 16 2.3 Data analysis …………………………………………………………………………………………… 17 3 The Pacific BlueÔ-probe retains similar spectrophysical properties as Pacific BlueÔ ……………………………………………………………………………………………… 19 3.1 Outline …………………………………………………………………………………………………… 19 3.2 Experimental methods …………………………………………………………………………… 19 3.3 Results and discussion ……………………………………………………………………………. 21 3.3.1 Effect of cilengitide linkage on Pacific BlueÔ fluorescence …………. 21 vii 3.3.2 Determination of the fluorescence quantum yield of the Pacific BlueÔ-probe ……………………………………………………………………. 23 3.3.3 Effects of solvent on the fluorescence properties of the Pacific BlueÔ-probe ……………………………………………………………… 25 3.3.4 The emission properties of the Pacific BlueÔ-probe are retained labeling cells in vitro ……………………………………………………… 27 3.3.5 Persistence of Pacific BlueÔ-probe fluorescence through time …… 29 3.4 Summary ..………………………………………………………………………………………………. 32 4 The Alexa FluorÒ 568 probe and cilengitide have similar binding affinities ……… 33 4.1 Outline ……………………………………………………………………………………………………. 33 4.2 Experimental methods ……………………………………………………………………………. 33 4.3 Results and discussion …………………………………………………………………………….. 34 4.4 Summary ..………………………………………………………………………………………………. 38 5 The Pacific BlueÔ-probe labels integrin aVb5 in vitro …………………………………….. 39 5.1 Outline ……………………………………………………………………………………………………. 39 5.2 Experimental methods ……………………………………………………………………………. 39 5.3 Results and discussion …………………………………………………………………………….. 41 5.3.1 Optimization of labeling parameters ……………………………………………. 41 5.3.2 Specificity of the Pacific BlueÔ-probe …………………………………………. 50 5.3.3 Application of the Pacific BlueÔ-probe in vivo ……………………………. 58 5.4 Summary ..………………………………………………………………………………………………. 62 6 The integrin labeling probes retain inhibitory effect on integrin aVb5 signaling in vitro ……………………………………………………………………………………………… 63 6.1 Outline ……………………………………………………………………………………………………. 63 6.2 Experimental methods ……………………………………………………………………………. 63 6.3 Results and discussion …………………………………………………………………………….. 66 6.3.1 Validation of BV-2 cells as a model system to evaluate phagocytic activity ………………………………………………………………………. 66 6.3.2 Effects of integrin aVb5 inhibitors on bead engulfment (confocal microscopy) ………………………………………………………………….. 68 6.3.3 Phagocytosis of vitronectin-coated beads quantified using flow cytometry …………………………………………………………………………….. 73 6.3.4 Effects of integrin aVb5 inhibitors on bead engulfment (flow cytometry) ………………………………………………………………………….. 79 viii 6.3.5 Inhibitory effect of an Alexa FluorÒ 568 probe on glial phagocytosis ……………………………………………………………………………….. 81 6.4 Summary ..………………………………………………………………………………………………. 84 7 Conclusion ………………………………………………………………………………………………………. 85 7.1 Limitations ………………………………………………………………………………………………. 86 7.2 Future directions …………………………………………………………………………………….. 87 References …………………………………………………………………………………………………………………. 89 Appendices ………………………………………………………………………………………………………………… 100 Appendix A: Drafted manuscript excerpt ……………………………………………………………. 100 Appendix B: Microglia and neurodegenerative disease ………………………………………. 102 Appendix C: Confocal microscopy acquisition parameters ………………………………….. 103 ix List of Tables Table 1.1 Dictionary of compounds used in this work …………………………………………. 13 Table 3.1 The Pacific BlueÔ-probe and Pacific BlueÔ have similar fluorescence quantum yields ………………………………………………………………… 25 Table 5.1 Integrin labeling by the Pacific BlueÔ-probe and immunocytochemistry techniques share 59% colocalization ………………… 56 Table 6.1 Physical attributes of polystyrene latex beads ……………………………………… 78 x List of Figures Figure 1.1 Cladogram of human integrin subunits ………………………………………………… 3 Figure 1.2 Outside-in integrin signaling cascade …………………………………………………… 4 Figure 1.3 Inside-out integrin signaling cascade ……………………………………………………. 6 Figure 1.4 Crystal structure of integrin aVb3 ………………………………………………………… 6 Figure 1.5 Immunostain of integrin subunits in adult rat retinal pigment epithelium …………………………………………………………………………………………… 7 Figure 1.6 Integrin aVb5 phagocytic signaling cascade …………………………………………. 8 Figure 1.7 Chemical structures of small-molecule labels ………………………………………. 11 Figure 3.1 The fluorescence excitation l max of the Pacific BlueÔ-probe and Pacific BlueÔ-linker are red-shifted 7.5 nm relative to Pacific BlueÔ …… 22 Figure 3.2 The fluorescence emission l max of the Pacific BlueÔ-probe and Pacific BlueÔ-linker are red-shifted 3 nm relative to Pacific BlueÔ ……… 22 Figure 3.3 Overlaid excitation and emission spectra ……………………………………………… 23 Figure 3.4 Fluorescence profiles of Pacific BlueÔ and Pacific BlueÔ-probe solutions supporting quantum yield calculations …………………………………. 25 Figure 3.5 Using F10 medium causes 2.5 nm a blue-shift in Pacific BlueÔ-probe excitation l max ………………………………………………………………………………….. 26 Figure 3.6 Using F10 medium causes 2.5 nm a red-shift in Pacific BlueÔ-probe emission l max ……………………………………………………………………………………. 27 Figure 3.7 Overlaid excitation and emission spectra …………………………………………….. 27 Figure 3.8 The emission properties of the Pacific BlueÔ-probe are retained labeling cells in vitro ……………………………………………………………………………. 28 Figure 3.9 Illumination of the Pacific BlueÔ-probe in F10 over 2 h results in a 20.5% increase in emission intensity …………………………………………………. 30 Figure 3.10 Illumination of F10 over 2 h results in no increase in emission intensity at 459 nm ……………………………………………………………………………… 31 Figure 3.11 Illumination of the Pacific BlueÔ-probe in PBS over 2 h does not affect emission intensity ……………………………………………………………………… 32 Figure 4.1 Fluorescence enhancement of the Alexa FluorÒ 568 probe upon binding to integrin aVb5 ……………………………………………………………………… 35 Figure 4.2 Effect of tumbling rate on emission anisotropy ……………………………………. 36 xi Figure 4.3 Fluorescence anisotropy increases upon binding of the Alexa FluorÒ 568 probe to integrin aVb5 ……………………………………………………… 37 Figure 4.4 Competitive binding of cilengitide reduces the fluorescence anisotropy of Alexa FluorÒ 568 probe bound to integrin aVb5 ……………. 38 Figure 5.1 Fluorescence intensity of labeled cells in vitro increases proportionally with Pacific BlueÔ-probe concentration ………………………. 43 Figure 5.2 Fluorescence intensity of labeled cells in vitro increases proportionally with Pacific BlueÔ-probe incubation time …………………… 45 Figure 5.3 The Pacific BlueÔ-probe is amphiphilic ………………………………………………. 46 Figure 5.4 Fluorescence intensity of labeled cells in vitro changes with permeabilization but not wash number ………………………………………………. 48 Figure 5.5 Prolonged illumination of BV-2 cells effects morphology …………………….. 50 Figure 5.6 Effect of cilengitide on the fluorescence intensity of cells labeled with the Pacific BlueÔ-probe ……………………………………………………………… 51 Figure 5.7 Fluorescent labeling is most intense in non-permeabilized cells treated with the Pacific BlueÔ-probe …………………………………………………. 52 Figure 5.8 Fluorescent labeling occurs following treatment with cytochalasin B ….. 55 Figure 5.9 Immunocytochemistry and the Pacific BlueÔ-probe label cells simultaneously …………………………………………………………………………………… 56 Figure 5.10 Zebrafish embryo 72 h post-fertilization ……………………………………………… 58 Figure 5.11 The Pacific BlueÔ-probe does not label macrophages in zebrafish embryos following direct incubation …………………………………………………… 59 Figure 5.12 The Pacific BlueÔ-probe colocalizes with macrophages in zebrafish embryos following injection into the yolk ……………………………………………. 60 Figure 5.13 The Pacific BlueÔ-probe does not label macrophages in zebrafish embryos following injection into circulation ………………………………………… 61 Figure 6.1 Effect of lipopolysaccharide-stimulation on latex bead engulfment by BV-2 microglia ……………………………………………………………… 67 Figure 6.2 Effect of cilengitide on vitronectin-coated bead engulfment by unstimulated BV-2 microglia ……………………………………………………………….. 68 Figure 6.3 Cilengitide blocks the uptake of vitronectin-coated beads by stimulated BV-2 microglia ……………………………………………………………………. 69 Figure 6.4 The Pacific BlueÔ-probe blocks the uptake of vitronectin-coated beads by stimulated BV-2 microglia ………………………………………………………………. 70 Figure 6.5 Cilengitide and the Pacific BlueÔ-probe are not cytotoxic …………………… 71 xii Figure 6.6 The effect of cilengitide and the Pacific BlueÔ-probe on bead engulfment is amplified at high levels of phagocytosis ………………………… 72 Figure 6.7 Engulfment of vitronectin-coated beads is actin-dependent and more efficient than uncoated beads ……………………………………………………. 74 Figure 6.8 Treatment with cytochalasin B reduces BV-2 cell viability …………………… 76 Figure 6.9 Time elapsed before flow cytometry analysis does not affect fluorescence intensity …………………………………………………………………………. 77 Figure 6.10 The number of beads incubated per cell affects fluorescence intensity …………………………………………………………………………. 79 Figure 6.11 Cilengitide and the Pacific BlueÔ-probe are equally effective at blocking vitronectin-coated bead engulfment ……………………………………… 80 Figure 6.12 Cilengitide and the Alexa FluorÒ 568 probe block vitronectin-coated bead engulfment …………………………………………………… 82 Figure 6.13 The Alexa FluorÒ 568 probe is not cytotoxic ………………………………………. 83 Figure 7.1 Second-generation Pacific BlueÔ-probe concept ………………………………… 88 Figure A.1 Synthesis of a modified cyclo(-RGDf-MeV-) bearing a synthetic handle for molecular cargo attachment ………………………………………………. 101 xiii List of Abbreviations Ab Amyloid beta AD Alzheimer’s disease ANOVA Analysis of variance AU Arbitrary units BF Band-pass filter C.A. Condenser aperture Cas9 CRISPR associated protein 9 CBS Calf bovine serum CNS Central nervous system CRISPR Clustered regularly interspaced short palindromic repeats cyclo(-RGDf-MeV-) Cyclic RGD tripeptide coupled to N-methylated D-phenylalanine and valine d Diameter (µm) DAPI 2-(4-amidinophenyl)-1H-indole-6-carboxamidine deg. Degree DM Dichroic mirror DMEM/F12 Dulbecco’s modified Eagle medium-nutrient mixture F12 Ham DMF N,N-dimethylformamide DMSO Dimethylsulfoxide EC50 Concentration resulting in 50% activity stimulation ECM Extracellular matrix EDTA Ethylenediaminetetraacetic acid F10 DMEM/F12 media supplemented with 10% CBS, 100U/mL penicillin, and 100µg/mL streptomycin FAK Focal adhesion kinase FITC Fluorescein isothiocyanate FSC Forward scatter Gas6 Growth arrest-specific 6 GFP Green fluorescent protein HA Hemagglutinin IC50 Concentration resulting in 50% activity inhibition IgG Immunoglobulin G KD Dissociation constant logP Octanol-water partition coefficient xiv LPS Lipopolysaccharide LSM Laser scanning microscope Mag. Magnification MERTK Proto-oncogene tyrosine-protein kinase MER MES 2-(4-morpholino)-ethane sulfonic acid MFG-E8 Milk fat globule-EGF factor 8 MCC Mander’s colocalization coefficient MTT 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide NA Numerical aperture PB Pacific BlueÔ PBS Phosphate buffered saline PCC Pearson correlation coefficient PET Positron emission tomography PHANTAST Phase contrast microscopy segmentation toolbox F Fluorescence quantum yield PI Propidium iodide PKC Protein kinase C PMT Photomultiplier tube RGD Arginine-glycine-aspartate tripeptide SDS Sodium dodecyl sulfate S.E.M. Standard error of the mean SSC Side scatter TBS Tris buffered saline TBSCa TBS supplemented with 2 mM calcium chloride, 0.1% sodium azide, and 100 mM octyl-β-d-glucopyranoside TRITC Tetramethylrhodamine xv Acknowledgements I offer my enduring gratitude to the faculty and staff at UBC, who have supported me in all of my pursuits. I am grateful to Dr. Frederic Menard for his tireless effort in helping me become both a better scientist and individual. I am thankful to Dr. Andis Klegeris for providing keen insight on my research and supporting my academic pursuits as well as my personal development. I owe a particular debt of gratitude to Dr. Vojtech Kapras, without whom this research would not be possible. Similar thanks are offered to Dr. Jonathan Little, Dr. Russ Algar, Dr. Pui Ying Lam, and Dr. Kirsten Wolthers for sharing equipment and expertise. Special thanks are extended to my family and friends who have supported me along the way. xvi In loving memory of Nat Slattery 1 1 Introduction 1.1. The central nervous system The central nervous system (CNS) is composed of two main cell types: neurons and glia. Neurons are the electrically active cells of the brain that form elaborate networks with one another to facilitate information processing [1]. Electrical and chemical signals transmitted throughout neuron networks in the CNS are the basis of cognition, movement, and sensation [2]. Glial cells are generally considered to support neurons and are broken down into three subcategories: oligodendrocytes, astrocytes, and microglia – all of which help maintain CNS homeostasis [3]. Oligodendrocytes form myelin sheaths around neuron axons, facilitating saltatory conduction and increasing action potential velocity [4]. Astrocytes are closely involved with the maintenance of neuronal synapses (aiding in synaptic transmission and providing metabolic support to neurons) as well as the blood-brain barrier [5]. Lastly, microglia participate in CNS homeostasis as innate immune cells; they respond to pro-inflammatory stimuli with the secretion of cytotoxic factors and phagocytosis directed at eliminating foreign substances or cellular debris [6]. 1.1.1 Glia and synaptic elimination Glial populations including microglia and astrocytes conduct synaptic elimination processes in a variety of physiological and pathological conditions [7]. In physiology, synaptic pruning plays a critical role throughout development in which abundant synaptic connections are selectively refined to form mature neural circuits [8, 9]. Moreover, learning and memory processes throughout life are dependent on synaptic plasticity resulting from both the formation of new synapses and the elimination of those that are old and weak [10]. The evidence that both astrocytes and microglia eliminate synapses through phagocytic processes in vivo is robust; however, the role this process plays later in life and throughout neuropathology is less clear [11]. It has been described in Alzheimer’s disease (AD) that astrocytes and microglia phagocytose plaques of aggregated amyloid b (Ab), functioning to clear this disease hallmark with potentially beneficial effect [12]. Separately, it has been demonstrated in vitro that microglia stimulated with lipopolysaccharide (LPS) actively induce neuronal death through phagocytosis, even in cases where neurons are otherwise viable [13-16]. Microglia are considered to exist in one of two phenotypic states, termed M1 and M2. Microglia with an M2 phenotype are considered essential for maintaining CNS homeostasis and exert neuroprotective effects through inflammatory resolution and repair processes [17, 18]. 2 However, exposure of microglia to pro-inflammatory stimuli (such as LPS) induces a phenotypic change to an M1 state, characterized by alterations in the secretory profile and increased phagocytic activity [13, 19, 20]. While these responses are aimed at eliminating pathogens within the CNS, chronically activated M1 pro-inflammatory microglia may lead to irreversible neuron loss [17]. It is unclear whether different subpopulations of M1 or M2 microglia may exert beneficial and deleterious effects on neuron viability through phagocytic processes simultaneously, or whether there might be a transition from a neuroprotective role to one that is neurotoxic throughout pathogenesis. For a more complete overview of microglia in inflammatory processes relevant to neurodegenerative disease, the reader is referred to Appendix B. As professional phagocytes, microglia rely on both the complement system and signaling cascades initiated by the proto-oncogene tyrosine-protein kinase MER (MERTK) to mediate synaptic elimination [21, 22]. Separate studies have identified that involvement of the integrin aVb5 heterodimer is necessary for the effective initiation of phagocytic cascades by MERTK [23]. Significantly, a transcriptomic analysis of glia has revealed that all proteins involved in the MERTK-integrin aVb5 phagocytic signaling cascade are highly expressed by astrocytes during active pruning phases of brain development [24]. This suggests that astrocytes can act as phagocytes and be active participants in synaptic elimination processes. Considering that both microglia and astrocytes possess phagocytic machinery and participate in synaptic pruning, we hypothesize that this process actively induces neuron death throughout CNS pathology. This hypothesis is consistent with the findings that activated glia have an increased propensity for phagocytosis following inflammatory insult and that neurons may reversibly expose signals of cell damage in response to such conditions, triggering their own engulfment [13, 25]. It is likely that signals mediated by integrin aVb5 play a central role in such a process; however, a more precise evaluation of integrin aVb5 functions throughout physiological and pathological conditions is necessary to confirm or refute this hypothesis. Thus far, related fields of neuroscience have focused extensively on the effects of stimuli or therapies on neuronal viability in disease models, while largely neglecting the underlying mechanisms. Deducing the role of proteins central to both synaptic elimination and neuron phagocytosis will likely indicate future therapeutic targets to block neurodegenerative disease processes. Importantly, such a study may also provide insight into a critical threshold at which the effect of glia-mediated phagocytosis transitions from neural circuit refinement and synaptic plasticity to irreversible neuronal loss. 3 1.2 Integrin heterodimer complexes Integrins are a diverse family of transmembrane receptors that derive their name from integrating extracellular matrix (ECM) constituents with the intracellular cytoskeleton [26]. In mammals, 18 unique a subunits combine with 8 unique b subunits to form 24 obligate ab heterodimers that are distinct in both function and tissue distribution [26]. While all integrin heterodimers function as the primary mediators of cell adhesion to the ECM, individual subunits combine differentially to impart various ligand specificities and transduce signals through unique pathways [27]. Integrin heterodimers are broadly separated into four functional categories based on their expression and ligand-binding specificities: leukocyte-specific receptors, laminin receptors, GFOGER-binding collagen receptors, and RGD-binding receptors [28]. The latter two categories are named after the single-letter amino acid sequences recognized by the integrin. Despite their functional classifications, each integrin heterodimer shares a certain degree of interrelatedness or homology (Fig. 1.1). Figure 1.1 Cladogram of human integrin subunits Human A) a and B) b integrin subunits were aligned using Clustal2.1 Omega. Numerical values indicate relative sequence identity. The sum of horizontal line values linking two subunits equating to 1.00 indicates no similarity whereas a value of 0.00 indicates complete sequence identity. Subunits boxed in red have been identified to form heterodimers that bind the RGD-motif. Main Point: Following the identification of residues critical to integrin signaling in Figure 3., I think it would be a great transition to discussing the implications of proper and improper (both underactive and overactive) signaling of alphaV beta5 in endocytic and immunological processes; specifically, in the MerTK phagocytic pathway which is upregulated in astrocytes. 0.02301 b4 b2 b7 b1 b8 b6 b3 b5 0.01354 0.02919 0.35464 0.26302 0.26352 0.25431 0.02883 0.03125 0.22806 0.21240 0.25359 0.34402 a11 a10 a1 a2 aE aL aMaD aX a3 a6 a7 aiib a5 aV a8 a4 a9 0.02047 0.054500.017250.007560.023510.27943 0.28409 0.28801 0.30007 0.36437 0.33687 0.19789 0.17562 0.17093 0.30933 0.24993 0.25792 0.31341 0.26576 0.26444 0.26151 0.29054 0.28737 0.03567 0.02553 0.02684 0.05608 0.00611 0.07339 0.04564 0.12804 0.05381 0.03240 A B 4 RGD-binding integrins are some of the most extensively characterized due to their involvement in diverse cell signaling events. RGD-binding integrin heterodimers recognize a broad array of ligands that contain the arginine-glycine-aspartate (RGD)-motif, including the extracellular proteins fibronectin, vitronectin, fibrinogen, Von Willebrand factor, osteopontin, and the soluble opsonin milk fat globule-EGF factor 8 (MFG-E8) [26, 29-31]. 1.2.1 Integrins function in signaling events While integrins are known to function in a relatively passive capacity by anchoring the intracellular cytoskeleton to the ECM, they can also mediate complex bidirectional cell signaling cascades in response to appropriate stimuli. In the classical form of signal transduction across a cell membrane, termed “outside-in,” integrin binding to an extracellular ligand initiates intracellular signaling cascades mediated by adapter proteins and soluble kinases (Fig. 1.2) [32]. Such cascades have broad implications for cellular remodeling. Integrin-mediated sensing of the chemical composition and mechanical status of the ECM has significant effects on development, angiogenesis and thrombosis, differentiation, proliferation, adhesion, migration and invasion, survival, as well as on the activation of transforming growth factor-b and fibrosis [26, 32, 33]. Figure 1.2 Outside-in integrin signaling cascade Cartoon depiction of integrin aVb5 embedded in a lipid-bilayer cross-section (yellow-grey). Extracellular ligands bind the apex of the heterodimer at the interface of the aV (green) and b5 (blue) subunits. Signals are transduced primarily through the interaction of the b5 cytoplasmic domain with intracellular kinases. 5 Although integrins themselves lack enzymatic activity, associations with soluble kinases such as Src-family kinases, focal adhesion kinase (FAK), phosphoinositide-3 kinase, and mitogen-activated protein kinase allow for the propagation of chemical signals in response to structural or scaffolding properties of the integrin [31, 34, 35]. Additionally, adapter proteins such as the heterotrimeric G protein subunit Ga13 modulate integrin signaling by stimulating the activation of Src-family kinases and by dynamically regulating integrin contractility to promote outside-in signaling [36]. In addition to outside-in signaling, integrin associations with adapter proteins and other kinases are involved in “inside-out” signaling: a process whereby the extracellular integrin domain undergoes a conformation change in response to intracellular interactions to regulate affinity for extracellular ligand (Fig. 1.3) [32]. Current crystal-structure models portray integrins in a bent conformation (Fig. 1.4) which represents its inactive state [37, 38]. Intracellular events leading to the binding of cytoskeletal proteins, such as binding of talin or kindlin to the integrin b cytoplasmic domain, induce global conformational changes that are characterized by the separation of the cytoplasmic integrin tails [31, 32, 37]. Separation of the cytoplasmic integrin tails converts the heterodimer to a fully extended form, representing an active state that is able to bind ligand and initiate signaling cascades [37]. While integrin heterodimers do not need to be fully extended to bind to a ligand, cytoplasmic tails that are artificially fused together (unable to separate from one another) result in the permanent adoption of a bent conformation that cannot bind ligand or signal effectively [37]. 6 Figure 1.3 Inside-out integrin signaling cascade Cartoon depiction of integrin aVb5 embedded in a lipid-bilayer cross-section (yellow-grey). Intracellular cytoskeletal proteins bind the b5 cytoplasmic domain to induce a conformational change from a bent (inactive) state to an extended form able to bind extracellular ligand. The aV and b5 subunits are represented in green and blue, respectively. Figure 1.4 Crystal structure of integrin aVb3 A) Inactive, bent conformation of the extracellular portion of integrin aVb3 bound to RGDF (yellow). The protein is visualized from an intracellular perspective with the pink arrow indicating extracellular space perpendicular to the plasma membrane. B) Zoomed region of the RGD-binding site located at the interface of the aV (green) and b3 subunits (blue, boxed region in A). Protein Data Bank #1L5G [39]. 1.2.2. Integrin signaling in phagocytosis Integrin aVb3 and aVb5 are two RGD-binding heterodimers that are particularly interesting due to their participation in engulfment processes [24, 40]. In retinal pigment epithelial cells, it has A B 7 been reported that integrin aVb5 heterodimers involved in engulfment processes exhibit unique subcellular localization relative to integrin aVb3 heterodimers that mediate cellular adhesion (Fig. 1.5) [41]. It was found that mutually exclusive expression patterns restrict each heterodimer to the apical surface and the basolateral membrane, respectively [41]. If a similar pattern of segregation is true in glia, it is likely that subcellular localization of integrin aVb5 will allow for functional interpretations. Integrin aVb5 expressed on the cell surface away from contact points with the ECM may be involved in engulfment processes, while other populations likely mediate cellular adhesion. Simultaneous labeling strategies targeting adapter proteins, complexing proteins, and related soluble signaling proteins might be necessary to confirm the functional conclusions drawn by integrin localization. Figure 1.5 Immunostain of integrin subunits in adult rat retinal pigment epithelium Reproduced from Finnemann et al. [41]. Integrin b5 was detected only on the apical surface of the retinal pigment epithelium (triangles) while b3 was restricted to the basolateral surface (arrows). Integrin aV was present both apically and basally. Scale bar = 5 µm. Integrin aVb5 has been reported to initiate phagocytic cascades in primary, immature dendritic cells in response to binding MFG-E8, a soluble opsonizing protein that recognizes externalized phosphatidylserine on apoptotic cells [42]. Integrin aVb5 promotes early phagosome formation through an outside-in mechanism. After binding MFG-E8, the integrin b5 cytoplasmic domain associates with FAK then downstream signaling is transduced by a complex of the adapter molecule CrkII, dedicator of cytokinesis (Dock180), and Ras-related C3 botulinum toxin substrate 1 (Rac1, Fig. 1.6) [42, 43]. Functional integrin aVb5 is also required for effective stimulation of phagocytosis by the transmembrane protein MERTK [23]. MERTK has been implicated in phagocytic processes including synaptic elimination and functions by recognizing phosphatidylserine on damaged cells through the soluble protein growth arrest-specific 6 (Gas6) [44]. Importantly, effective endocytosis and FAK association by integrin aVb5 are dependent on prior activation by protein kinase C (PKC) through an inside-out mechanism [45]. This is in contrast to integrin aVb3, which constitutively associates with FAK. It is suggested that prior 8 activation by PKC is necessary for the association of integrin aVb5 with FAK due to the unique b5 cytoplasmic domain that features an eight-amino acid insertion relative to other b subunits [23, 45]. Finally, several reports have indicated that the pro-inflammatory stimulus LPS, a component of the gram-negative bacterial cell well, is a potent inducer of PKC activation [46-49]. Figure 1.6 Integrin aVb5 phagocytic signaling cascade Cartoon depiction of integrin aVb5 embedded in a lipid-bilayer cross-section (yellow-grey). Phosphatidylserine exposed by apoptotic cells is recognized by soluble MFG-E8. Complexed phosphatidylserine and MFG-E8 bind to integrin aVb5, initiating a signaling cascade across the plasma membrane. The integrin b5 cytoplasmic domain interacts directly with FAK, promoting early phagosome formation through the CrkII-Dock180-Rac1 signaling complex. 1.3 Methods to monitor proteins in cells The accurate subcellular localization of endogenous proteins has been the focus of many biological studies as a means for both the discovery of protein-protein interactions and to provide insight into protein function [50, 51]. Most commonly, proteins are labeled through: 1) the introduction of recombinant gene constructs where a target protein is fused to an epitope tag or fluorescent protein, or 2) immunochemistry techniques. Recently, small-molecule labeling methods have been introduced that benefit from both high specificity and rapid applicability [52]. 9 Most precedents in this field have been derived from activity-based profiling of enzymes, or through radioactive modification for subsequent macroscale detection by positron emission tomography (PET) [53, 54]. More recently, this technique has also been demonstrated with fluorescent reporters directed at non-enzymatic proteins [55]. 1.3.1 Recombinant protein expression The fluorescent detection of target proteins can be accomplished through the construction and expression of a recombinant fusion protein consisting of both a protein of interest and a fluorescent protein, such as green fluorescent protein (GFP) [56-58]. These constructs can be introduced to a model system by transfection. This technique benefits from having 100% selectivity; however, it cannot be used to label endogenous proteins and has several other drawbacks. Firstly, it is a technically-intensive process to design an appropriate fusion construct. Once successfully designed, the construct must then be ligated correctly and inserted into a plasmid to yield a functional vector. Purification and sequencing analyses must be conducted at each step. Alternatively, purchasing a functional vector alleviates the need for construct design and development, but does not necessarily circumvent the downsides of recombinant protein expression. Depending on the application, transfection methods may result in cell death or trigger an inflammatory response, inherently altering the model system in study [59, 60]. In the event that transfection can be conducted without issue, the strong promoters commonly employed in vectors result in protein overexpression and mislocalization, relative to proteins derived endogenously [61]. Alternatively, functional fusion proteins can be more effectively introduced using clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR associated protein 9 (Cas9) targeted genome editing. CRISPR/Cas9 gene editing is a method to study proteins through the expression of recombinant constructs in vitro. It is a rapidly growing technique due to the precision and accuracy with which it can be used to modify the genome. It represents a significant improvement over transfection methods as genes can be modified in situ, thereby preventing any issues resulting from overexpression and subsequent mislocalization. However, it too suffers from critical drawbacks that prevent its rapid application to study proteins in a range of cell systems. Firstly, the cellular machinery required for effective genome editing using CRISPR/Cas9 is predominantly expressed during cell division [62]. This limits the timeframe in which the technique can reliably be applied. Moreover, it cannot be guaranteed that the gene of interest will be taken-up and expressed by target cells, limiting the number of proteins in a population that are detectable [63]. These downsides are generally outweighed by the minimal perturbation 10 the method imposes on a cellular environment and the ease of use it offers once a model system has been successfully edited. 1.3.2 Endogenous protein labeling The most common method of labeling endogenous proteins both in vitro and in vivo is using immunodetection techniques. Immunocytochemistry, the in vitro application, involves the binding of a primary antibody, raised against a specific epitope, to its subcellular target. When the primary antibody is itself detectable through fluorescence or radioactive decay, the process is termed direct [64]. More commonly, secondary antibodies are employed that bind the primary antibody and are subsequently detected through fluorescence or other methods, termed indirect [64]. Immunocytochemistry is a fairly rapid technique and is not technically complicated. Two common downsides, however, are firstly that samples must be fixed with formaldehyde or methanol prior to analysis, preventing the longitudinal study of proteins through time [50]. Secondly, there have been several reports of antibodies lacking specificity and labeling off-target proteins [65]. This downside has been circumvented by strict antibody vetting procedures or, alternatively, through the immunodetection of an epitope tag such as hemagglutinin (HA) recombinantly fused to a protein of interest [61, 66]. Importantly though, the process to overcome the issue of antibody specificity by thorough vetting is time consuming and effort-intensive. Alternatively, the detection of epitope tags suffers from the same drawbacks as recombinant protein expression introduced by transfection or CRISPR/Cas9. 1.3.3 Small-molecule labeling techniques Endogenous protein labeling methods employing small molecules are still an underdeveloped technique for the detection and localization of protein targets. Generally, the mass of these molecules is on the order of 1000 g/mol, which is approximately 150 x smaller than an antibody and 27 x smaller than GFP [67, 68]. Structurally, a small-molecule label commonly consists of a highly specific substrate or ligand covalently linked to a reporter motif which may be either radioactive or fluorescent in nature [52, 69]. Several reports have employed the cilengitide moiety as the ligand for labeling molecules directed at integrin, although in these studies it has been coupled exclusively to radioactive substrates for analysis by PET (Fig. 1.7A) [54, 70]. Fluorescent reporters have been well-studied in other small-molecule labeling applications, most notably, for the activity-based profiling of enzymes (Fig. 1.7B) [53]. In these applications, the fluorescent reporter is typically coupled to a substrate through a covalent linkage that is 11 susceptible to enzymatic cleavage. As a result, the fluorescent moiety is covalently linked to an active-site residue of the target enzyme, leading to a robust and specific means of detection at the expense of enzyme function. Other research has targeted non-enzymatic proteins with small-molecules based on potent binding motifs with a high degree of success [52]. Figure 1.7 Chemical structures of small-molecule labels A) Modified cyclo(-RGDFK-) coupled to a radioactive isotope of fluorine, 18F, at the lysine residue [70]. B) Modified arachidonic acid derivative coupled to a coumarin dye by a cleavable linker. Enzymatic activity of phospholipase A2 facilitates the covalent linkage of the fluorophore to the active site of the enzyme [53]. 1.4 Cilengitide as a small-molecule integrin inhibitor Originally reported in 1995, cilengitide (cyclo(-RGDf-MeV-)) is a potent inhibitor of integrin aVb3, aVb5, and a5b1 [71]. Its chemical structure, presented in Table 1.1, is based on the amino acid sequence recognized by RGD-binding integrins and features a cyclization that imparts rigidity. These structural features have been strategically developed to yield one of the most potent RGD-mimetic integrin inhibitors currently available [72]. Cilengitide was synthesized in response to reports that integrin aVb3 plays a critical role in angiogenic processes necessary for tumor development [73]. Initial studies reported that cilengitide is well-tolerated pharmacodynamically and the drug went through phase III clinical trials as a treatment for glioblastoma in 2014 [74, 75]. At the conclusion of these trials, cilengitide was found to lack meaningful anti-tumor activity administered as an adjuvant with temozolomide chemoradiotherapy [75]. It has been suggested that integrin inhibitors and anti-angiogenic therapies may still be useful for cancer treatment; however, this research is ongoing [73, 75]. A B 12 1.5 Objective of this study Current methods for the fluorescent labeling of endogenous integrin aVb5 suffer limitations. No existing technique is both rapidly applicable and suitable to study endogenous integrin aVb5 in live cell systems [61]. To address this gap, two ligand-directed labeling probes for integrin aVb5 have been synthesized that are comprised of a high-affinity integrin inhibitor connected to a fluorophore via a linker motif. The chemical structure of these probes and other compounds used in this work are presented in Table 1.1. 13 Table 1.1 Dictionary of compounds used in this work. Chemical names, chemical structures, cartoon depictions, and how each chemical will be referred to throughout this work are presented. Chemical structures are colored to indicate the integrin aVb5 inhibitor cilengitide (red), Pacific Blueä (PB, blue), and Alexa FluorÒ 568 (purple) motifs. Pacific BlueÔ, PB 6,8-difluoro-7-hydroxycoumarine-3-carboxylic acid Cilengitide Cyclo(-RGDf-MeV-) PB-Linker 2-(6,8-difluoro-7-hydroxycoumarine-3-carboxamido)ethoxyacetic acid PB-Probe PB-linker coupled to cyclo(-RGDf-MeV-) at the D-phenylalanine meta position Alexa FluorÒ 568 Probe Alexa FluorÒ 568 coupled to cyclo(-RGDf-MeV-) at the D-phenylalanine meta position ONHOOOFFOHOHOHOOOOFFOHHNHNNNHNHCOO-HNH2N+H2NO OOOOONHOOOFFOHOHOHOOOOFFOHONHOOOFFOHONHHNHNNNHNHCOO-HNH2N+H2NO OOOOHNNHNHNHNOOOOOCOO-HNH2NNH2+HNONHONH+SO3--O3SCOOHOHNOONNNONHOOOFFOHONHHNHNNNHNHCOO-HNH2N+H2NO OOOOHNNHNHNHNOOOOOCOO-HNH2NNH2+HNONHONH+SO3--O3SCOOHOHNOONNN 14 The primary objective of this study is to evaluate the pharmacodynamic and spectrophysical properties of the Pacific BlueÔ (PB)-probe in order to characterize its utility as a means of labeling endogenous integrin aVb5 in vitro. 1.6 Thesis overview For this investigation, it was first necessary to test the spectrophysical properties of the PB-probe and quantify any deviation from unaltered PB fluorophore. As previous reports indicate that the carboxylic acid at position 3 of PB can be modified without affecting the spectrophysical properties of the molecule, it was hypothesized that the PB-probe would retain similar spectrophysical properties to PB as well [76]. This hypothesis is addressed in Chapter 3; the fluorescence excitation and emission, photostability, and fluorescence quantum yield of the PB-probe were analyzed and compared to PB. The binding affinity of an Alexa FluorÒ 568 probe towards purified integrin aVb5 was determined next. Binding affinity could not be quantified for the PB-probe due to instrumental constraints. Since modification of the meta position on the D-phenylalanine ring of cilengitide is well-tolerated in terms of binding affinity, it was hypothesized that the Alexa FluorÒ 568 probe would have a comparable binding affinity towards integrin aVb5, relative to cilengitide [77]. This hypothesis was investigated in Chapter 4, using fluorescence anisotropy to indicate the fraction of bound and unbound Alexa FluorÒ 568 probe over a range of protein concentrations, in the presence or absence of cilengitide. The labeling properties of the PB-probe towards integrin aVb5 were optimized using BV-2 murine microglial cells in vitro. Chapter 5 evaluates the effects of incubation time, PB-probe concentration, and cell sample preparation on labeling efficiency. Binding specificity of the PB-probe was quantified relative to simultaneous labeling of endogenous integrin aVb5 using immunocytochemistry. Finally, a collaborative effort was pursued in which the PB-probe was applied to zebrafish embryos in vivo through a variety of methods. Together, these results indicate the optimal parameters for labeling endogenous integrin aVb5 with the PB-probe both in vitro and in vivo. The effect of both probes on integrin aVb5-mediated signaling cascades is presented in Chapter 6. Integrin inhibitors, such as cilengitide, have been found to block integrin-mediated signaling events implicated in both pathological and physiological functions [71, 78]. Since the probes were expected to retain their binding affinity towards integrin aVb5 (compared to cilengitide) it was hypothesized that they would maintain their inhibitory effect as well. To test this hypothesis, an experiment was developed in which fluorescent latex beads engaged RGD- 15 binding integrins on LPS-stimulated microglial cells to initiate their own engulfment. Cells were preincubated with a range of concentrations of cilengitide, the PB-probe, or the Alexa FluorÒ 568 probe and inhibitory effects towards integrin aVb5 were quantified through bead fluorescence. 16 2 Materials and Methods 2.1 Reagents 6,8-difluoro-7-hydroxycoumarine-3-carboxylic acid (Pacific BlueÔ, PB), 2-(6,8-difluoro-7-hydroxycoumarine-3-carboxamido)ethoxyacetic acid (PB-linker), PB-linker coupled to cyclo(-RGDf-MeV-) at the D-phenylalanine meta position (PB-probe), and Alexa FluorÒ 568 coupled to cyclo(-RGDf-MeV-) at the D-phenylalanine meta position (Alexa FluorÒ 568 probe) were all synthesized in-house according to reported organic syntheses and the manuscript excerpt included as Appendix A [79]. Phosphate buffered saline (PBS) solution was prepared first as a 10 x stock by dissolving 80.0 g sodium chloride, 2.0 g potassium chloride, 14.2 g sodium phosphate dibasic anhydrous (all from Fisher Scientific, Ottawa, ON, Canada), and 2.4 g potassium phosphate monobasic anhydrous (Sigma-Aldrich, Oakville, ON, Canada) in 1.0 L deionized water. Prior to use, 10 x PBS stock was diluted to a 1 x solution using deionized water, the pH was adjusted to 7.4 using 1 M hydrochloric acid (VWR, Mississauga, ON, Canada) or 1 M sodium hydroxide (Fisher Scientific), and the solution was autoclave-sterilized. Cell culture media (F10) was prepared by supplementing Dulbecco’s modified Eagle medium (Gibco #11995-065) containing 4.5 g/L D-glucose, 4 mM L-glutamine, 15 mg/L phenol red, and 110 mg/L sodium pyruvate with final concentrations of 100 U/mL penicillin, 100 µg/mL streptomycin, and 10% v/v calf bovine serum (CBS #SH3007303, all from Fisher Scientific). Dimethyl sulphoxide (DMSO) was purchased from VWR. Pacific BlueÔ, PB-linker, PB-probe, and Alexa FluorÒ 568 probe were all stored at concentrations above 1 mM in DMSO at -20 oC when not in use. Throughout experimentation, all compounds were light-protected. 2.2 Cell culture Experiments performed in vitro were conducted using artificially immortalized cell cultures [80]. Murine BV-2 microglial cells donated by Dr. Andis Klegeris (Department of Biology, University of British Columbia, Kelowna, BC, Canada) were used to model microglia of the CNS [81]. Cells were grown and maintained in a water-jacketed CO2 incubator (Lab-Line Instruments Inc. #1199-0060, Melrose Park, IL, USA) in 100 mm dishes (Sarstedt Inc. #83.3902, Montreal, QC, Canada). Sterile procedures and cell culture experiments were performed in a class II, type A2 biological safety cabinet. Cell cultures were visualized using a phase-contrast inverted microscope (Motic #AE2000, Kowloon Bay, Kowloon, Hong Kong) and split every 2-3 days when @ 80% confluent. Adherent 17 BV-2 cells were removed from the surface of the cell culture dishes with trypsin as follows. Cell culture media was removed from the dish, then cells were washed once with 7 mL PBS. Next, 1 mL of 0.25% trypsin with 0.2 g/L ethylenediaminetetraacetic acid (EDTA, Fisher Scientific #SH3004202) was added to the cell culture for an ensuing five-minute incubation at 37oC in humidity and 5% CO2 atmosphere. The dish was removed from the incubator and the cell suspension was mixed to dislodge adherent cells. Pre-warmed F10 medium (4 mL) was added to the dish and washed over the growth surface to inactivate the remaining trypsin. Finally, a desired volume of cell suspension was transferred to a new 100 mm dish and mixed with pre-warmed F10 medium to yield a final volume of 10 mL. For use in experiments, BV-2 cells were prepared as outlined above with the following modifications. After the addition of pre-warmed F10 medium (4 mL) to the cell culture dish, an 8 µL sample of cell suspension was counted using an improved Neubauer hemocytometer (QIUJING #02270113, China). A desired number of cells was then added to a 15 mL centrifuge tube (VWR #89039-666) and centrifuged at 500 xg for 7 min using a Heraeus Megafuge 40R centrifuge (Fisher Scientific #75004503). Supernatants were discarded and pelleted cells were suspended in 1 mL of fresh, pre-warmed F10, then diluted with additional F10 to reach a desired concentration of cells. Plating volumes varied by experimental procedure and are outlined in subsequent chapters. 2.3 Data analysis Confocal microscopy images were collected using a Fluoview FV1000 Confocal Microscope (Olympus Canada Inc., Richmond Hill, ON, Canada) and processed using Fiji V1.0 and Adobe Photoshop CC 2018 V19.1.4 [82, 83]. All data were analyzed using Microsoft Excel for Mac V16.13. Graphpad Prism V7.0d software was used to generate figures and conduct one-way and two-way analyses of variance (ANOVA), two-tailed unpaired t test statistical analyses, and regression modeling. Dunnett’s multiple comparisons and Tukey’s multiple comparisons post hoc tests were used to determine statistical significance. Data are presented mean ± standard error of the mean (S.E.M.). P-values less than 0.05 were considered statistically significant. Inhibitor versus response curves were assumed to have a standard Hill slope of -1 and modeled using the following nonlinear regression formula: y = bottom + (top – bottom)/(1 + x/IC50) [2.1] 18 Where top and bottom values represent the upper and lower plateaus of y axis values, respectively. IC50 expresses the concentration (x) resulting in a half-maximal inhibitory effect (IC50). The dependent variable (y) is experiment-specific. 19 3 The PB-probe retains similar spectrophysical properties as PB 3.1 Outline The following chapter characterizes the spectrophysical properties of the PB-probe. This characterization includes the fluorescence excitation maxima, emission maxima, quantum yield, and photostability, along with the effects of physiologically compatible solvents on its excitation and emission spectra. Additionally, the PB-probe is applied to cell culture in vitro, further confirming its spectrophysical properties in a relevant application. 3.2 Experimental methods Fluorescence spectrometry For spectroscopic analyses, PB, PB-linker, or PB-probe stocks were diluted 1000 x with PBS or F10 to generate solutions containing 0.1% DMSO. Each sample was further diluted, as necessary, with 0.1% DMSO in PBS or 0.1% DMSO in F10 to achieve a concentration that would not saturate the fluorescence detector of the spectrophotometer. For each analysis, the light source was attenuated with a 1.0 neutral density filter and a 0.3 neutral density filter where indicated (both from Thorlabs Inc., Newton, NJ, USA). Emission spectra were collected over the range of 400-800 nm with an excitation wavelength of 405 nm. Excitation spectra were collected over the range of 300-800 nm with a detection wavelength of 456 nm. Excitation and detection wavelengths were chosen based on the excitation and emission maxima of PB [84]. Each spectrum represents the mean value of three individual scans. Fluorescence spectra of vehicle solutions were collected on the same day as sample spectra. The fluorescence quantum yield of the PB-probe was determined relative to PB according to literature precedents [85, 86]. The quantum yield of PB has been reported as 0.75 in 0.1 M phosphate buffer pH 10; therefore, the same buffer was employed herein [84]. A 0.1 M phosphate buffer was prepared by mixing 0.1 M solutions of tribasic and monobasic sodium phosphate (both from Sigma Aldrich) to reach a pH of 10. Stock concentrations of PB (7.2 mM) and the PB-probe (1.82 mM) in DMSO were diluted with 0.1 M phosphate buffer until an absorbance of 0.3-0.5 was observed for both compounds following excitation at 396 nm. These solutions were then diluted 50-200 x and emission spectra were collected from 415-700 nm as per established protocols [76]. Fluorescence measurements were performed using a PerkinElmer LS 45 Fluorescence Spectrometer temperature-controlled at 25 oC using a PerkinElmer PTP 1 Peltier Temperature Programmer (PerkinElmer Inc., Waltham, MA, USA). Absorbance measurements were performed 20 using a Varian Cary 100 Conc Ultraviolet-Visible Spectrophotometer at ambient temperature (Agilent Technologies Canada Inc., Mississauga, ON, Canada). Fluorescence and absorbance values were reported in arbitrary units (AU). All samples were prepared in a quartz fluorometer cell (10 mm path-length, Starna Cells, Inc., Atascadero, CA, USA). Cells labeled with the PB-probe in vitro were visualized using confocal microscopy. Data analysis The fluorescence intensity values of vehicle solutions were subtracted from sample spectra to correct for solvent fluorescence and light scatter. Each value of a given sample spectrum was then normalized relative to the maximum fluorescence intensity of that spectrum, except when fluorescence emission intensities were compared. Relative quantum yield was calculated using the relationship between absorbance and fluorescence intensity outlined in the following equation: Fs = (Au/As)(Fs/Fu)Fu [3.1] Where F represents the fluorescence quantum yield, A is the absorbance at 396 nm, and F is the area under the emission curve. Subscripts s and u refer to the standard and to the unknown, respectively. This equation assumes that the refractive indices of the standard and unknown solvents are equivalent, that the absorbance values of both solutions are <0.05, and that the intensity of the excitation light is equivalent [85]. Cell imaging Cell analyses in vitro were conducted using BV-2 cells suspended at 0.25 million cells/mL in F10. 50 µL of cell suspension were added to a pre-sterilized square, 18 mm microscope cover glass 0.13 – 0.17 mm thick (Fisher Scientific #12-542A) and incubated for 2 h at 37 oC in humidity and 5% CO2. Cells were washed three times with PBS and fixed with 4% formaldehyde (Fisher Scientific) in F10 at room temperature for 15 min prior to an additional three washes with PBS. BV-2 cells were treated with 1 µM PB-probe and incubated for 3 h at room temperature in a light-protected environment. Cells were washed twice more with PBS and mounted on rectangular, 25 x 75 mm microscope slides 1 mm thick (Fisher Scientific #12-550-A3) with 80% glycerol in PBS for visualization. The phase contrast microscopy segmentation toolbox (PHANTAST) plugin for Fiji was used to segment confocal microscopy images into cell area and background for analysis of fluorescent signal (PHANTAST parameters: s = 1.2, e = 0.02) [87]. 21 3.3 Results and discussion The wavelengths of light absorbed and reemitted by a molecule through fluorescence are intimately linked to the extent of p-conjugation within its chemical structure [88-90]. The chemical structure of PB was inherently altered upon covalent linkage to cilengitide in the synthesis of the PB-probe. It was therefore necessary to characterize the spectrophysical properties of the PB-probe, relative to PB. Specifically, any differences in the fluorescence excitation, emission, or quantum yield would be of critical importance prior to the application of the PB-probe in fluorescence microscopy experiments. Although the design of the covalent linkage between PB and cilengitide did not extend the p-conjugation of the coumarin moiety, small alterations in chemical structure of PB are known to affect its spectrophysical properties [76, 84]. Nonetheless, it was hypothesized that the fluorescence properties of the PB-probe, along with its fluorescence quantum yield, would remain similar to PB. It was additionally hypothesized that the fluorescence excitation and emission properties of the PB-probe would be persistent through time in both PBS and F10. These solvents were chosen as they are physiologically compatible and as such, were used in subsequent experiments to label cells in vitro. 3.3.1 Effect of cilengitide linkage on PB fluorescence Knowledge of the fluorescence excitation and emission profiles of the PB-probe are prerequisite for its application as a tool to study endogenous integrin aVb5 both in vitro and in vivo. To this end, the excitation and emission spectra of PB, PB-linker, and PB-probe were characterized in the physiologically relevant buffers PBS (0.1% DMSO) and F10 (0.1% DMSO). Gratifyingly, Figure 3.1 confirms that these compounds all have very similar excitation profiles in PBS. The excitation maximum of PB (398 nm) is slightly lower than that of the PB-linker (404 nm) and PB-probe (406 nm), both of which have maxima that are within the detection limit of the spectrophotometer (± 1 nm) from each other. Figure 3.2 depicts that the emission maxima of all three compounds are similar as well, with the maximum emission of the PB-probe at a slightly higher wavelength (459 nm) than PB (455 nm) and PB-linker (456 nm). A similar red-shift has been previously reported to occur when the carboxylic acid at position 3 of PB is modified to a more electron-rich ester group [84]. This trend is seen to continue with modification of PB to a similarly electron-rich amide group in the PB-linker and PB-probe. The similarities between the excitation and emission spectra of all three compounds are further supported in Figure 3.3, where all six spectra are merged for easier comparison. 22 Throughout the fluorescence spectra presented in Figures 3.1 – 3.3, blank subtraction does not completely correct for the background signal resulting from light scatter at the excitation or emission wavelengths (405 nm and 456 nm). Every effort was made to ensure consistency between sample and blank preparations, including using the same aliquots of buffer and measuring blanks on the same day as the samples; however, it has been previously reported that the blank intensity primarily due to light scatter varies relative to samples [91]. It was deemed that blank subtraction was a sufficient correction method for the purpose of this study as the analyses herein focus on the comparison of excitation and emission maxima, which are remote from the light scatter peaks. Figure 3.1 The fluorescence excitation l max of the PB-probe and PB-linker are red-shifted 7.5 nm relative to PB A) Fluorescence excitation spectra of PB, PB-linker, and PB-probe in PBS buffer supplemented with 0.1% DMSO. Emission was detected at 456 nm. B) Zoomed region of the fluorescence excitation spectra with labeled maxima (boxed region in A). The mean value of three individual scans from one independent experiment are presented. Figure 3.2 The fluorescence emission l max of the PB-probe and PB-linker are red-shifted 3 nm relative to PB A) Fluorescence emission spectra of PB, PB-linker, and PB-probe in PBS buffer supplemented with 0.1% DMSO. Fluorophores were excited at 405 nm. B) Zoomed region of the fluorescence emission spectra with labeled maxima (boxed region in A). The mean value of three individual scans from one independent experiment are presented. 400 500 600 700 8000.00.51.0λ (nm)Normalized Fluorescence Intensity (AU)PBPB-LinkerPB-Probe390 395 400 405 410 4150.900.920.940.960.981.00λ (nm)Normalized Fluorescence Intensity (AU)397.5 406.0404.0500 600 700 8000.00.51.0λ (nm)Normalized Fluorescence Intensity (AU)PBPB-LinkerPB-Probe445 450 455 460 465 4700.900.920.940.960.981.00λ (nm)Normalized Fluorescence Intensity (AU)454.5 459.0456.0A B A B 23 Figure 3.3 Overlaid excitation and emission spectra Fluorescence spectra of PB, PB-linker, and PB-probe in PBS buffer supplemented with 0.1% DMSO. Fluorophores were excited at 405 nm and emission was detected at 456 nm. Solid lines indicate excitation spectra, dashed lines indicate emission spectra. The mean value of three individual scans from one independent experiment are presented. Together, these spectra confirm that the fluorescence excitation and emission properties of the PB-probe and PB-linker are largely unchanged relative to PB in a simple buffer solution. This is significant for the widespread adoption of the PB-probe as a useful tool for in vitro and in vivo investigations, as the 406 nm excitation maximum corresponds closely with commonly available 405 nm lasers used for fluorescence microscopy. According to these data, excitation of the PB-probe at 405 nm returns fluorescence emission that is 99.8% efficient, relative to its maximum (when detected at 456 nm). It should be cautioned that the spectral properties of the PB-probe do not closely align with some of the most commonly available filter cubes such as 2-(4-amidinophenyl)-1H-indole-6-carboxamide (DAPI), GFP, fluorescein isothiocyanate (FITC), tetramethylrhodamine (TRITC), or Texas Red. Of these, DAPI overlaps most closely with the PB-probe, however the excitation maximum of DAPI at 375 nm only excites the PB-probe with 7.6% efficiency. It is therefore suggested that the PB-probe be applied in vitro and in vivo using its optimal excitation and emission wavelengths of 406 nm and 459 nm, respectively. 3.3.2 Determination of the fluorescence quantum yield of the PB-probe Fluorescence quantum yield is a physical property that relates the number of photons absorbed by a molecule to the number that are subsequently reemitted as light through fluorescence. The fluorescence quantum yield of the PB-probe is significant in the context of protein labeling, as each molecule of the PB-probe is designed to deliver a single fluorophore to its protein target. In this regard, the fluorescence quantum yield of the PB-probe is inherently linked to its effectiveness as a means of labeling endogenous integrin aVb5 since a high fluorescence quantum yield will ease its subsequent detection. The original synthesis of PB reports its quantum 400 500 600 700 8000.00.51.0λ (nm)Normalized Fluorescence Intensity (AU)PBPB-LinkerPB-Probe 24 yield as 0.75. Modification of the carboxylic acid at position 3 to an ester is well-tolerated in terms of fluorescence quantum yield, resulting in a value of 0.78 (4% change) [84]. Since ester and amide groups have similar electronic properties [92], it was hypothesized that the PB-probe would maintain a quantum yield that is similar to PB. In line with this reasoning, previous work has demonstrated that an amide group at position 3 of PB does not adversely affect the fluorescence quantum yield, with the value reported as 0.74 [76]. The data collected for PB and the PB-probe to examine quantum yield are shown in Table 3.1 and spectra are presented in Figure 3.4. While quantum yield can be solved with one data point relating absorbance and fluorescence for each of a standard and an unknown, Figure 3.4 depicts the experimentally-determined absorbance and fluorescence values of both PB and the PB-probe at three different dilutions. In addition to yielding a more precise relationship between absorbance and fluorescence, these data confirm that the fluorophores are significantly dilute to possess a linear relationship between their absorbance and fluorescence (R2 = 0.9999). This linear relationship is necessary to confirm that the quantum yield equation accurately relates the absorbance and fluorescence of the standard and unknown and is typically only achieved at absorbance values <0.05 [86]. Satisfied that this criterion was met, the quantum yield of the PB-probe was calculated as 0.76 ± 0.08 relative to PB, considering that fluorescence quantum yields are commonly accepted with precision no greater than ± 10% error [86, 93-95]. These data confirm the hypothesis that the fluorescence quantum yield of the PB-probe is essentially unchanged relative to PB in PBS. This result is significant in the context of protein labeling, considering that the fluorescence emission of the PB-probe is limited compared with other labeling techniques, such as indirect immunocytochemistry. Using immunocytochemistry, fluorescent signal is amplified when more than one fluorescent secondary antibody binds each primary antibody, delivering several fluorophores to each protein target. Since the PB-probe is currently limited to delivering a single fluorophore to each protein target, the finding that the coumarin dye retains its quantum yield while linked to cilengitide is of critical importance. 25 Table 3.1 The PB-probe and PB have similar fluorescence quantum yields Absorbance readings were collected at 396 nm, which was subsequently used as an excitation wavelength in accordance with previous work on PB [76]. Where F represents the fluorescence quantum yield calculated using equation [3.1], A is the absorbance at the excitation wavelength (396 nm), and F is the area under the emission curve. Data from one independent experiment are presented. Sample Dilution Alex F (AU2) F PB 1 x 0.42242 - 0.75 [84] 50 x - 34801 100 x - 17545 200 x - 8624 PB-Probe 1 x 0.36422 - 0.76 ± 0.08 50 x - 29870 100 x - 14773 200 x - 7154 Figure 3.4 Fluorescence profiles of PB and PB-probe solutions supporting quantum yield calculations A) Pacific Blueä (Alex=0.42242) and PB-probe (Alex=0.36422) solutions in 0.1 M phosphate buffer pH 10 were excited at 396 nm and emission spectra were recorded at 50 x, 100 x, and 200 x dilutions. B) Plot of integrated fluorescence intensity corresponding to A) (calculated as total area under the curve) vs absorbance of PB and PB-probe solutions (R2 = 0.9999). The mean value of three individual scans from one independent experiment are presented. Data were fit with a linear regression. The relationship between fluorescence intensity and absorbance was significantly non-zero; PB P = 0.0053, F = 14472; PB-probe P = 0.0015, F = 185318. 3.3.3 Effects of solvent on the fluorescence properties of the PB-probe Solvent properties such as polarity, viscosity, and the propensity to form hydrogen bonds have all been shown to affect fluorescence excitation and emission properties [96, 97]. It was therefore necessary to investigate whether the composition of a labeling solution amenable to culturing live cells in vitro might affect the spectrophysical properties of the PB-probe. As such, the effects of solvent on the fluorescence excitation and emission properties of the PB-probe were 500 600 7000200400600λ (nm)Fluorescence Intensity (AU) PBPB-Probe0.000 0.002 0.004 0.006 0.008 0.010010000200003000040000AbsorbanceFluorescence Intensity (AU)PBPB-ProbeA B 26 investigated. Figure 3.5 depicts that the PB-probe in PBS (0.1% DMSO) and F10 (0.1% DMSO) have very similar excitation spectra, with the PB-probe in PBS red-shifted 2.5 nm relative to F10. Likewise, Figure 3.6 illustrates that the PB-probe emission maximum in PBS is red-shifted 2.5 nm relative to F10. Interestingly, the shape of emission spectra appears to be solvent-dependent. The exact maximum of the PB-probe emission spectrum in Figure 3.6 exists towards the left side of the emission peak in PBS buffer and towards the right side when measured using F10, forming two distinct “shoulders.” This may suggest the overlap of two emission peaks that are affected differently by changes in solvent. Nonetheless, it can be concluded that any shift in the fluorescence profile of the PB-probe resulting from buffer composition is minimal and inconsequential with regards to experimentally appropriate excitation and detection wavelengths. Figures 3.5 and 3.6 therefore confirm the hypothesis that the fluorescence excitation and emission properties of the PB-probe are largely unchanged in PBS and F10. These two buffering solutions were chosen as they are commonly employed in cell labeling studies and are physiologically compatible [99-102]. The extent of the similarity between the fluorescence properties of the PB-probe in both PBS (0.1% DMSO) and F10 (0.1% DMSO) can be more clearly visualized in the merged excitation and emission spectra presented in Figure 3.7. Figure 3.5 Using F10 medium causes a 2.5 nm blue-shift in PB-probe excitation l max A) Fluorescence excitation spectra of the PB-probe in PBS buffer or F10 supplemented with 0.1% DMSO. Emission was detected at 456 nm. B) Zoomed region of the fluorescence excitation spectra with labeled maxima (boxed region in A). The mean value of three individual scans from one independent experiment are presented. 400 500 600 700 8000.00.51.0λ (nm)Normalized Fluorescence Intensity (AU)F10 (0.1% DMSO)PBS (0.1% DMSO)390 395 400 405 410 4150.900.920.940.960.981.00λ (nm)Normalized Fluorescence Intensity (AU)406.0403.5A B 27 Figure 3.6 Using F10 medium causes a 2.5 nm red-shift in PB-probe emission l max A) Fluorescence emission spectra of the PB-probe in PBS buffer or F10 supplemented with 0.1% DMSO. The PB-probe was excited at 405 nm. B) Zoomed region of the fluorescence emission spectra with labeled maxima (boxed region in A). The mean value of three individual scans from one independent experiment are presented. Figure 3.7 Overlaid excitation and emission spectra Fluorescence spectra of the PB-probe in PBS buffer or F10 supplemented with 0.1% DMSO. The PB-probe was excited at 405 nm and emission was detected at 456 nm. Solid lines indicate excitation spectra, dashed lines indicate emission spectra. The mean value of three individual scans from one independent experiment are presented. 3.3.4 The emission properties of the PB-probe are retained labeling cells in vitro Importantly, the fluorescence emission profile for the PB-probe was reproduced in cell imaging experiments in vitro (Fig. 3.8). Murine microglial cells are known to express vitronectin receptors, of which integrin aVb5 represents @ 60% of the aV surface integrins present on this cell type [103-105]. Artificially immortalized murine BV-2 cells have been shown to accurately model primary microglia and as such, were employed in this study [106]. BV-2 microglia were fixed with formaldehyde and incubated with the PB-probe for 3 h, then washed with PBS and visualized using confocal microscopy. Cells were illuminated with a 405 nm laser and fluorescence was 500 600 700 8000.00.51.0λ (nm)Normalized Fluorescence Intensity (AU)F10 (0.1% DMSO)PBS (0.1% DMSO)445 450 455 460 465 4700.900.920.940.960.981.00λ (nm)Normalized FluorescenceIntensity (AU)459.0 461.5400 500 600 700 8000.00.51.0λ (nm)Normalized Fluorescence Intensity (AU)F10 (0.1% DMSO)PBS (0.1% DMSO)A B 28 observed over ten 10 nm band-pass filter ranges, spaced at 5 nm increments. The mean fluorescence intensity of labeled cells was calculated for each emission range and plotted in Figure 3.8. Figure 3.8 The emission properties of the PB-probe are retained labeling cells in vitro Above) Representative, negative images of labeled cells visualized using 10 nm band-pass filter ranges centered at 440 nm, 455 nm, and 480 nm (Scale bar = 80 µm). Below) Fluorescence emission spectrum of BV-2 cells labeled with the PB-probe for 3 h in PBS buffer supplemented with 0.1% DMSO. Cells were excited at 405 nm and emission was detected over 10 nm band-pass filters spaced at 5 nm increments. Data from one independent experiment are presented. Figure 3.8 indicates that the 10 nm band-pass filter corresponding to the maximum fluorescence emission of the PB-probe is 450-460 nm. This is consistent with what is to be expected for the PB fluorophore, which is reported to have an emission maximum between 450 nm and 457 nm [84, 107]. However, the emission spectra presented in Figures 3.2 & 3.6 indicate that the emission maximum of the PB-probe may be closer to 460 nm. As such, it could be hypothesized that the 10 nm band-pass filter corresponding to the maximum fluorescence emission of the PB-probe might be 455-465 nm. Re-examining Figure 3.8, it can be seen that the difference in mean 400 420 440 460 480 500 52001234Emission Band-Pass Filter Center (λ)Mean Fluorescence Intensity (AU)440 nm nmn 455 nm nm 480 nm 29 fluorescence intensity between the band-pass filters centered at 455 nm and 460 nm is only 2%, indicating that both ranges are appropriate to detect the fluorescence emission of the PB-probe at its maximum. 3.3.5 Persistence of PB-probe fluorescence through time Finally, the persistence of the fluorescence intensity of the PB-probe through time was determined. Pacific blueÔ is reported to have a low propensity for photobleaching [84]. It was therefore expected that the PB-probe would also be stable for extended periods of time following constant illumination at its excitation maximum. This is an important characteristic considering that the ultimate goal for its application is to conduct longitudinal studies through time in vivo. As the PB-probe is not expected to be cell-permeable, due to its two charges at physiological pH, its photobleaching half-life will have important implications in how frequently the PB-probe must be applied and, by extension, how convenient its application will be to study endogenous integrin aVb5 in vivo. The decrease in fluorescence intensity of the PB-probe over time was initially tested in F10 medium (0.1% DMSO). High levels of fluorescent signal were observed in the medium prior to the addition of the PB-probe. This signal was centered at 405 nm, the same wavelength used for excitation (Fig. 3.9 and 3.10). As such, it is most likely that the peak in fluorescent signal produced by F10 medium is due to light scattering [108]. It is possible, however, that this background signal resulted from the fluorescence of tryptophan residues present in the proteins comprising the CBS supplement [109]. Alternatively, the vitamins B2 and B9 (riboflavin and folic acid, respectively) are also both present in DMEM/F12 and F10 and have extended conjugation motifs that are similar to PB [109]. While these media additives are both present in DMEM/F12 and F10 and fluorescently active, it is likely that their fluorescence emission profile would be visually distinguishable from the peaks observed in Figures 3.9 and 3.10, further supporting the conclusion that increased signal association with F10 is a result of light scattering [109-111]. Due to this effect, it was necessary to use 0.3 and 1.0 neutral density filters so as to not saturate the detector. Measurements of the emission spectrum following excitation at 405 nm were collected once every minute for the course of 2 h. Contrary to the initial expectation that the fluorescence intensity of the PB-probe over time would decrease slowly, Figure 3.9 reveals that there was an increase of 20.5% over the course of two hours. 30 Figure 3.9 Illumination of the PB-probe in F10 over 2 h results in a 20.5% increase in emission intensity A) Fluorescence emission spectra of the PB-probe in F10 supplemented with 0.1% DMSO. The PB-probe was excited at 405 nm and scans were collected once every minute for 120 min. B) Zoomed region of the fluorescence emission spectra with labels indicating the lower and upper limits of emission maxima, respectively (boxed region in A). Data from one independent experiment are presented. It was hypothesized that this increase in fluorescence intensity might be due to additives present in F10 media, physical attributes of the fluorimeter, or nonspecific binding of the PB-probe to proteins in solution. In examining the effects of F10 additives, it was possible that the fluorescently active cofactors riboflavin and folic acid were oxidized throughout the experiment, gradually increasing their emission at 456 nm [112]. It is important to note that of the possible medium constituents contributing to the fluorescence of F10, the emission spectrum of folic acid overlaps most closely with PB [109]. With regards to the physical attributes of the fluorimeter, two possible explanations are offered. First, it may be that the xenon flash lamp warmed during the experiment, increasing the power of the excitation light and thereby, the emission of the probe [113]. Second, it could be that initial scans were acquired at ambient temperature and the Peltier system gradually warmed the sample over time, increasing its fluorescence intensity. The latter of these explanations is admittedly unlikely as the dependence of the quantum yield of fluorophores on temperature generally ranges from a 0-1% change in quantum yield per degree Celsius [94, 114]. This difference hardly accounts for the 20.5% change observed in Figure 3.9 following a temperature increase of at most 5 oC, and it is also notable that the quantum yield of coumarin dyes actually decreases with increased solvent temperatures [114]. Fluorophores generally exhibit lower fluorescence quantum yields at higher temperatures due to a proportional increase in intermolecular collisions within a solution. A higher number of intermolecular collisions provides more opportunity for excited fluorophores to return to an electronic ground state via vibrational relaxation, as opposed to fluorescence. To a similar end, fluorophores have been found to exhibit enhanced fluorescence upon binding to 500 600 700 800-2000200400600800λ (nm)Fluorescence Intensity (AU)0 min10 min20 min30 min40 min50 min60 min70 min80 min90 min100 min110 min120 min20.5%445 450 455 460 465 470600650700750800λ (nm)Fluorescence Intensity (AU)458.0 462.5A B 31 macromolecules in solution as this results in less intermolecular collisions with solvent molecules, providing more opportunity for excited fluorophores to return to an electronic ground state through the release of a photon. This phenomenon has been especially well reported for the fluorescence of tryptophan residues either in denatured polypeptides or buried in tertiary/quaternary protein structures [115, 116]. In order to determine which explanation may be responsible for the enhanced fluorescent signal depicted in Figure 3.9, the experiment was repeated using F10 supplemented with 0.1% DMSO in the absence of PB-probe. If the fluorescence enhancement resulted from physical attributes of the fluorimeter or the oxidation of media additives, then a similar increase would be seen in the absence of PB-probe. Figure 3.10 illustrates that no such increase in fluorescence is observed in the absence of PB-probe, suggesting that the increase in fluorescence results from nonspecific binding of the PB-probe to proteins in solution. Figure 3.10 Illumination of F10 over 2 h results in no increase in emission intensity at 459 nm A) Fluorescence emission spectra of F10 supplemented with 0.1% DMSO. The solution was excited at 405 nm and scans were collected once every minute for 120 min. B) Zoomed region of the fluorescence emission spectra (boxed region in A). Data from one independent experiment are presented. To confirm this conclusion, the experiment was repeated to test the time-dependent fluorescence emission of the PB-probe in PBS buffer supplemented with 0.1% DMSO. Measurements of the emission spectrum following excitation at 405 nm were collected once every minute for the course of 2 h. Figure 3.11 indicates that there is no change in the fluorescence intensity of the PB-probe in PBS supplemented with 0.1% DMSO throughout the experiment. These data further support that the increase in fluorescence intensity of the PB-probe in F10 depicted in Figure 3.9 is most likely due to nonspecific binding to, or association with, proteins in solution resulting in fluorescence enhancement. Moreover, Figure 3.11 indicates that the PB-probe is highly photostable following constant illumination for an extended period 400 500 600 700 80005001000λ (nm)Fluorescence Intensity (AU)0 min10 min20 min30 min40 min50 min60 min70 min80 min90 min100 min110 min120 min445 450 455 460 465 470020406080100λ (nm)Fluorescence Intensity (AU)A B 32 of time. This is consistent with reports that PB alone is photostable following 33 min of illumination at its wavelength of maximum absorption [84]. Figure 3.11 Illumination of the PB-probe in PBS over 2 h does not affect emission intensity A) Fluorescence emission spectra of the PB-probe in PBS supplemented with 0.1% DMSO. The PB-probe was excited at 405 nm and scans were collected once every minute for 120 min. B) Zoomed region of the fluorescence emission spectra with labels indicating the lower and upper limits of emission maxima, respectively (boxed region in A). Data from one independent experiment are presented. 3.4 Summary The above experiments confirm the hypothesis that the PB-probe retains similar spectrophysical properties to PB. The excitation and emission maxima of the PB-probe (406 nm and 459 nm, respectively) are slightly red-shifted relative to PB but photostable over the time period of at least 2 h and highly amenable to detection using a commonly available 405 nm laser. These spectrophysical properties are stable in two physiologically compatible buffers and are retained when used to label cell culture in vitro. Moreover, the PB-probe does not have a reduced fluorescence quantum yield relative to PB. One important phenomenon that these data bring to light is that visualization of the PB-probe in cell culture media over time may be accompanied by the gradual increase in background signal most likely due to nonspecific association with proteins in solution. 500 600 700 8000200400λ (nm)Fluorescence Intensity (AU)0 min10 min20 min30 min40 min50 min60 min70 min80 min90 min100 min110 min120 min445 450 455 460 465 470300350400450500λ (nm)Fluorescence Intensity (AU)455.5 463.5A B 33 4 The Alexa FluorÒ 568 probe and cilengitide have similar binding affinities 4.1 Outline This chapter reports the binding affinity of an Alexa FluorÒ 568 probe towards purified integrin αVβ5. Fluorescence enhancement and anisotropy reveal that the Alexa FluorÒ 568 probe binds integrin αVβ5 with a dissociation constant (KD) of 0.11 ± 0.07 µM. A competitive displacement assay showed that cilengitide reduces integrin binding in a concentration-dependent manner. These data support a specific and high-affinity interaction between the Alexa FluorÒ 568 probe and purified integrin αVβ5. 4.2 Experimental methods Reagents Human integrin αVβ5 formulated with 100 mM octyl-β-d-glucopyranoside and 0.1% sodium azide in Tris-buffered saline (TBS) was purchased from Sigma Aldrich. Cilengitide was acquired from Tocris Bioscience (Oakville, ON, Canada). TBS with calcium (TBSCa) was prepared as a solution of 20 mM Tris hydrochloride, 150 mM sodium chloride, 0.1% sodium azide, 2 mM calcium chloride, and 100 mM octyl-β-d-glucopyranoside (all from Fisher Scientific) in deionized water. TBSCa was adjusted to a pH of 7.44 using 1 M hydrochloric acid (VWR) or 1 M sodium hydroxide (Fisher Scientific). Fluorescence enhancement and anisotropy measurements of the Alexa FluorÒ 568 probe were performed using a Tecan INFINITEÒ M1000 Pro microplate reader (Tecan US, Inc., Morrisville, NC, USA) at a constant temperature of 30 oC. Excitation and detection wavelengths of 530 nm and 608 nm (5 nm bandwidths) were chosen based on the fluorescence properties of Alexa FluorÒ 568. Samples were prepared in TBSCa with 5 nM Alexa FluorÒ 568 probe, 100 mM octyl-β-d-glucopyranoside, 0.1% sodium azide, and variable concentrations of integrin αVβ5 (0 nM – 512 nM). All analyses were conducted in opaque 384-well microplates (Corning Inc. #3575, Corning, NY, USA) following a 30 min incubation at 30 oC. Data analysis Total fluorescence intensity of the Alexa FluorÒ 568 probe was calculated as a function of polarized intensities [117], according to the following equation: S = IVV + 2GIVH [4.1] 34 where I is fluorescence intensity with subscripts indicating vertical (V) or horizontal (H) orientation of polarizers and G is the grating factor, a wavelength-dependent value defined as: G = IHV/IVV [4.2] G was measured as 1.33, using the Rhodamine B fluorophore (l max = 610 nm) as a standard. Anisotropy (A) was calculated using the following equation: A = (IVV – GIVH)/(IVV + 2GIVH) [4.3] Dissociation constants were calculated using the following quadratic isotherm equation [118]: A = D(([ITG] + [P] + KD) – ((([ITG] + [P] + KD)^2) – 20[ITG])^0.5)/2[P] + M [4.4] where [P] is the concentration of the Alexa FluorÒ 568 probe (5 nM), [ITG] is the concentration of integrin αVβ5, and D and M represent parameters of the fit corresponding to the range of anisotropy values and the lower plateau of anisotropy values, respectively. Half-maximal inhibitory concentrations were calculated as previously described (equation [2.1]). 4.3 Results and discussion While the binding activity of cilengitide towards integrin αVβ5 has been well-characterized, the PB-probe and the Alexa FluorÒ 568 probe may have reduced binding activities or affinities as a result of their covalent modification with fluorophore. This phenomenon has previously been reported for cilengitide, with changes in amino acid sequence or methylation increasing its IC50 towards integrin αVβ5 function by up to 40-fold [72]. Consequently, it was necessary to quantify the binding affinity of the labeling probes towards purified integrin αVβ5. The binding affinity of the probes towards their integrin target is critically important for their applicability in labeling endogenous protein; high-affinity, specific, interactions such as those between streptavidin and biotin currently represent the gold-standard in protein detection methods [119, 120]. For the analyses presented herein, binding affinity is quantified for the Alexa FluorÒ 568 probe in place of the PB-probe. The Alexa FluorÒ 568 probe was more technically challenging to synthesize and, as such, was only available for select experiments near the end of this thesis work. The Alexa FluorÒ 568 probe improves on the previously outlined PB-probe in several ways. Most notably, the spectrophysical properties of the Alexa FluorÒ 568 fluorophore 35 enable deeper tissue imaging, compared to PB, and the wavelength of its fluorescence excitation maximum is better-tolerated by live-cells [121, 122]. In order to measure the binding affinity of the Alexa FluorÒ 568 probe towards purified integrin αVβ5, a fluorescence enhancement experiment was performed. Briefly, fluorescence enhancement is based on the principle that in solution, a fluorophore bound to a macromolecule such as a protein will have fewer intermolecular collisions with solvent molecules than unbound fluorophore. Consequently, a greater proportion of excited fluorophores return to an electronic ground state through the release of a photon rather than vibrational relaxation, resulting in fluorescence enhancement [115, 116]. Experimentally, fluorescence intensity will increase as a function of protein concentration since the ratio of bound:unbound fluorophore will also increase. Figure 4.1 illustrates that the binding of the Alexa FluorÒ 568 probe to integrin αVβ5 results in a slight enhancement of fluorescence intensity with the KD calculated as 0.1 ± 0.2 µM; however, the trend was not statistically significant. Figure 4.1 Fluorescence enhancement of the Alexa FluorÒ 568 probe upon binding to integrin aVb5 Samples were prepared in TBSCa with 5 nM Alexa FluorÒ 568 probe and variable concentrations of integrin αVβ5 (0 nM – 512 nM). Fluorescence intensity values were calculated using equation [4.1]. Data from three independent experiments are presented as mean ± S.E.M and fit with a nonlinear quadratic isotherm (equation [4.4]). The effect of treatments was assessed by the one-way ANOVA. ANOVA F and P values are presented on the graph. No significant differences were observed. To obtain a more accurate measure of binding affinity, a fluorescence anisotropy experiment was performed. Anisotropy is the fraction of fluorescence emission that is polarized in a defined axis. As a technique, fluorescence anisotropy enables highly precise measurements of binding affinity through changes in light polarization. This experiment operates under the principle that molecules in solution change orientation or “tumble” more rapidly when they are small than when they are large [123]. When polarized light is used to excite a fluorophore, the photons 1 10 100 10008.5×1049.0×1049.5×1041.0×105Integrin αVβ5 (nM)Fluorescence Intensity (AU)F = 0.9317P = 0.5241Alexa Fluor® 568 ProbeKD = 0.1 ± 0.2 μM 36 released are also polarized. Since there is a delay between the electronic excitation and subsequent relaxation of a fluorescent molecule, the polarization of emitted light can be used to calculate the rate of molecular tumbling (Fig. 4.2). Figure 4.2 Effect of tumbling rate on emission anisotropy The Alexa FluorÒ 568 probe is excited with polarized light at 530 nm. Fluorescence emission is detected at 608 nm. In a theoretical case where the tumbling rate of the probe approaches zero, 100% of the fluorescence emission would share the polarization of the excitation light. In contrast, an infinitely fast tumbling rate would allow the probe to adopt random orientations in solution before a photon is released, resulting in a loss of polarization information being emitted by the fluorescing probe. Experimentally, small changes in the tumbling rate of the probe can be calculated by measuring fluorescence emission anisotropy. According to this principle, a probe bound to integrin αVβ5 in solution will change its orientation more slowly than a free probe and therefore, will have a higher anisotropy. Due to instrumental constraints, fluorescence anisotropy could only be quantified for the Alexa FluorÒ 568 probe (polarized filters for the 405 nm excitation laser were unavailable). The results of this experiment are presented in Figure 4.3. The Alexa FluorÒ 568 probe was found to bind integrin αVβ5 with a KD of 0.11 ± 0.07 µM. Anisotropy of the bound fluorophore only increased by approximately 9% relative to the unbound fluorophore; however, this relatively small difference can be explained by the long linker motif imparting several degrees of rotational freedom to the Alexa FluorÒ 568, even in the bound state [123]. 37 Figure 4.3 Fluorescence anisotropy increases upon binding of the Alexa FluorÒ 568 probe to integrin aVb5 Samples were prepared in TBSCa with 5 nM Alexa FluorÒ 568 probe and variable concentrations of integrin αVβ5 (0 nM – 512 nM). Fluorescence anisotropy values were calculated using equation [4.3]. Data from three independent experiments are presented as mean ± S.E.M and fit with a nonlinear quadratic isotherm (equation [4.4]). The effect of treatments was assessed by the one-way ANOVA followed by Dunnett’s multiple comparisons test. ANOVA F and P values are presented on the graph. **P<0.01; significantly different from Alexa FluorÒ 568 probe incubated in the absence of integrin αVβ5. The competitive binding of cilengitide and Alexa FluorÒ 568 probe were tested next for the dual purpose of indicating: 1) that Alexa FluorÒ 568 probe binding can be competitively inhibited by an RGD-domain-binding ligand, and 2) how the binding activity of the Alexa FluorÒ 568 probe compares to vitronectin. Most literature precedent for binding activity of integrin inhibitors is presented in the context of an IC50 value towards ECM ligand-binding. Vitronectin is commonly employed for integrin αVβ5 studies, with cilengitide consistently reported to have an IC50 on the order of 10 nM [72, 78]. Since variation is tolerated at the D-phenylalanine meta position of cilengitide, it was hypothesized that the Alexa FluorÒ 568 probe would bind integrin αVβ5 with a similar activity as unmodified cilengitide (IC50 towards integrin αVβ5 = 11.7 ± 1.5 nM) [77, 78]. Figure 4.4 confirms that binding of the Alexa FluorÒ 568 probe to purified integrin αVβ5 can be competitively inhibited by an RGD-domain-binding ligand. The IC50 value (30 ± 30 nM) of cilengitide is higher than what has previously been reported for vitronectin-binding, suggesting that integrin αVβ5 may bind the Alexa FluorÒ 568 probe with greater activity than its ECM ligand. However, due to the high experimental error of the measurements, it cannot be asserted that the IC50 of cilengitide towards Alexa FluorÒ 568 probe-binding is significantly higher than the 11.7 ± 1.5 nM reported for vitronectin-binding [78]. Nonetheless, these data support the finding that the Alexa FluorÒ 568 probe binds integrin αVβ5 with an activity similar to both vitronectin and cilengitide. 1 10 1000.1200.1250.1300.1350.1400.145Integrin αVβ5 (nM)Fluorescence Anisotropy****F = 6.465P = 0.0001Alexa Fluor® 568 ProbeKD = 0.11 ± 0.07 μM 38 Figure 4.4 Competitive binding of cilengitide reduces the fluorescence anisotropy of Alexa FluorÒ 568 probe bound to integrin αVβ5 Samples were prepared in TBSCa with 10 nM Alexa FluorÒ 568 probe, 100 nM integrin αVβ5, and variable concentrations of cilengitide (0 nM – 200 nM). Fluorescence anisotropy values were calculated using equation [4.3]. Data from three independent experiments are presented as mean ± S.E.M and fit with a nonlinear inhibitor versus response curve (equation [2.1]). The effect of treatments was assessed by the one-way ANOVA followed by Dunnett’s multiple comparisons test. ANOVA F and P values are presented on the graph. **P<0.01; significantly different from Alexa FluorÒ 568 probe incubated with integrin αVβ5 in the absence of cilengitide. 4.4 Summary In conclusion, changes in the fluorescence intensity and anisotropy of the Alexa FluorÒ 568 probe reveal a binding affinity towards integrin αVβ5 of 0.11 ± 0.07 µM. The finding that cilengitide competitively inhibits Alexa FluorÒ 568 probe binding to integrin αVβ5 with an IC50 value of 30 ± 30 nM confirms that its binding activity is similar to both cilengitide and vitronectin. Taken together, these data demonstrate that the Alexa FluorÒ 568 probe retains a high affinity towards integrin αVβ5, supporting its applicability in labeling endogenous integrin αVβ5. 1 10 1000.1800.1820.1840.1860.1880.190Cilengitide (nM)Fluorescence Anisotropy****F = 8.036P = 0.0003Cilengitide IC50 = 30 ± 30 nM 39 5 The PB-probe labels integrin aVb5 in vitro 5.1 Outline This chapter presents the first in vitro applications of a fluorescently active cilengitide molecule modified at the aromatic substituent. The fluorescent activity of the PB-probe confirmed in previous chapters suggested that it could be used to both label endogenous integrin aVb5 in cell cultures and be detectable through confocal microscopy. The effects of incubation time, probe concentration, and cell sample preparation were evaluated in order to fully characterize the labeling properties of the PB-probe and indicate its future applicability to in vivo studies. 5.2 Experimental methods Reagents Immunocytochemistry experiments were conducted using polyclonal rabbit anti-integrin b5 (#ab15459, Abcam Inc., Toronto, ON, Canada) and polyclonal goat anti-rabbit IgG conjugated with DyLightÒ 550 (#84541, Fisher Scientific). Cell culture imaging BV-2 cells were prepared for analysis by confocal microscopy by suspending cells at 0.25 million cells/mL and seeding 50 µL onto pre-sterilized square, 18 mm microscope cover glass 0.13 – 0.17 mm thick (Fisher Scientific #12-542A) in a 2 cm dish (Sarstedt Inc. #83.3900) for a 2 h incubation at 37 oC in humidity and 5% CO2. The cell culture media was then made up to 2 mL with F10 and incubated overnight. Next, the cell culture media was aspirated and cells were washed three times with 2 mL PBS prior to fixation with 2 mL 4% formaldehyde (Fisher Scientific) diluted in F10 for 15 min at room temperature. To prepare non-permeabilized cells, an additional three washes with PBS were conducted. To prepare permeabilized cells, cells were washed 3 x 5 min with 0.1% v/v Triton X-100 diluted in PBS immediately after fixation, then another three times with PBS. In both cases, 1 mL of PBS was added to the cell culture after washing for subsequent labeling. Stock solutions of the PB-probe and PB-linker in DMSO were added directly to the cell culture at a 1000 x dilution for a final DMSO concentration of 0.1% v/v. Permeabilized and non-permeabilized cells were incubated with the PB-probe or PB-linker at room temperature in the dark. Unless otherwise indicated, cells were incubated with 1 µM labeling molecules for 3 h, washed twice with PBS, and mounted on rectangular, 25 x 75 mm microscope slides 1 mm thick (Fisher Scientific #12-550-A3) with 80% glycerol (Sigma Aldrich) in PBS for visualization with confocal microscopy. 40 Alternatively, live BV-2 cell experiments were conducted by first, suspending cells at 0.25 million cells/mL and seeding 2 mL of the cell suspension in glass-bottom dishes (Matsunami Glass USA Inc. #10810-054, Bellingham, WA, USA) for an overnight incubation at 37 oC in humidity and 5% CO2. Stock solutions of the PB-probe and PB-linker in DMSO were then added directly to the cell culture media at a 1000 x dilution for a final DMSO concentration of 0.1% v/v. Cells were incubated with 1 µM PB-probe or PB-linker for 3 h at 37 oC in humidity and 5% CO2. Next, cells were washed twice with PBS and replenished with F10 for visualization with confocal microscopy immediately afterwards. For dual labeling with immunocytochemistry, the above steps for the preparation of permeabilized BV-2 cells were adhered to with the following modifications. After permeabilization with 0.1% v/v Triton X-100 in PBS, cells were washed three times with PBS and incubated overnight at 4 oC with anti-integrin b5 primary antibody diluted 1:25 in 1 mL of 4% CBS (Fisher Scientific) in PBS. The following day, cells were washed 4 x 10 min with PBS and replenished with 1 mL 4% CBS in PBS. The PB-probe was added directly to this solution for a final concentration of 1 µM (0.1% v/v DMSO). Cells were incubated for 1.5 h at room temperature in the dark prior to the addition of 1 µL 1 mg/mL anti-rabbit IgG secondary antibody conjugated to DyLightÒ 550. After another 1.5 h incubation had elapsed, cells were washed twice with PBS and replenished with 1 mL 80% glycerol in PBS for visualization with confocal microscopy. Unless otherwise indicated, confocal microscopy images were all collected at 60 x objective magnification with z-stacks acquired every 2 µm through the vertical span of the cells. Laser intensities were set to 2.5% and complete microscope parameters including HV, offset, and gain are presented in Appendix C. Zebrafish imaging Labeling experiments with zebrafish in vivo were conducted by Dr. Pui Ying Lam of the Peterson Lab at the University of Utah. Transgenic zebrafish embryos at three days post-fertilization expressing the fluorescent protein mCherry in macrophages were used. Images were collected using confocal microscopy and, in all cases, the PB-probe was used at a concentration of 18.2 µM. Three experimental conditions were tested, including direct incubation of the zebrafish embryo with the PB-probe solution, a 2 nL injection of the PB-probe into the yolk, and a 2 nL injection of the PB-probe into the circulatory system. Data analysis Maximum projections of the fluorescent signal from the PB-probe, PB-linker, or anti-rabbit IgG antibody conjugated to DyLightÒ 550 were overlaid on the transmitted detector (or white-light) channel to produce final images. The PHANTAST plugin for Fiji was used to segment images into cell area and background for analysis of fluorescent signal [87]. The 41 parameters utilized for cell segmentation are as follows: live/non-permeabilized cells, s = 1.2 and e = 0.02; permeabilized cells, s = 2 and e = 0.02. Mean fluorescence intensity of the backgrounds were subtracted from the cell area to yield absolute values for quantitative analyses. Colocalization analyses were conducted using the Coloc 2 plugin for Fiji V1.0. Maximum projections of the PB-probe and immunocytochemistry fluorescence channels were obtained as 16-bit images. Each field of view was analyzed independently using ten Costses randomisations and a point-spread function value of 1.0. Pearson correlation coefficients (PCC) and Mander’s colocalization coefficients (MCC) were reported. 5.3 Results and discussion RGD-mimetic compounds with radioactive tracers have previously been used to label integrins in vivo in the context of cancer [54, 69, 70]. Detectable through PET, these radioactive labels have yielded important findings in the field of oncology; however, they lack the spatial resolution to track integrins at the subcellular scale [124]. Fluorescent reporters enable more precise resolution than their radioactive counterparts: on the order of 250 nm with conventional microscopy techniques and 20 nm using super-resolution microscopy [125]. Consequently, a fluorescent chemical probe should provide a rapidly applicable method for the study of endogenous integrins with a resolution @ 10,000 x higher than existing PET techniques. Before applying the PB-probe to study the function of endogenous integrin though, it was necessary to fully characterize its labeling properties in vitro. 5.3.1 Optimization of labeling parameters The parameters for labeling endogenous integrin aVb5 with the PB-probe were optimized in vitro using BV-2 microglia. These model cells were chosen because integrin aVb5 represents @ 60% of the RGD-binding integrins expressed by this cell type [103-105]. Moreover, BV-2 cells have been shown to robustly model primary murine microglia, increasing the translatability of in vitro results presented herein to future studies in vivo [126]. First, the optimal concentration of PB-probe for labeling cell cultures in vitro was determined. The parent molecule of the PB-probe, cilengitide, is known to inhibit the binding of other substrates to integrin aVb5 with an IC50 of 11.7 ± 1.5 nM [78]. This concentration was determined by assays that used purified integrin protein. Here, 10 nM was chosen as the minimum PB-probe labeling concentration since it was expected that the addition of PB to cilengitide would not significantly impact its binding affinity towards integrin aVb5. Two 42 additional concentrations (100 nM and 1 µM) of the PB-probe were used initially to ensure that labeling would still be detected even if its binding affinity was reduced relative to cilengitide. A 3 h incubation at room temperature was chosen for integrin labeling to mimick secondary antibody labeling times used in immunocytochemistry [50]. Figure 5.1 indicates that the fluorescence intensity of cells labeled with PB-probe increases proportionally to PB-probe concentration. As expected, the highest fluorescence intensity of labeled cells was obtained following incubation with 1 µM PB-probe for 3 h (Fig. 5.1C and 5.1D). Interestingly, there was no significant difference in fluorescence intensity between treatment with 10 or 100 nM, yet both concentrations were statistically different from cells treated with 1 µM PB-probe. 43 Figure 5.1 Fluorescence intensity of labeled cells in vitro increases proportionally with PB-probe concentration Non-permeabilized, fixed BV-2 cells were incubated with A) 0.01 µM, B) 0.1 µM, or C) 1 µM PB-probe at room temperature for 3 h; representative images from three independent experiments are presented. After incubation, cells were washed twice with PBS and visualized using confocal microscopy. Absolute fluorescence intensity values of each treatment were quantified (lem 445-455 nm) from z-stacks representing three fields of view. Scale bar = 20 µm. D) Data from three independent experiments are presented as mean ± S.E.M. and were fit with a linear regression (significantly non-zero; P=0.0498, F=162.8). The effect of treatments was assessed by the one-way ANOVA, followed by Tukey’s multiple comparisons post-hoc test. ANOVA F and P values are presented on the graph. **P<0.01; significantly different from cells incubated with 0.01 µM PB-probe. ##P<0.01; significantly different from cells incubated with 0.1 µM PB-probe. Next, the time-dependence of PB-probe labeling in vitro was examined. The PB-probe was used to label BV-2 cells at a concentration of 1 µM with incubation times varying from 20 min to 9 h. 0.01 0.1 10510PB-Probe Concentration (µM)Fluorescence Intensity (AU)**##F = 78.69P <0.0001D 44 Figure 5.2 shows that the fluorescence intensity of cells labeled with the PB-probe increases proportionally with incubation time. The highest fluorescence intensity of labeled cells was obtained following a 9 h incubation with 1 µM PB-probe (Fig. 5.2D and 5.2E). This experiment revealed several important findings. First, incubation of the PB-probe with cell culture for 1 h is not significantly different from the 20 min incubation, indicating that a longer incubation time is necessary to allow for sufficient binding of the PB-probe. Second, Figure 5.2E illustrates that the relationship between fluorescence intensity and incubation time begins to plateau on the order of 9 h. This indicates that following an extended incubation period, changes in fluorescence intensity become less dependent on time. This is important for integrin aVb5 quantification because a large time-dependence may cause variation in fluorescence intensity that does not result from changes in integrin expression. However, the data also demonstrate that there is no significant difference between the fluorescence intensity of cells incubated with the PB-probe for 3 h and those incubated for 9 h. Consequently, a 3 h incubation time was used for subsequent experimentation to both take advantage of the rapid applicability of the PB-probe and minimize variation in fluorescence intensity resulting from incubation time. 45 Figure 5.2 Fluorescence intensity of labeled cells in vitro increases proportionally with PB-probe incubation time Non-permeabilized, fixed BV-2 cells were incubated with 1 µM PB-probe at room temperature for: A) 20 min, B) 1 h, C), 3 h, or D) 9 h. Representative images from three independent experiments are presented. After incubation, cells were washed twice with PBS and visualized using confocal microscopy. Absolute fluorescence intensity values of each treatment were quantified (lem 445-455 nm) from z-stacks representing three fields of view. Scale bar = 20 µm. E) Data from three independent experiments are presented as mean ± S.E.M. and were fit with a nonlinear regression. The effect of treatments was assessed by the one-way ANOVA, followed by Tukey’s multiple comparisons post-hoc test. ANOVA F and P values are presented on the graph. *P<0.05 and **P<0.01; significantly different from cells incubated with the PB-probe for 20 min. ##P<0.01; significantly different from cells incubated with the PB-probe for 1 h. 0 2 4 6 8 100510Incubation Time (h)Fluorescence Intensity (AU)F = 10.60P = 0.0037***##E 46 As a further remark on the data presented in Figure 5.2, highly localized regions of intense fluorescence are present in panel D that are absent from other panels representing shorter incubation times (Fig. 5.2A-C). To explain this phenomenon, one must consider the physiochemical properties of the PB-probe. As illustrated in Figure 5.3, the PB-probe possesses a polar chemical motif that contains two formal charges and another non-polar, aromatic component [127, 128]. The polarities of these motifs are quantified by the partition coefficients of each molecule in a solution of octanol and water (logP), quantified as an XlogP3-AA value [129]. XlogP3-AA is a predictive method of calculating logP values that uses an atom-additive model to increase its accuracy over other calculation methods [129]. Since the different regions of polarity on the PB-probe are separated by a covalent linker, the molecule as a whole is amphiphilic. Amphiphilic molecules, with detergents being a primary example, are able to form micelles in polar solutions such as the aqueous PBS buffer used for labeling in this experiment [130]. Moreover, these associations are stabilized at the interface of polar and non-polar regions (such as a plasma membrane) [131]. The highly localized regions of intense fluorescence present after a 9 h incubation with the PB-probe, but less so with shorter incubation times, may therefore be the result of micelle formation by the PB-probe. Such formations are enthalpically favored and may require a prolonged incubation time to form in great numbers [132]. It is therefore suggested that in practical applications of the PB-probe, incubation times greater than 3 h should be avoided when using the PB-probe at a concentration of 1 µM. However, since micelle formation is usually limited to solutions that are more concentrated than some threshold value, the application of lower PB-probe concentrations for longer than 3 h may both increase the quantitative accuracy of the PB-probe while avoiding nonspecific intermolecular associations [133]. Figure 5.3 The PB-probe is amphiphilic The chemical structure of the PB-probe is colored to indicate the polar cilengitide motif (red, XlogP3-AA = -1) and the non-polar PB motif (blue, XlogP3-AA = 0.9) [126, 127]. 47 A key finding from the above set of experiments is that the PB-probe effectively labels cells over the course of 3 h at a concentration of 1 µM. Compared to other integrin aVb5 labeling techniques, such as immunocytochemistry or transgene expression, application of the PB-probe is much more rapid. From first seeding cells to the completion of confocal microscopy image collection, the entire PB-probe-labeling workflow can be completed in approximately 6 h for one sample. In contrast, experiments with immunocytochemistry and transgene techniques commonly last at least two days [50, 134, 135]. This is in cases where antibodies or plasmids can be purchased from a supplier (sometimes at great expense), as their development in situ can frequently span weeks or months [136]. Moreover, these methods are limited in their scope of application as immunocytochemistry requires sample fixation prior to analysis and the expression of transgenes has been demonstrated to alter the subcellular localization of the recombinantly expressed proteins being investigated [61]. While the fluorescent labeling of endogenous protein in live cells for analysis through time is achievable with genetic modification, the PB-probe improves on this technique through both its ease-of-use and rapid applicability [61]. To further optimize the experimental parameters of the PB-probe, the effect of wash number on fluorescence intensity was investigated. It was hypothesized that washing the cells after incubation with the PB-probe would decrease fluorescence intensity by clearing free or non-specifically bound PB-probe. This trend was expected to continue until only specifically bound PB-probe remained, at which time the fluorescence intensity would remain unchanged with additional washes. This analysis was conducted simultaneously using fixed, non-permeabilized and permeabilized BV-2 microglia. Permeabilized cells were of interest for two primary reasons. First, permeabilization removes the plasma membrane of fixed cells. It was hypothesized that if the PB-probe is being non-specifically trapped inside the membrane of non-permeabilized cells, the signal returned from permeabilized cells would be weaker but more specific. Second, permeabilization is an important step in many immunocytochemistry procedures. Later in this chapter, antibody labeling techniques are applied simultaneously with the PB-probe. While the precise epitope of anti-integrin b5 is unknown because it is proprietary, the supplier indicated that the recognition motif of the antibody is cytoplasmic. Since antibodies are not permeable to the cell membrane, it was therefore necessary to permeabilize these cells for effective labeling and subsequent colocalization analysis. Prior to this analysis though, the effect of permeabilization on the efficacy of PB-probe labeling in vitro needed to be examined. BV-2 cells were fixed with formaldehyde and permeabilized or not with 0.1% v/v Triton X-100 in PBS prior to labeling. The PB-probe was added 48 at 1 µM for 3 h, then cells were washed 1-3 times with PBS and visualized using confocal microscopy. Representative images from the permeabilized and non-permeabilized cell treatments are presented in Figure 5.4A and 5.4B, while the wash data are quantified in Figure 5.4C. Figure 5.4 Fluorescence intensity of labeled cells in vitro changes with permeabilization but not wash number Fixed BV-2 cells were washed with PBS alone or permeabilized with 0.1% v/v Triton X-100 prior to incubation with 1 µM PB-probe at room temperature for 3 h. After incubation, cells were washed 1-3 with PBS and visualized using confocal microscopy. Representative images from three independent experiments are presented for A) non-permeabilized cells and B) permeabilized cells washed twice with PBS prior to visualization with confocal microscopy. Scale bar = 20 µm. C) Absolute fluorescence intensity values of each treatment were quantified (lem 445-455 nm) from z-stacks representing three fields of view. Data from three independent experiments are presented as mean ± S.E.M. The effect of treatments was assessed by the two-way ANOVA, followed by Tukey’s multiple comparisons post-hoc test. No significant differences were observed for different wash numbers within either cell treatment. ANOVA F and P values are presented on the graph. **P<0.01; significantly different from non-permeabilized cells incubated with 1 µM PB-probe and washed any number of times. Figure 5.4C demonstrates that there is no significant difference in the fluorescence intensity of cells washed 1-3 times with PBS after labeling with the PB-probe. A trend is seen, however, in for cells washed twice to have a lower fluorescence intensity than those washed once. Accordingly, two washes were used for subsequent experiments to minimize cell perturbation yet ensure that any unbound PB-probe was sufficiently cleared. Figure 5.4 also reveals that the fluorescence intensity of permeabilized cells labeled with the PB-probe is significantly less than those not permeabilized. Two possible explanations for this trend are offered. First, it is possible that the hypothesis outlined previously is correct and that 1 2 30510Wash NumberFluorescence Intensity (AU)Non-PermeabilizedPermeabilized**** **Wash TreatmentF = 5.065 F = 111.4P = 0.0254 P <0.0001C 49 the higher fluorescence intensity of non-permeabilized cells results from the PB-probe collecting non-specifically inside the cell membrane. This would suggest that while the signal returned from permeabilized cells is weaker, it is likely more specific. Alternatively, it may be that in both cases the PB-probe is labeling integrin aVb5 specifically but, as integrins are transmembrane proteins, the permeabilization step washes away some of the integrin aVb5 targets, thereby decreasing the fluorescent labeling. Intermolecular cross-linking associated with formaldehyde fixation should prevent integrin aVb5 from washing away in significant amounts, making the first explanation more likely [137]. However, the selectivity of the PB-probe in labeling permeabilized and non-permeabilized BV-2 cells was explored further through subsequent experimentation and will be discussed later. Despite the decreased fluorescence intensity associated with labeling permeabilized cells, it is important that the PB-probe is detectable using a preparation technique amenable to immunocytochemistry [50]. This is a prerequisite for the simultaneous application of immunocytochemistry techniques with the PB-probe, necessary to determine the degree that these methods colocalize. Moreover, it is proposed that the fluorescence intensity of permeabilized cells labeled with the PB-probe could be increased by using a longer incubation time or higher PB-probe concentration, however the effects of these changes have not been fully explored in permeabilized cells. More intense fluorescence could also be observed by increasing the detector sensitivity or laser power, although the reader is cautioned that laser illumination of live cells, especially at the relatively high-energy wavelength of 405 nm, is destructive to cells. In this set of experiments, the 405 nm laser was used at 2.5% power to prevent rapid cell death [122, 138]. Results from a separate experiment (Fig. 5.5) illustrate a population of BV-2 cells that were illuminated with a 455 nm laser at 5% power for 100 min. These cells exhibited changes in morphology that are consistent with the formation of apoptotic bodies. These changes were not evident in adjacent cells that were not illuminated throughout the duration of the experiment (data not shown). 50 Figure 5.5 Prolonged illumination of BV-2 cells effects morphology Live BV-2 cells were continuously irradiated over 100 min using the 455 nm laser at 5% power. Prior to visualization, cells were stimulated with 0.5 µg/mL LPS overnight. Latex beads were added 1:2000 in the cell culture media and cells were visualized immediately afterwards. Images from the same field of view at: A) t = 0 min and B) t = 100 min. Scale bar = 10 µm. Representative images from one independent experiment are presented. 5.3.2 Specificity of the PB-probe The specificity of the PB-probe towards RGD-binding integrins was investigated next. The highly potent inhibitor of integrin aVb3 and aVb5, cilengitide, was applied to cell cultures together with the probe at an equal concentration of 1 µM [71, 78]. It was hypothesized that simultaneous treatment with cilengitide would reduce the fluorescence intensity of labeled cells by competitively inhibiting PB-probe binding to integrin. Cells treated with cilengitide and the PB-probe had a lower mean fluorescence intensity than cells treated with PB-probe alone, but this effect was not statistically significant (Fig. 5.6). It is anticipated that with additional replicates, pre-incubation with cilengitide, or use of a higher cilengitide concentration, statistical significance would be achieved. However, it was not certain that such a finding would provide additional insight into the selectivity of the PB-probe. An alternative measure of PB-probe selectivity was therefore pursued. 51 Figure 5.6 Effect of cilengitide on the fluorescence intensity of cells labeled with the PB-probe Fixed, non-permeabilized BV-2 cells were incubated with 1 µM PB-probe in the presence or absence of 1 µM cilengitide at room temperature for 3 h. After incubation, cells were washed twice with PBS and visualized using confocal microscopy. Absolute fluorescence intensity values of each treatment were quantified from z-stacks representing three fields of view. Data from three independent experiments are presented as mean ± S.E.M. and were assessed with an unpaired, two-tailed t test. No significant differences were observed. The confocal microscope acquisition parameters of this experiment deviated from those outlined previously, with fluorescence emission detected from 420-540 nm. To further determine the specificity of the PB-probe in labeling integrin aVb5 in live, non-permeabilized, or permeabilized cells, its labeling efficacy was compared to the PB-linker. It was hypothesized that if the PB-probe is non-specifically accumulating in cells, then treatment with the PB-linker should result in a similar fluorescence intensity of labeled cells, independently of the cilengitide motif. Alternatively, if the PB-probe binds integrin aVb5 specifically, then incubation of cells with PB-linker should not result in any notable fluorescent signal. Both the PB-probe and PB-linker were incubated at a concentration of 1 µM for 3 h with BV-2 cells that were: live, fixed and non-permeabilized, or fixed and permeabilized. All samples were washed twice with PBS after the incubation time had elapsed and visualized using confocal microscopy. Representative images of the PB-probe-treated cells are presented in Figure 5.7A-C and PB-linker-treated cells are represented in Figures 5.7D-F with vertical columns indicating cell treatments. These data are quantified in Figure 5.7G. 0.1% DMSO1 µM Cilengitide0.00.51.01.52.02.5Fluorescence Intensity (AU) P = 0.05981 μM PB-Probe 52 Live cells Non-permeabilized cells Permeabilized cells 53 Figure 5.7 Fluorescent labeling is most intense in non-permeabilized cells treated with the PB-probe BV-2 cells were incubated with 1 µM PB-probe, 1 µM PB-linker, or 0.1% DMSO at room temperature for 3 h following fixation alone or with Triton X-100 permeabilization. Alternatively, live BV-2 cells were incubated with 1 µM PB-probe, 1 µM PB-linker, or 0.1% DMSO at 37 oC for 3 h in humidity. After incubation, cells were washed twice with PBS and visualized using confocal microscopy. A-F) Representative images from three independent experiments. Scale bar = 20 µm. G) Absolute fluorescence intensity values of each treatment were quantified (lem 445-455 nm) from z-stacks representing three fields of view. Data from three independent experiments are presented as mean ± S.E.M. The effect of treatments was assessed by the two-way ANOVA, followed by Tukey’s multiple comparisons post-hoc test. ANOVA F and P values are presented on the graph. **P<0.01; significantly different from cells incubated with 1 µM PB-probe that were non-permeabilized. #P<0.05 and #P<0.01; significantly different from cells of the same treatment incubated with 1 µM PB-probe. Since the PB-linker shares the same chemical structure as the PB-probe but lacks the cilengitide moiety, it was expected that the PB-linker would share any nonspecific interactions of the PB-probe caused by association of the fluorescent handle with off-target proteins. Figure 5.7 reveals that fixed, non-permeabilized cells treated with the PB-probe fluoresced almost three-fold more intensely than those treated with PB-linker. These data suggest that the majority of PB-probe labeling is both selective for RGD-binding integrins and results from a cilengitide-mediated directing effect. In no treatment did the fluorescence intensity of cells labeled with the PB-linker significantly differ from cells incubated with DMSO alone. This finding further supports that the cilengitide moiety is necessary for effective labeling of integrin with PB-probe. However, the fluorescent signal of permeabilized cells treated with PB-probe was also not significantly different from cells incubated with DMSO alone. Differences between live or permeabilized cells treated Live Non-PermeabilizedPermeabilized0510Cell TreatmentFluorescence Intensity (AU)0.1% DMSO1 µM PB-Linker1 µM PB-Probe****#####P = 0.0618P = 0.2205P = 0.8908Drug TreatmentF = 41.81 F = 13.72P <0.0001 P = 0.0002G 54 with the PB-probe and those treated with PB-linker may require a more sensitive detection technique to be fully resolved. An additional finding of Figure 5.7 is that the fluorescence intensity of live cells labeled with the PB-probe is less than non-permeabilized cells and comparable to permeabilized cells. Intriguingly, the subcellular localization of the PB-probe in live cells appeared to be different in Figure 5.7A than the labeling patterns observed in non-permeabilized and permeabilized cells, Figures 5.7B and 5.7C respectively. Live cells appeared to have very centrally located deposits of PB-probe while non-permeabilized and permeabilized cells displayed fluorescent signal that was excluded from the nucleus and more evenly distributed throughout the cytosol. While integrin aV and b5 subunits have been previously reported to localize throughout both the plasma membrane and the cytosol [50], the effect of cell treatment on PB-probe labeling was of interest. Therefore, it was investigated whether live cells might actively engulf the PB-probe, leading to subcellular localization unique from other cell treatments. It was hypothesized that if live cells are actively engulfing the PB-probe through phagocytic processes, then pre-treatment of the cells with an actin reorganization inhibitor, cytochalasin B, should result in a labeling pattern more closely resembling those observed in non-permeabilized and permeabilized cells by blocking endocytosis [139-142]. Figure 5.8 reveals that, contrary to this last hypothesis, live cells pre-treated with cytochalasin B and labeled with the PB-probe demonstrate a fluorescent labeling pattern very similar to live cells incubated in the absence of cytochalasin B. These data suggest that the PB-probe is not being actively engulfed by live cells through phagocytic processes. However, another interesting observation was that in all representative images of live cells labeled with the PB-probe (Fig. 5.7A, 5.7D, and 5.8), some cells present a spherical morphology. This morphology suggests the presence of cell stressors, such as those associated with confocal microscopy analysis at room temperature and altered pH levels resulting from atmospheric CO2 concentrations. Morphological changes are an initial step in apoptosis and may also be associated with integrin internalization [143, 144]. In order to monitor integrin internalization under these conditions, two independent methods to label integrin would be required that are distinct in fluorescence emission profiles and amenable to sequential application. At the time of writing, such methods were unavailable. While both the Alexa FluorÒ 568 probe and immunocytochemistry labeling methods may adequately serve in this capacity for future work, the fluorescence stability of the PB-probe following incubation with 4% formaldehyde has yet to be investigated. 55 Figure 5.8 Fluorescent labeling occurs following treatment with cytochalasin B Live BV-2 cells were treated with A) 0.1% DMSO or B) 2.5 µg/mL cytochalasin B for 15 min prior to incubation with 1 µM PB-probe at 37 oC for 3 h in humidity. Cells were washed twice with PBS and visualized using confocal microscopy. Representative image from three independent experiments. Scale bar = 20 µm. C) Absolute fluorescence intensity values of each treatment were quantified (lem 445-455 nm) from z-stacks representing three fields of view. Data from three independent experiments are presented as mean ± S.E.M. The effect of treatments was assessed by an unpaired, two-tailed t test. No significant differences were observed (P=0.9479). In a final investigation of the labeling selectivity of the PB-probe towards RGD-binding integrins, the subcellular colocalization of the PB-probe with a concomitant immunostain was explored. Immunocytochemistry is a widely accepted and robust method for the detection of cellular proteins, although it is not always reliably selective [65]. An integrin b5-targeting antibody was employed for this investigation as no commercial antibodies were available that target the murine integrin aVb5 heterodimer. Integrin b5 was chosen as a molecular target instead of integrin aV because, while it is known to only form obligate heterodimers with integrin aV, the aV subunit is promiscuous in its associations [145]. Therefore, immunodetection of integrin b5 was expected to provide a more accurate representation of integrin aVb5 localization. Standard immunocytochemistry labeling procedures were followed, with the modification that after the wash steps succeeding primary antibody incubation were complete, the PB-probe was added to the cell culture in blocking buffer (4% CBS in PBS) for 1.5 h prior to the addition of secondary antibody. The fluorescence emission of the probe and the DyLightÒ 550 fluorophore of the secondary antibody were detected at their respective emission maxima (459 nm and 576 nm). It was hypothesized that the labeling of one technique would be indicated 0.1% DMSO2.5 µg/mLCytochalasin B0.00.51.01.52.02.5Fluorescence Intensity (AU) 1 μM PB-ProbeC 56 by PCC and MCC analyses to entail the other. Representative images from this experiment are presented in Figure 5.9, with the coefficient analyses results outlined in Table 5.1. Figure 5.9 Immunocytochemistry and the PB-probe label cells simultaneously BV-2 cells were incubated in the A) presence or B) absence of anti-integrin b5 primary antibody overnight prior to labeling with 1 µM PB-probe and anti-rabbit IgG antibody conjugated to DyLightÒ 550. Cells were washed twice with PBS and visualized using confocal microscopy. Representative images from three independent experiments are presented. Scale bar = 30 µm. Table 5.1 Integrin labeling by the PB-probe and immunocytochemistry techniques share 59% colocalization Data from three independent experiments are presented. Pearson correlation coefficient values range from 1 to -1, indicating perfect correlation and perfect inverse correlation of signals, respectively. Mander’s colocalization coefficients offer a similar measure but range from 0 to 1 and express the fraction of signal from one channel that is accompanied by the other. PCC MCC PB-Probe 0.18 ± 0.02 0.52 ± 0.01 Immunocytochemistry / 0.16 ± 0.03 Importantly, Figure 5.9 demonstrates that the labeling parameters of the PB-probe are compatible with commonly employed immunocytochemistry labeling techniques, confirming the PB-probe as an easily applicable labeling strategy amenable to simultaneous immunodetection [50]. This is important for future investigations of integrin aVb5 localization and function with the PB-probe, as these studies will likely benefit from concomitant detection of associated 57 signaling molecules. For example, intracellular colocalization with talin or FAK could suggest that integrin aVb5 is functioning in ECM anchorage processes or phagocytosis, respectively. Images collected from these experiments were analyzed to quantify the colocalization of the PB-probe and immunocytochemistry fluorescent signals. Perfect overlap of these labeling techniques would indicate accurate and precise targeting of integrin aVb5. Two common measures of colocalization are the PCC and the MCC [146]. In this experiment, it was hypothesized that both the PCC and MCC values would indicate a high correlation of fluorescence labeling by the PB-probe and immunocytochemistry. Data from the colocalization analyses presented in Table 5.1 indicate that the PB-probe and immunocytochemistry techniques share 59 ± 1% colocalization as assessed by the PCC (0.18 ± 0.02). This value corresponds closely to the 65% specific labeling observed in Figure 5.7. Generally, PCC values >0.5 are taken to indicate colocalization while values <0 suggest random overlap [147]. Notably, the PCC value reported for the PB-probe (0.18 ± 0.02) exists between these classifications. As small-molecule labeling techniques are still an underdeveloped technique for the detection and localization of protein targets, very few literature precedents are available for comparison regarding labeling selectivity. Other reports have assessed labeling specificity in the absence of quantitative data [148], characterized colocalization with entire organelles [149], or investigated specific off-target protein interactions [150]. Of these, the most informative studies report the binding activity of labeling probes towards on-target and off-target proteins [150]. Consistent with our proposed explanation that the PB-probe labels other RGD-binding integrins while immunocytochemistry only labels integrin aVb5, off-target interactions are typically associated with proteins that bind similar chemical motifs (such as endocannabinoids in the case of [150]). The MCC value (0.52 ± 0.01) of fluorescent signal above an intensity threshold indicates that 52 ± 1% of immunocytochemistry signal is accompanied by PB-probe labeling. This coefficient is reported for above-threshold values only because the raw data returned an MCC value of 1 ± 0. This suggests that the confocal microscope photodetector may have been too sensitive for the fluorescence detection of the PB-probe. The most likely explanation for 100% overlap of PB-probe labeling with immunocytochemistry (± 0) is that light scattering was recorded as weak fluorescent signal for all pixels. Intensity thresholds correct for this background signal. The MCC for above-threshold PB-probe signal accompanied by immunocytochemistry was 0.16 ± 0.03. This value is significantly lower than the MCC of the inverse relationship. A possible explanation is that the PB-probe labels other RGD-binding integrins while immunocytochemistry only labels integrin aVb5. Integrin aVb3, aVb1, and aVb8 are also 58 expressed by murine microglia, representing @ 40% of surface aV integrins on this cell type [103-105]. As was found previously, the MCC was much higher when calculated from the raw data (0.94 ± 0.02); however, this value may be artificially inflated by background signal as well. Future investigations will be necessary to determine whether the low MCC for PB-probe labeling accompanied by immunocytochemistry is due to associations of the PB-probe with other RGD-binding integrins or attributable to nonspecific interactions. 5.3.3 Application of the PB-probe in vivo Finally, the labeling profile of the PB-probe was investigated in vivo through a collaborative effort with Dr. Pui Ying Lam of the University of Utah. In this experiment, zebrafish embryos (Fig. 5.10) expressing mCherry in macrophages were employed due to their utility as a model organism for neurodegenerative disease and ease of use (rapid doubling time, translucent tissue) [151]. This section presents preliminary results as studies were still ongoing at the time of writing. Figure 5.10 Zebrafish embryo 72 h post-fertilization Protruding mouth stage of embryogenesis. reproduced from Kimmel et al. [152]. Scale bar = 250 µM. Three different application methods were employed to test the capacity of the PB-probe to label tissue in vivo. First, a zebrafish embryo three days post-fertilization was incubated directly with an 18.2 µM solution of the PB-probe for 3 h and images were collected using confocal microscopy. Representative images of the PB-probe-treated embryo are shown in Figure 5.11 with vertical columns indicating the fluorescence detection channel and rows indicating different fields of view. Figure 5.11 illustrates that the PB-probe labels tissue structures in this application but does not appear to colocalize with fluorescent signal from macrophages. 59 Figure 5.11 The PB-probe does not label macrophages in zebrafish embryos following direct incubation Zebrafish embryos expressing mCherry in macrophages were incubated three days post fertilization in an 18.2 µM PB-probe solution for 3 h. Fluorescent signal at 450 nm (blue) is only found in tissues – none above background is observed in macrophage cells (red). Samples were visualized using confocal microscopy. Data from one independent experiment were collected by Dr. Pui Ying Lam of the Peterson Lab, University of Utah. Second, 2 nL of an 18.2 µM PB-probe solution were injected directly into the yolk of a zebrafish three days post-fertilization and incubated for 3 h prior to visualization with confocal microscopy. Representative images of the yolk-injected embryo are presented in Figure 5.12 with vertical columns indicating the fluorescence detection channel. Figure 5.12 demonstrates a higher overlap of PB-probe fluorescence with macrophage signal than that observed following the direct-incubation method. This field of view also presents a much higher number of macrophages. Since these images were collected proximal to the yolk-injection site, it is likely that macrophages were recruited by chemoattractant signals released in response to tissue damage. Interestingly, Dr. Lam remarked that recruited macrophages appeared to be engulfing the PB-probe; however, it is unclear whether this process was mediated by integrins or not. An additional finding of Figure 5.12 is that the injection of an 18.2 µM solution of PB-probe in the A B C D E F PB-Probe mCherry Merge 60 relatively confined yolk seems to have resulted in highly localized regions of intense fluorescence resembling the micelle formation described previously in Figure 5.2. Figure 5.12 The PB-probe colocalizes with macrophages in zebrafish embryos following injection into the yolk Zebrafish embryos expressing mCherry in macrophages were injected three days post fertilization with 2 nL of 18.2 µM PB-probe solution into the yolk. Fluorescent signal at 450 nm (blue) is observed in tissues surrounding the injection site and is engulfed by macrophages (red). Samples were visualized 3 h post-labeling using confocal microscopy. Data from one independent experiment were collected by Dr. Pui Ying Lam of the Peterson Lab, University of Utah. Finally, 2 nL of an 18.2 µM PB-probe solution were injected directly into the circulation of a zebrafish embryo three days post-fertilization and incubated for 3 h prior to visualization with confocal microscopy. Representative images are presented in Figure 5.13 with vertical columns indicating the fluorescence detection channel and rows indicating different fields of view. The images in Figure 5.13 illustrate similar trends to those observed in both Figures 5.11 and 5.12. It can be seen once again that PB-probe labeling does not colocalize with the fluorescent signal of macrophages and, as with the previous injection method, highly localized regions of intense fluorescent signal are observed. PB-Probe mCherry Merge A B C 61 Figure 5.13 The PB-probe does not label macrophages in zebrafish embryos following injection into circulation Zebrafish embryos expressing mCherry in macrophages were injected three days post fertilization with 2 nL of 18.2 µM PB-probe solution into their circulation. Samples were visualized 3 h post-labeling using confocal microscopy. Data from one independent experiment were collected by Dr. Pui Ying Lam of the Peterson Lab, University of Utah. The zebrafish labeling experiments conducted by Dr. Lam yielded several insights into the labeling characteristics of the PB-probe in vivo. Firstly, 18.2 µM of PB-probe appeared to be sufficient for labeling cellular structures through a direct incubation method but may be too concentrated for more localized applications achieved through injection. Secondly, there does not seem to be any specific macrophage labeling. To the contrary, there appears to be no macrophage labeling at all. It has been previously reported that macrophages have low integrin aVb5 expression; however, these findings are not specific to zebrafish [153, 154]. If there is low expression of integrin aVb5 by zebrafish macrophages three days post-fertilization, it may be that PB-probe labeling is specific, simply not directed at macrophages. Together, the preliminary results of this ongoing study represent an exciting first application of the PB-probe in vivo, demonstrating that it is able to label cellular structures discriminately and through a variety of application methods. A B C D E F PB-Probe mCherry Merge 62 5.4 Summary The above data confirm that the PB-probe is a robust and rapidly applicable labeling method for the fluorescence detection of endogenous RGD-binding integrins. The fluorescence intensity of labeled cells is dependent on both the incubation time and concentration of the PB-probe employed in the study. The optimal parameters for labeling experiments utilizing the PB-probe were determined to be a 3 h incubation with 1 µM PB-probe, followed by two washes with PBS and subsequent visualization. These parameters improve significantly on the timescale required by other techniques for protein labeling, such as immunocytochemistry and transgene expression [134, 135]. Significantly, the PB-probe has been demonstrated to label cells that are live, fixed but not permeabilized, and fixed and permeabilized. These findings indicate that the PB-probe is compatible with immunocytochemistry techniques and these methods have indeed been shown to be suitable for simultaneous application. Moreover, labeling with the PB-probe and immunocytochemistry techniques simultaneously demonstrate modest colocalization, suggesting the detection of a shared molecular target. The selectivity of the PB-probe is further supported by the finding that cells are labeled much more effectively by the PB-probe compared to the PB-linker, indicating that cilengitide has a directing effect towards RGD-binding integrins. Lastly, labeling with the PB-probe was observed in vivo in zebrafish embryos three days post-fertilization. 63 6 The integrin labeling probes retain inhibitory effect on integrin aVb5 signaling in vitro 6.1 Outline This chapter demonstrates that stimulated BV-2 microglial cells are an effective model system for the investigation of cellular signaling pathways engaged in phagocytosis. Analyses with confocal fluorescence microscopy and flow cytometry reveal that vitronectin-coated latex bead engulfment is integrin-dependent. Phagocytic activity towards vitronectin-coated beads is blocked by the integrin probes, confirming that inhibitory effect towards integrin-mediated signaling is retained, relative to cilengitide. 6.2 Experimental methods Reagents Carboxylate-modified polystyrene latex beads 1 µm in diameter, fluorescing yellow-green (lex @ 470 nm, lem @ 505 nm), were purchased from Sigma Aldrich (#L4655). 2-(-4-morpholino)-ethane sulfonic acid (MES) buffer was prepared as a 50 mM stock by dissolving MES free acid monohydrate (Fisher Scientific #BP300-100) in deionized water. Fluorescent beads were coated with vitronectin (Sigma Aldrich #SRP3186-250UG) through passive adsorption, as outlined according to the supplier. Briefly, 1 mg/mL vitronectin was diluted 2 x with MES buffer (final concentration 15 mM) and bead suspension, resulting in a 0.5% solids solution. The vitronectin-bead adsorption suspension was gently rocked overnight at 4 oC then centrifuged at 12,000 xg for 15 min. After centrifugation, the supernatant was aspirated and coated beads were resuspended to their original volume with PBS immediately before use. Cell imaging BV-2 cells were prepared for analysis by confocal microscopy by suspending cells at 0.2 million cells/mL and seeding 30 µL onto pre-sterilized square, 18 mm microscope cover glass 0.13 – 0.17 mm thick (Fisher Scientific #12-542A) in a 2 cm dish (Sarstedt Inc. #83.3900, Montreal, QC, Canada) for a 2 h incubation at 37 oC in humidity and 5% CO2. F10 was then added to the cell culture to bring the total volume of media up to 2 mL. Cells were incubated overnight before the media was aspirated and replenished with fresh F10 30 min prior to stimulation with LPS (0.5 µg/mL, Sigma Aldrich #L6529-1MG). BV-2 cells were subsequently incubated for 24 h to induce a pro-inflammatory, M1 phenotype. Next, BV-2 cell culture media was aspirated to leave 1 mL remaining. Cells were treated with 10 nM of cilengitide, PB-probe, or vehicle solution (0.1% DMSO) for 15 min prior to addition 64 of 10 µL of vitronectin-coated bead solution and incubated for 2.5 h at 37 oC in humidity and 5% CO2. BV-2 cells were then washed once with PBS and treated with 200 µL 0.25% trypsin with 0.2 g/L EDTA (Fisher Scientific #SH3004202) for an ensuing five-minute incubation to remove most of the extracellular beads [155-157]. The dish was then removed from the incubator, tapped lightly on the side to loosen adherent cells, and visually inspected to ensure no cells remained attached to the dish. Pre-warmed F10 medium (2 mL) was added to the dish and washed over the growth surface to inactivate the remaining trypsin and further loosen adherent cells. Following gentle mixing, the cell suspension was removed and transferred to a new 15 mL tube (VWR) for centrifugation at 500 xg for 7 min. The cell pellet was resuspended in 50 µL of F10 and seeded on a pre-sterilized microscope cover glass for a 2 h incubation at 37 oC in humidity and 5% CO2 to allow for adherence of cells. After 2 hours had elapsed, BV-2 cells were washed three times with PBS and fixed with 2 mL 4% formaldehyde (Fisher Scientific) diluted in F10 for 15 min at room temperature. Coverslips were then mounted on rectangular, 25 x 75 mm microscope slides 1 mm thick (Fisher Scientific #12-550-A3) with 80% glycerol (Sigma Aldrich) in PBS and visualized using confocal microscopy. Methods for cell fixation and slide preparation were conducted in accordance to commonly employed techniques for fluorescence microscopy sample preparation [50]. Flow cytometry For analysis with flow cytometry, the protocols described above were followed with the following modifications. BV-2 cells were prepared by suspending cells at 0.25 million cells/mL and seeding 300 µL into 2 cm cell culture dishes. The cell culture media was immediately made up to 2 mL with F10 and incubated overnight, then stimulated with LPS as described above. Following incubation with LPS, BV-2 cells were treated with 5 nM – 50 µM of inhibitor or vehicle solution (0.05% DMSO) for 15 min prior to addition of 10 µL of vitronectin-coated bead solution, then incubated for 2.5 h at 37 oC in humidity and 5% CO2. Next, BV-2 cells were washed three times with PBS and resuspended in F10 for transport to the flow cytometer. BV-2 cells were then washed once more with PBS and treated with 200 µL 0.25% trypsin with 0.2 g/L EDTA for five minutes. Once all cells had dislodged, pre-warmed F10 medium (300 µL) was added to the dish and washed over the growth surface to inactivate the remaining trypsin. Following gentle mixing, the cell suspension was removed and transferred to a new 5 mL tube (Miltenyi Biotec Inc. San Diego, CA, USA) for centrifugation at 500 xg for 7 min. The cell pellet was resuspended in 500 µL of PBS and 1 µL of propidium iodide (PI, Miltenyi Biotec Inc. #130-093-233) was added to each cell suspension immediately before analysis. Flow cytometry analyses were conducted using a MACSQuant Analyzer and MACSQuantify Software V2.11 (Miltenyi Biotec Inc.). 65 The flow cytometer was configured prior to experimentation to include several gating parameters that filtered out data from cells that had clumped together or were dead at the time of analysis, as well as any cellular debris or free-floating beads. Data from cell aggregates had to be filtered out because they are recorded as a single ‘event’ and return fluorescence values that are artificially high. These values were excluded from analysis using a gating parameter set to compare two measures of cell size, recorded by the instrument as the area and height of light intensity side-scatter (SSC), versus cell position relative to the laser/detector. For a single cell, the ratio between SSC area and height follows an approximately linear relationship; clumped cells deviate from this trend [158]. Next, cells that were dead at the time of recording were eliminated from the dataset. Dead cells do not actively engulf vitronectin-coated beads, leading to a lower fluorescence intensity resulting from decreased cell viability rather than an effect of inhibitors on integrin signaling. The fluorescent marker PI was employed to distinguish between living and dead cells. Propidium iodide permeates cells with a compromised plasma membrane, indicating loss of viability [159]. Lastly, flow cytometry data were filtered once more based on the forward-scatter (FSC) size parameter to eliminate cellular debris or free-floating beads from the final analysis. Median fluorescence intensities were calculated from cells above an experimentally-determined threshold and the percent of cells that were actively engulfing beads was quantified based on the number of cells above and below this threshold. Where possible, flow cytofluorometry data are presented as absolute median fluorescence intensity values, as reported by the flow cytometer. Some experiments include replicates that were acquired with fluorescent beads from different lot numbers. These data are presented as relative median fluorescence intensity values to account for any variation resulting from differences in the fluorescence intensity of beads from different lots. Cell viability In order to confirm that reduced phagocytic activity of BV-2 cells following treatment with integrin inhibitors was not due to cell death, cell viability was measured using 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT). This water-soluble tetrozolium compound is converted to a formazan dye by the mitochondrial enzyme succinate dehydrogenase in live cells, precipitating as purple crystals. Colorimetric quantification of formazan dye allows for the proportional determination of cellular viability. Briefly, 5 µL of a 10 mg/mL MTT in deionized water were added to the cell culture medium of each well in a 96-well plate (Greiner Bio-One Inc. #655061, Monroe, NC, USA) reaching a final MTT concentration of 1 mg/mL [160]. Plates were incubated for 1 hour at 37 oC in humidity and 5% CO2 to allow for cellular metabolism of the tetrozolium dye. Next, a 20% sodium dodecyl sulfate (SDS)/50% N,N-dimethylformamide (DMF) solution in milliQ water was added 1:1 with the cell culture volume in 66 each well to lyse the cell membranes and solubilize the formazan crystals. Plates were then incubated overnight at 37 oC in humidity until the solution in each well was homogenous. The absorbance of each well at 570 nm was subsequently measured and cell viability was calculated as a percent relative to fully viable cells incubated in cell media only. These data were first standardized against cell culture media devoid of cells, indicating no enzymatic activity, according to the following equation: Percent cell viability = 100(Atreatment well – Amedia)/(Afully viable well – Amedia) [6.1] It is important to note that the absorbance of the Alexa FluorÒ 568 probe overlaps closely with the 570 nm detection wavelength used to measure cell viability. In this case, the absorbance at 570 nm of the Alexa FluorÒ 568 probe solutions in cell media, corresponding in concentration to the treatment conditions, were subtracted from the treatment wells prior to analysis. 6.3 Results and discussion Fluorescent beads coated with ECM protein are readily engulfed by phagocytic cells in vitro [161]. ECM proteins have been shown to engage select transmembrane receptors and facilitate bead endocytosis through specific interactions [162, 163]. Since vitronectin is recognized by integrin aVb5, the relative efficacy of cilengitide and the integrin probes in blocking vitronectin-coated fluorescent bead engulfment was evaluated to determine whether inhibitory effect towards integrin is retained, relative to cilengitide [164]. 6.3.1 Validation of BV-2 cells as a model system to evaluate phagocytic activity To determine whether pre-treatment of cells with cilengitide or the integrin probes block the cellular uptake of fluorescent beads, it was first necessary to develop a robustly phagocytic model system in vitro. BV-2 cells were chosen for use in this study due to their demonstrated similarity to integrin aVb5-expressing primary microglia [103, 126]. Since previous reports have indicated that pro-inflammatory, M1 phenotype microglia exhibit more robust phagocytic activity than unstimulated cells, the effect of LPS-stimulation on BV-2 microglial phagocytosis was investigated [165-167]. BV-2 cells were incubated in the presence or absence of 0.5 µg/mL LPS then treated with fluorescent latex beads 1 µm in diameter. This bead diameter was selected to ensure it was large enough for easy visual detection through microscopy and small enough to initiate phagocytic 67 cascades, as opposed to cell-spreading events associated with larger bead diameters [168]. Bead phagocytosis was quantified using confocal microscopy. Representative images collected from each cell treatment and mean values of bead engulfment are presented in Figure 6.1. While these data represent a single experiment, they demonstrate a trend that M1 microglia have an enhanced phagocytic response relative to unstimulated control cells. This finding is consistent with literature reports that activated microglia are more phagocytic than those in a resting, or ramified, phenotypic state [25]. Figure 6.1 Effect of LPS-stimulation on latex bead engulfment by BV-2 microglia BV-2 microglia were treated with: A) vehicle (10 µL PBS) or B) LPS (0.5 µg/mL) for 24 h then incubated with latex beads (0.5% solid suspension diluted 100 x into cell culture dish) for 2.5 h. After washing, cells were analyzed with confocal microscopy. C) The mean number of fluorescent beads (± S.E.M.) engulfed per cell are presented from one independent experiment. Scale bar = 20 µm. The effect of integrin inhibition on the engulfment of vitronectin-coated beads was subsequently determined. Since vitronectin is the native ligand for integrin aVb5, it was hypothesized that the engulfment of vitronectin-coated beads would be blocked by integrin inhibition [164, 169, 170]. Unstimulated BV-2 cells were pre-treated with 10 nM cilengitide or vehicle solution (0.1% DMSO) then incubated with vitronectin-coated beads for 2.5 h before analysis with confocal microscopy. Representative images from these cell treatments and mean values of bead engulfment are presented in Figure 6.2. The concentration of cilengitide for this initial study was chosen to approximate the IC50 of cilengitide towards integrin aVb5 ligand-binding (11.7 ± 1.5 nM) [78]. While Figure 6.2 again represents a single experiment, the data indicate that Unstimulated CellsLPS-Stimulated Cells050100150200250Beads/Celln = 125n = 119C 68 vitronectin-coated bead engulfment by unstimulated microglial cells is very similar following treatment with cilengitide or vehicle. This finding suggests that there is no effect of integrin aVb5-blockade on vitronectin-coated bead engulfment in unstimulated cells. This is consistent with literature reports that integrin aVb5-signaling requires prior activation by PKC for recruitment to downstream signaling partners [171, 172]. Several studies have implicated LPS as a stimulus of PKC; however, the engagement of this pathway in integrin aVb5-mediated signaling was not explicitly studied herein [173, 174]. Figure 6.2 Effect of cilengitide on vitronectin-coated bead engulfment by unstimulated BV-2 microglia Unstimulated BV-2 microglia were treated with: A) 0.1% DMSO or B) 10 nM cilengitide then incubated with vitronectin-coated beads (0.5% solid suspension diluted 100 x into cell culture dish) for 2.5 h. After washing, cells were analyzed with confocal microscopy. C) The mean number of fluorescent beads (± S.E.M.) engulfed per cell are presented from one independent experiment. Scale bar = 20 µm. 6.3.2 Effects of integrin aVb5 inhibitors on bead engulfment (confocal microscopy) In accordance with these results, LPS-stimulated BV-2 cells were used for subsequent experimentation to determine the effect of pharmacological blockade of integrin aVb5 on vitronectin-coated bead engulfment. As an inhibitor of RGD-binding integrins, including integrin aVb5, it was hypothesized that pre-treatment of cells with cilengitide would block the cellular uptake of vitronectin-coated beads [71, 78]. To test the efficacy of cilengitide in blocking phagocytosis initiated by integrin aVb5, BV-2 cells were pre-treated with 10 nM cilengitide or 0.1% DMSO10 nM Cilengitide020406080Beads/Celln = 134n = 100C 69 vehicle solution (0.1% DMSO) then incubated with vitronectin-coated beads for 2.5 h before analysis with confocal microscopy. Figure 6.3 illustrates representative images from confocal microscopy analyses of vehicle-treated cells compared to cells treated with cilengitide. The data demonstrate a clear decrease in the number of fluorescent beads internalized following treatment with cilengitide, from a median value of 59 beads engulfed by vehicle-treated cells to 29 beads per cell following treatment with cilengitide (51% decrease). Figure 6.3 Cilengitide blocks the uptake of vitronectin-coated beads by stimulated BV-2 microglia LPS-stimulated BV-2 microglia were treated with A) 0.1% DMSO or B) 10 nM cilengitide then incubated with vitronectin-coated beads (0.5% solid suspension diluted 100 x into cell culture dish) for 2.5 h. After washing, cells were analyzed with confocal microscopy. C) The median number of engulfed beads per cell from 4 independent experiments are presented as mean ± S.E.M. The effects of the treatments were assessed using an unpaired, two-tailed t test. *P<0.05, significantly different from control cells incubated in the absence of drugs. Scale bar = 20 µm. It has been reported that variation at the meta position of cilengitide’s phenyl ring is well-tolerated in terms of binding affinity to integrin aVb5 [77]. Since this is where the PB-linker of the PB-probe is attached, it was hypothesized that the PB-probe would be equipotent, relative to cilengitide, in blocking signals transduced by integrin aVb5. To test his hypothesis, BV-2 cells were pre-treated with 10 nM PB-probe or vehicle solution (0.1% DMSO) then incubated with vitronectin-coated beads for 2.5 h before analysis with confocal microscopy. Representative images from confocal microscopy analyses of vehicle-treated cells compared to those treated with PB-probe are presented in Figure 6.4. Similar to cilengitide, a decrease in the number of beads engulfed by cell treated with the PB-probe is observed, although this effect was not 0.1% DMSO10 nMCilengitide020406080100Beads/Cell*C 70 statistically significant. On average, vehicle-treated cells engulfed a median value of 44 beads engulfed per cell while PB-probe-treated cells engulfed 21 (52% decrease). Figure 6.4 The PB-probe blocks the uptake of vitronectin-coated beads by stimulated BV-2 microglia LPS-stimulated BV-2 microglia were treated with A) 0.1% DMSO or B) 10 nM PB-probe then incubated with vitronectin-coated beads (0.5% solid suspension diluted 100 x into cell culture dish) for 2.5 h. After washing, cells were analyzed with confocal microscopy. Scale bar = 20 µm. C) The median number of engulfed beads per cell from 4 independent experiments are presented as mean ± S.E.M. The effects of the treatments were assessed using an unpaired, two-tailed t test. No significant differences from control cells incubated in the absence of PB-probe were observed. Figures 6.3 and 6.4 illustrate that BV-2 cells pre-treated with 10 nM cilengitide or PB-probe engulfed, on average, less beads relative to cells treated with vehicle alone, although this effect was only approaching significance in the case of the PB-probe. It is anticipated that with additional replicates, the trend for cells to engulf fewer beads after pre-treatment with the PB-probe would become statistically significant. However, due to the high interexperimental variation and the time-intensive data analysis, an alternative method of analysis (flow cytometry) was pursued and will be discussed later. In order to ensure that the decrease in bead engulfment associated with the various treatments employed in the above experiments was due to effects on signaling cascades, and not due to cell death, an MTT assay was performed. The MTT assay is an indirect but highly sensitive measurement of cell viability, indicating the reductive capacity of mitochondria through the active cellular conversion of the tetrazolium dye to an insoluble purple formazan [160]. Figure 0.1% DMSO10 nMPB-Probe020406080100Beads/CellP = 0.1153C 71 6.5 indicates that cilengitide and the PB-probe do not affect the viability of BV-2 cells at concentrations up to 100 µM over the course of 3 h. This incubation time was chosen to reflect the drug treatment period used in the vitronectin-coated bead engulfment assay. Figure 6.5 Cilengitide and the PB-probe are not cytotoxic BV-2 cells were treated with cilengitide or PB-probe (10 nM - 100 µM) then incubated for 3 h. Cell viability was measured using the MTT assay. Data from 6 independent experiments were calculated using equation [6.1] and are presented as mean ± S.E.M. The effects of cilengitide and PB-probe were assessed by the two-way ANOVA, followed by Dunnett’s multiple comparisons post-hoc test. ANOVA F and P values are presented on the graph. No significant differences from control cells incubated in the absence of drugs were observed. To further characterize the effects of cilengitide or the PB-probe on microglial bead engulfment, a histogram was created to compare the distribution of internalized beads in each cell population, relative to control cells incubated in the absence of drugs. Figure 6.6 shows that the effects of integrin inhibitors on vitronectin-coated bead engulfment were not evenly distributed throughout cell populations. Specifically, fewer cells engulfed more than 50 beads per cell when treated with cilengitide or the PB-probe compared to the control, although this effect was only statistically significant in select comparisons. Interestingly, more cells treated with cilengitide or the PB-probe engulfed 10-30 beads compared to those treated with the vehicle. These data suggest that cilengitide and the PB-probe are most efficacious at blocking high amounts of phagocytosis; however, the reasons underlying this phenomenon are unknown. Three potential explanations for this trend are offered. First, it may be that cells are able to engulf a certain number of beads non-specifically and that integrin aVb5 signaling is required for phagocytosis of >50 beads per cell. This explanation is supported by the data in Figure 6.1, showing that uncoated beads are internalized, but does not account for the trend in Figure 6.6 that more cells treated with cilengitide or the PB-probe engulf 10-30 beads per cell compared to the control. Second, it could be that integrin aVb5-blockade slows the process of vitronectin-0 0.01 0.1 1 10 100050100Concentration (µM)Cell Viability (%)Cilengitide PB-ProbeDrug ConcentrationF = 3.68 F = 3.755P = 0.606 P = 0.0056 72 coated bead engulfment. This explanation accounts for the trends illustrated in Figure 6.6 and is suggested to be the most likely. Lastly, it is possible that cilengitide and the PB-probe affect the inflammatory response of stimulated microglia and reduce phagocytic response by restoring a ramified, M2, phenotype. This explanation is admittedly unlikely, as anti-inflammatory effects of cilengitide are undocumented and both compounds were added 15 min prior to the addition of vitronectin-coated beads, which is a timescale more closely associated with pharmacological blockade than a cellular response at the genetic level. To a further point, no more than 7% of cells from any treatment did not engulf any beads. This suggests that it was not the phagocytic response of microglia that was reduced upon treatment with cilengitide or the PB-probe but rather, a sustained phagocytic response that was less efficacious or slowed, relative to the control. Figure 6.6 The effect of cilengitide and the PB-probe on bead engulfment is amplified at high levels of phagocytosis Relative frequency distribution of engulfed beads per cell. Bin width = 20 beads/cell. LPS-stimulated BV-2 microglia were treated with DMSO (0.1% v/v) or cilengitide or PB-probe (10 nM) then incubated with vitronectin-coated beads (0.5% solid suspension diluted 100 x into cell culture dish) for 2.5 h. After washing, cells were analyzed with confocal microscopy and engulfed beads of at least 50 cells per treatment were counted manually. Data from 7 independent experiments are presented as mean ± S.E.M. The effects of cilengitide and PB-probe were assessed by the two-way ANOVA, followed by Tukey’s multiple comparisons post-hoc test. Note that statistical differences are only illustrated for select comparisons. ANOVA F and P values are presented on the graph. #P<0.05, significantly different from control cells incubated in the absence of drugs from a different bin. Returning to the interexperimental variation mentioned above and the time-intensive data analysis associated with confocal microscopy analysis, it should be noted that several factors can affect the quality and quantity of data generated using this technique. Firstly, analysis with confocal microscopy was inherently time consuming and did not offer any significant benefits 0 20 40 60 80 10012014016018020022024026028030032001020304050Bin Center (Beads/Cell)Relative Frequency (%) 0.1% DMSO10 nM Cilengitide10 nM PB-Probe##nsDrugF = 0P >0.9999Beads/CellF = 67.19P <0.0001 73 over other image collection methods, such as epifluorescence microscopy. This was exacerbated by the time it took to count the number of beads in each individual cell for each cell treatment. Secondly, only some fields of view were suitable for manual bead counting; specifically, where individual cells could be distinguished. Because some fields of view were excluded from the final analysis, it is possible that analyzed cells did not represent the entire population. Next, a notable limitation of the confocal microscopy analysis is that, in this experimental design, images and data were not collected or analyzed in a blinded fashion, potentially introducing bias. Consequently, further analyses were pursued using flow cytometry. It has been reported that flow cytometry and confocal microscopy are highly correlated when used to analyze fluorescent bead engulfment in cell culture [175]. Moreover, automated analyses conducted with flow cytometry are more objective, faster, and are generally more powerful when compared to confocal microscopy as a result of the large sample size they are able to process [175]. 6.3.3 Phagocytosis of vitronectin-coated beads quantified using flow cytometry Before testing cilengitide and the PB-probe, it was necessary to confirm flow cytometry as an appropriate analysis technique to resolve differences in fluorescent bead engulfment by BV-2 microglia. Two negative controls were employed for this confirmation. First, cells were pre-treated with cytochalasin B, a mycotoxin that inhibits actin network formation and thereby blocks the endocytic mechanisms required to internalize beads @ 1 µm in diameter [139-142]. It was expected in this control that low levels of fluorescence would be detected as a result of beads bound to the surface of the cells but not internalized. Second, it has been reported that uncoated beads are also bound by cells, but not ingested, while cells coated with ECM substrate trigger select molecular signaling pathways to initiate phagocytosis [157, 162]. To further deduce the role of vitronectin-coating on bead engulfment, and whether integrin aVb5 is engaged, phagocytosis of uncoated beads by simulated BV-2 cells was tested after pre-treatment with 500 nM of cilengitide or PB-probe. It was hypothesized that if only beads coated with ECM protein are effectively internalized, then inhibition of integrin aVb5 by cilengitide or the PB-probe should not affect the fluorescence intensity of cells incubated with uncoated beads. The results of this experiment are presented in Figure 6.7. 74 Figure 6.7 Engulfment of vitronectin-coated beads is actin-dependent and more efficient than uncoated beads Flow cytometry data from 4 independent experiments are presented as mean ± S.E.M. LPS-stimulated BV-2 microglia were treated with DMSO (0.05% v/v), cilengitide or PB-probe (500 nM), or cytochalasin B (2.5 µg/mL) then incubated with beads (0.5% solid suspension diluted 200 x into cell culture dish) for 2.5 h. A) Fluorescent beads bound to, but not internalized, by stimulated BV-2 cells account for @ 28% of cellular fluorescence intensity. Median fluorescence intensities are reported as-collected from the flow cytometer. B) Only treatment with 2.5 µg/mL cytochalasin B reduces the percent of engulfing cells (24% decrease). The percent of engulfing cells was calculated as the number of cells with a fluorescence intensity greater than that of cells incubated in the absence of beads. The effects of treatments were assessed by the one-way ANOVA, followed by Dunnett’s post-hoc test. ANOVA F and P values are presented on the graphs. **P<0.01 and *P<0.05, significantly different from stimulated control cells incubated with vitronectin-coated beads in the absence of drugs. Figure 6.7A confirms that the engulfment of vitronectin-coated beads is actin-dependent, as indicated by the higher median fluorescence intensity of control cells relative to those treated with cytochalasin B. A similar trend was observed comparing the fluorescence intensity of BV-2 cells incubated with vitronectin-coated beads to those treated with uncoated beads, indicating that uncoated beads do not engage signaling pathways required for efficient phagocytosis. In particular, the finding that cilengitide and the PB-probe do not block the engulfment of uncoated beads (Fig. 6.7A) suggests that integrin aVb5 does not participate in phagocytosis directed at uncoated beads. To a further point, the similar values obtained for cells both pre-treated with cytochalasin B and incubated with vitronectin-coated beads, along with cells incubated with uncoated beads are consistent with literature reports that both of these treatments result in cells binding to, but not engulfing, fluorescent beads [157, 162]. Taken together, these data indicate that fluorescent beads bound to, but not internalized, by stimulated BV-2 cells account for @ 28% 0.1% DMSO2.5 µg/mLCytochalasin B0.1% DMSO500 nMCilengitide500 nM PB-Probe 051015Fluorescence Intensity (AU)********Vitronectin-coated beadsUncoatedbeadsF = 13.24P = 0.00020.1% DMSO2.5 µg/mLCytochalasin B0.1% DMSO500 nMCilengitide500 nM PB-Probe050100Engulfing Cells (%)*Vitronectin-coated beadsUncoatedbeadsF = 3.531P = 0.0368B A 75 of the fluorescence intensity recorded by the flow cytometer, with the remaining 72% resulting from internalization of the beads following engagement of select signaling pathways by vitronectin. Interestingly, the only treatment indicated by Figure 6.7B to reduce the percentage of engulfing cells was 2.5 µg/mL cytochalasin B. In this case, the axis label “Engulfing Cells (%)” is a misnomer, more accurately describing the percentage of cells with a higher fluorescence intensity than cells incubated in the absence of beads. Cells that have bound, but not internalized, fluorescent beads would therefore be above this threshold. It was necessary to segregate data in this manner to make distinct any effects of integrin aVb5 inhibitors in reducing phagocytosis rather than decreasing the number of phagocytic cells. Since cytochalasin B was employed to block bead engulfment completely, it may therefore be that the decrease in the percentage of engulfing cells observed with this treatment was instead due to fewer bound beads. To explore potential causes for this phenomenon, the effect of cytochalasin B on cell viability was investigated. Figure 6.8 illustrates the results of an MTT analysis following treatment of BV-2 microglia with cytochalasin B. Treatment with cytochalasin B demonstrates a significant cytotoxic effect at the concentration and time period employed in the vitronectin-coated bead engulfment assay. Compellingly, the 26% decrease in cell viability observed nearly duplicated the 24% decrease in the number of cells above the fluorescence threshold in the flow cytometry data presented in Figure 6.7B. It is therefore suggested that the decreased number of cells included by the fluorescence criterion following treatment with cytochalasin B is due to a reduction in cell viability. Since tetrazolium dye conversion techniques indicate cytotoxicity more accurately than exclusion dyes, this may be why data from cytochalasin B-treated cells were not excluded by the PI gating parameter [176]. 76 Figure 6.8 Treatment with cytochalasin B reduces BV-2 cell viability BV-2 cells were treated with cytochalasin B (2.5 µg/mL) then incubated for 3 h. Cell viability was measured using the MTT assay. Data from 3 independent experiments were calculated using equation [6.1] and are presented as mean ± S.E.M. The effect of cytochalasin B was evaluated using an unpaired, two-tailed t test. **P<0.01, significantly different from control cells incubated in the absence of drugs. Having confirmed that flow cytometry effectively resolves differences in fluorescent bead engulfment by BV-2 microglia, two additional experiments were conducted to control for intraexperimental variation prior to testing the concentration-dependent effects of cilengitide and the PB-probe. The effects of sample analysis order and the number of beads incubated per cell on resulting fluorescence intensity were investigated. First, the effect of sample analysis order on fluorescence intensity was tested since the workflow of the flow cytometry analysis lasts approximately 6 min per sample. Unlike the confocal microscopy experiments, where all cells are fixed with formaldehyde at the same time (preventing further bead engulfment), it was necessary to keep cells alive for the flow cytometry experiments. This ensured that nonviable cells could properly be controlled for using PI; however, it also meant that the first and last samples were analyzed nearly 40 min apart, or roughly 25% of the total bead incubation time. Although the cells had already been washed several times and trypsinized by this point, the still-viable cells might have continued to engulf beads throughout this delay, especially considering that at this timepoint they were effectively in the absence of any remaining inhibitors. The data from this experiment are presented in Figure 6.9. Figure 6.9A indicates that while there appears to be a slight trend for the fluorescence intensity of engulfing cells to increase with time, this variation was not statistically significant. Moreover, Figure 6.9B illustrates that delayed sample analysis does not have any effect on the percentage of cells actively engulfing beads throughout the experiment. It was therefore concluded that washing 0 2.5050100Cytochalasin B(µg/mL)Cell Viability (%)** 77 and trypsinization were sufficient to both clear free-floating beads and prevent continued phagocytosis during the delay period. Figure 6.9 Time elapsed before flow cytometry analysis does not affect fluorescence intensity LPS-stimulated BV-2 microglia were incubated with vitronectin-coated beads (0.5% solid suspension diluted 200 x into cell culture dish) for 2.5 h. Flow cytometry data from 3 independent experiments are presented and fit with linear regressions. A) Median fluorescence intensities were normalized against cells that were not incubated following sample preparation. The effect of sample incubation time was not significantly non-zero; P=0.4485, F=0.6444. B) The percent of engulfing cells was calculated as the number of cells with a fluorescence intensity greater than that of cells incubated in the absence of beads. The effect of sample incubation time was not significantly non-zero; P=0.0817, F=10.76. The effect of treatments was further assessed by the one-way ANOVA. ANOVA F and P values are presented on the graphs. No significant differences were observed. Second, the effect of bead concentration on fluorescence intensity was investigated. Engulfment of vitronectin-coated beads by LPS-stimulated microglia is limited by bead availability and by overall phagocytic capacity. Since, on average, the number of beads engulfed by cilengitide- and PB-probe-treated cells (Fig. 6.3 and 6.4) did not approach the maxima seen in untreated cells, bead availability was expected to have a larger influence on engulfment than saturation of phagocytic machinery. It was therefore hypothesized that the number of beads incubated per cell would proportionately increase the resulting fluorescence intensity. The number of polystyrene beads per millilitre was calculated using the following equation provided by the supplier: N = SPL(6 x 1010)/pPsd3 [6.2] Where S represents the % solids (w/w), Ps is the density of bulk polymer (g/mL), PL is the density of latex (g/mL), and d is the diameter of the beads in µm. The supplier-reported values for these parameters are reported in Table 6.1. 0 10 20 30 40050100150Time (min)Relative Fluorescence Intensity (%)y = 0.5786x + 95.85R2 = 0.08429F = 0.1641P = 0.85240 10 20 30 40050100Incubation Time (min)Engulfing Cells (%)y = 0.0250x + 97.25R2 = 0.8433F = 0.05787P = 0.9805B A 78 Table 6.1 Physical attributes of polystyrene latex beads Values provided by the supplier, Sigma Aldrich. S Ps PL d 2.5% 1.05 g/mL (polystyrene) 1.005 g/mL 1 µm Substituting the values presented in Table 6.1 into equation [6.2] returns a value of N = 4.57 x 1010 beads/mL. When the beads were prepared with vitronectin, they were diluted 5 x resulting in a concentration of 9.14 x 109 beads/mL. Finally, 10 µL of vitronectin-bead solution were diluted into 2 mL of cell culture media (200 x), further resulting in a concentration of 4.57 x 107 beads/mL (9.14 x 107 beads total). Approximately 75,000 cells were plated per experiment, resulting in a final concentration of 1218 beads per cell. Rather than manipulate the number of cells to determine the effect of the number of beads incubated per cell on fluorescence intensity, three parallel conditions were set up where cells were treated with 1 µL, 5 µL, or 10 µL of bead solution (ranging from approximately 120-1200 beads per cell) for the 2.5 h incubation period. The data in Figure 6.10 confirm the hypothesis that the number of beads per cell has a robust effect on the fluorescence intensity of engulfing cells, as detected by flow cytometry. Similarly, Figure 6.10B demonstrates that reducing the number of beads incubated per cell has a significant effect on the percentage of cells that actively engulfed beads throughout the experiment. While the statistical significance of this last trend was not supported by the linear regression model, this is indicative only that the variables do not share a linear dependence. Overall, the findings in Figure 6.10 have two key implications. Firstly, that both the cell culture densities and bead solutions need to be diluted with a high level of accuracy to ensure that error does not propagate to the experimental results. Second, that cells incubated with more beads have a proportionally greater phagocytic response. 10 µL of bead solution were used for subsequent experimentation. 79 Figure 6.10 The number of beads incubated per cell affects fluorescence intensity LPS-stimulated BV-2 microglia were incubated with vitronectin-coated beads (0.5% solid suspension diluted 200 – 2000 x into cell culture dish) for 2.5 h. Flow cytometry data from 3 independent experiments are presented and fit with linear regressions. A) The effect of bead concentration on median fluorescence intensity was significantly non-zero; P<0.0001, F=815.6. B) The percent of engulfing cells was calculated as the number of cells with a fluorescence intensity greater than that of cells incubated in the absence of beads. The effect of bead concentration was not significantly non-zero; P=0.3455, F=2.749. The effect of treatments was further assessed by the one-way ANOVA, followed by Tukey’s multiple comparisons post-hoc test. ANOVA F and P values are presented on the graphs. **P<0.01; significantly different from cells incubated with 1 µL of bead solution added. ##P<0.01; significantly different from cells incubated with 5 µL of bead solution added. 6.3.4 Effects of integrin aVb5 inhibitors on bead engulfment (flow cytometry) Satisfied that intraexperimental variations could adequately be controlled for, the effects of cilengitide and the PB-probe on phagocytic processes in BV-2 cells were analyzed. Since variation is tolerated at the D-phenylalanine meta position of cilengitide, it was hypothesized that cilengitide and the PB-probe, through integrin aVb5 inhibition, would block the engulfment of vitronectin-coated beads with equal efficacy and in a concentration-dependent manner [77]. Cells were treated with 5 nM – 50 µM of cilengitide or PB-probe for 15 min prior to a 2.5 h incubation with vitronectin-coated beads and subsequent analysis with flow cytometry. The data from these experiments are presented in Figure 6.11. Figure 6.11A indicates that cilengitide and the PB-probe inhibit the engulfment of vitronectin-coated beads by stimulated microglia in vitro with IC50 values of 0.6 ± 2 µM and 0.9 ± 0.3 µM, respectively. Statistical analyses concluded that only the 5 µM and 50 µM concentrations of both compounds are significantly different from cells treated in the absence of inhibitors (two-way ANOVA, followed by Tukey’s multiple comparisons post-hoc test). Importantly, none of the effects of cilengitide or the PB-probe were 0 5 10020406080100Bead Solution Added (µL)Fluorescence Intensity (AU)y = 7.837x-1.253R2 = 0.9915F = 356.4P <0.0001****##0 5 10050100Beads/CellEngulfing Cells (%)y = 21.28x + 81.13R2 = 0.7332F = 83.7P <0.0001** **B A 80 significantly different from one another and their IC50 value are within the range of error from each other. Statistically then, both inhibitors have an equivalent efficacy in blocking vitronectin-coated fluorescent bead engulfment, confirming that the PB-probe maintains integrin-binding potency, relative to cilengitide. Figure 6.11 Cilengitide and the PB-probe are equally effective at blocking vitronectin-coated bead engulfment LPS-stimulated BV-2 microglia were treated with DMSO (0.05% v/v) or cilengitide or PB-probe (5 nM – 50 µM) then incubated with vitronectin-coated beads (0.5% solid suspension diluted 200 x into cell culture dish) for 2.5 h. Flow cytometry data from 12 independent experiments are presented as mean ± S.E.M. and fit with nonlinear inhibitor versus response curves (equation [2.1]). A) Median fluorescence intensities were normalized against cells incubated in the absence of drugs. Twenty eight percent relative fluorescence intensity is indicated by a dashed line, which was determined experimentally (Fig. 6.7) to result from binding, not engulfment, of fluorescent beads. B) The percent of engulfing cells was calculated as the number of cells with a fluorescence intensity greater than that of cells incubated in the absence of beads. The effects of treatments were assessed by the two-way ANOVA, followed by Tukey’s multiple comparisons post-hoc test. ANOVA F and P values are presented on the graphs. **P<0.01 and *P<0.05, significantly different from stimulated control cells incubated in the absence of PB-probe; ##P<0.01, significantly different from stimulated control cells incubated in the absence of cilengitide. Notably, the IC50 values obtained from the experiments presented in Figure 6.11A are higher, by 2-3 orders of magnitude, than what has previously been reported for cilengitide (integrin aVb3 = 0.65 ± 0.07 nM; integrin aVb5 = 11.7 ±1.5 nM) [72, 78]. However, the data presented herein describe the efficacy of cilengitide and the PB-probe in blocking microglial phagocytic response to vitronectin-coated beads, while literature values indicate the concentration at which cilengitide competitively inhibits ligand binding to integrin aVb3 and aVb5 [78]. Therefore, it is expected that a higher concentration of cilengitide and PB-probe are necessary to affect a physiological response, compared to the concentration required for competitive inhibition of the 100 101 102 103 104 105050100Concentration (nM)Relative Fluorescence Intensity (%)PB-ProbeCilengitide## ##PB-Probe IC50 = 903 ± 348 nMCilengitide IC50 = 623 ± 235 nM** **Drug F = 5.742 P = 0.0232Concentration F = 55.2P <0.0001100 101 102 103 104 105050100Concentration (nM)Engulfing Cells (%)Cilengitide PB-Probe**Drug ConcentrationF = 3.604 F = 11.35P = 0.0649 P <0.0001B A 81 binding site. A detailed analysis of the structurally-related Alexa FluorÒ 568 probe, its binding affinity to purified aVb5, and its IC50 in the context of purified protein is presented in Chapter 4. 6.3.5 Inhibitory effect of an Alexa FluorÒ 568 probe on glial phagocytosis An alternate probe bearing a different fluorescent dye (one with an Alexa FluorÒ 568 dye in place of PB, Figure 6.12A) was created due to instrumental constraints when determining the KD of the probe, explained in Chapter 4. Because covalent modifications have been shown to affect binding affinity, we sought to understand the relative efficacy of the PB-probe and the Alexa FluorÒ 568 probe in blocking integrin aVb5 [72]. Since the binding affinity of the PB-probe could not be quantified relative to purified integrin aVb5, the vitronectin-coated bead engulfment assay was repeated with the Alexa FluorÒ 568 probe. All experimental parameters were kept the same as the previous analyses with cilengitide and the PB-probe. Data are presented in Figure 6.12B-C, alongside the previously reported cilengitide data for ease of comparison. The Alexa FluorÒ 568 probe was found to inhibit the engulfment of vitronectin-coated beads by stimulated microglia in vitro with an IC50 value of 71 ± 23 nM (Fig. 6.12). While the differences in IC50 values between the Alexa FluorÒ 568 probe and cilengitide are not within the range of error from each other, it is important to note that none of the effects of cilengitide or the Alexa FluorÒ 568 probe were significantly different from one another. 82 Figure 6.12 Cilengitide and the Alexa FluorÒ 568 probe block vitronectin-coated bead engulfment A) The chemical structure of the Alexa FluorÒ 568 probe contains the integrin aVb5 inhibitor cilengitide and the Alexa FluorÒ 568 fluorophore, coupled at the D-phenylalanine meta position. B-C) LPS-stimulated BV-2 microglia were treated with DMSO (0.05% v/v) or cilengitide or Alexa FluorÒ 568 probe (5 nM – 50 µM) then incubated with vitronectin-coated beads (0.5% solid suspension diluted 200 x into cell culture dish) for 2.5 h. Flow cytometry data from 9 independent experiments are presented as mean ± S.E.M. and fit with nonlinear inhibitor versus response curves (equation [2.1]). The effects of treatments were assessed by the two-way ANOVA, followed by Tukey’s multiple comparisons post-hoc test. ANOVA F and P values are presented on the graphs. ffP<0.01 and fP<0.05, significantly different from stimulated control cells incubated in the absence of Alexa FluorÒ 568 probe; ##P<0.01, significantly different from stimulated control cells incubated in the absence of cilengitide. B) Median fluorescence intensities were normalized against cells incubated in the absence of drugs. Twenty eight percent relative fluorescence intensity is indicated by a dashed line, which was determined experimentally (Fig. 6.7) to result from binding, not engulfment, of fluorescent beads. C) The percent of engulfing cells was calculated as the number of cells with a fluorescence intensity greater than that of cells incubated in the absence of beads. Three possible explanations are offered for the apparently higher activity of the Alexa FluorÒ 568 probe compared to cilengitide and the PB-probe. First, Alexa FluorÒ 568 has more formal 100 101 102 103 104 105050100Concentration (nM)Engulfing Cells (%)Alexa Fluor® 568 ProbeCilengitideDrug ConcentrationF = 0.09 F = 4.85P = 0.764 P = 0.002100 101 102 103 104 105050100Concentration (nM)Relative FluorescenceIntensity (%)Cilengitideφφ φφCilengitide IC50 = 623 ± 235 nMAlexa Fluor® 568 Probe IC50 = 71 ± 23 nMAlexa Fluor® 568 Probe## ##φφφDrug F = 0.5308 P = 0.4728Concentration F = 56.43P <0.0001C B A 83 charges at physiological pH than PB, so it is possible that Alexa FluorÒ 568 forms additional electrostatic interactions with integrin aVb5. Through these stabilizing interactions, Alexa FluorÒ 568 may effectively decrease the KD of the Alexa FluorÒ 568 probe, increasing its lifetime in the RGD-binding domain and thereby its capacity for competitive inhibition towards other ligands. This explanation may be indicated by the results in Chapter 4 (Fig. 4.4) that the Alexa FluorÒ 568 probe occupies more RGD-binding domains than cilengitide at an equivalent concentration (10 nM). Second, it was hypothesized that the Alexa FluorÒ 568 probe may negatively affect the viability of BV-2 cells. While PI should eliminate most cells with reduced viability from the final analysis, a more precise measure was warranted. Figure 6.13 presents the results of an MTT analysis on BV-2 cells treated with 0 – 100 µM of the Alexa FluorÒ 568 probe, indicating that this range of concentrations has no cytotoxic effects on stimulated microglia over the time period used for engulfment experiments. Figure 6.13 The Alexa FluorÒ 568 probe is not cytotoxic BV-2 cells were treated with Alexa FluorÒ probe (10 nM - 100 µM) then incubated for 3 h. Cell viability was measured using the MTT assay. Data from 4 independent experiments were calculated using equation [6.1] and are presented as mean ± S.E.M. The effects of the Alexa FluorÒ probe were assessed by the one-way ANOVA, followed by Dunnett’s multiple comparisons post-hoc test. ANOVA F and P values are presented on the graph. No significant differences from control cells incubated in the absence of drugs were observed. As a final, and most likely explanation, it was noted that the lot number of the beads used for the Alexa FluorÒ 568 experiments was different than those used to test cilengitide and the PB-probe. Visually, these beads were whiter than the previous lot, possibly indicating the presence of less fluorescent dye. Since all data are presented as relative values standardized to intraexperimental controls, this was not expected to be a meaningful discrepancy. However, the gain settings, gating parameters, and fluorescence threshold for the flow cytometer were all set based on the properties of the more fluorescently intense lot of beads. Since the cells treated with Alexa 00.01 0.1 1 10 100050100Alexa Fluor® 568 Probe (µM)Cell Viability (%)F = 3.032P = 0.037 84 FluorÒ 568 were incubated with less fluorescent beads, it is possible that cells engulfing fewer beads were excluded by these thresholds, skewing the data to yield an artificially low IC50 value for the Alexa FluorÒ 568 probe. In support of this explanation, it can be seen in Figure 6.12C that the 500 nM concentration of Alexa FluorÒ 568 probe has the lowest percentage of engulfing cells relative to cilengitide. Importantly, this data point weighs the most in the calculation of the IC50 value as it defines an inhibitory effect that is between zero and complete inhibition. These findings prevent positively concluding that the Alexa FluorÒ 568 probe has a more potent effect than cilengitide and the PB-probe in blocking integrin aVb5-mediated engulfment of fluorescent beads. Nevertheless, the suite of bead engulfment experiments described herein was designed to determine whether the PB-probe and the Alexa FluorÒ 568 probe have a reduced capacity to block integrin aVb5-mediated signaling cascades. This has been shown unequivocally to not be the case, as both probes have comparable, or lower, IC50 values relative to cilengitide in this assay. Moreover, the effects of the PB-probe and the Alexa FluorÒ 568 probe were similar to cilengitide, further supporting that these molecules have retained an analogous potency towards integrin aVb5. Overall, these findings support the conclusion that both the PB-probe and the Alexa FluorÒ 568 probe maintain a similar activity as cilengitide in the context of binding integrin aVb5 and inhibiting its associated signaling cascades in vitro. 6.4 Summary Together, the experiments described above demonstrate that stimulated BV-2 microglia are a robust in vitro model for the investigation of cellular signaling pathways engaged in phagocytosis. This activity is quantifiable through both confocal microscopy and flow cytometry. Flow cytometry was shown to be a superior detection method to measure bead engulfment due to its objectivity, speed, and capacity for high-throughput analysis. Through this technique, it was discovered that vitronectin-coated beads engaged RGD-binding integrins to initiate phagocytosis in an actin-dependent manner. Both cilengitide and the PB-probe were able to inhibit this phenomenon with equal potency, confirming the hypothesis that the PB-probe maintains inhibitory effect towards integrin with equal efficacy as cilengitide. Similarly, the Alexa FluorÒ 568 probe demonstrated a potency that is equal to or higher than cilengitide, further supporting that modification of the D-phenylalanine ring at the meta position of cilengitide does not reduce binding potency towards integrin aVb5. An additional conclusion of these experiments was that all three integrin aVb5 inhibitors were well tolerated by cell culture, affecting no decrease in cell viability at concentrations of up to 100 µM. 85 7 Conclusion In conclusion, this study demonstrates a selective and robust means of fluorescently labeling endogenous integrin aVb5 in vitro. The current state of the art in labeling endogenous protein relies on immunostaining or genetic modification techniques; however, both methods suffer from limitations. For instance, immunocytochemistry is a fairly rapid technique, but the need to fix cells prior to labeling prevents the study of proteins through time (moreover, commercial antibodies are known to sometimes be unreliably specific) [66]. Genetic modification techniques such as fusing a protein of interest to a fluorescent protein (e.g., GFP) offers 100% specific labeling, but the expression of recombinant proteins through transfection often perturbs the model system. While CRISPR/Cas9 targeted genome editing enables the study of endogenous protein and causes minimal perturbation to the model system, its key drawbacks are the time- and labor-intensive processes of developing a model organism. The PB-probe discussed in this thesis improves on the existing methods of protein labeling listed above through two main advantages: its ease-of-use and its rapid applicability. Chemically, the PB-probe retained similar spectrophysical properties as PB. In spectral characterizations, the PB-probe was found to be photostable through time, amenable to detection in physiologically relevant buffers, and well-suited for excitation by commonly available 405 nm lasers. In assays performed with purified integrin αVβ5 protein, the Alexa FluorÒ 568 probe maintained a high binding affinity towards integrin aVb5. Changes in fluorescence intensity and anisotropy assays indicate that the Alexa FluorÒ 568 probe binds integrin αVβ5 with a KD of 0.11 ± 0.07 µM. Competitive displacement assays revealed a similar finding: that the Alexa FluorÒ 568 probe has a binding activity comparable to other integrin αVβ5 ligands, cilengitide and vitronectin. In cell assays, application of the PB-probe demonstrated that the optimal labeling parameters are a 3 h incubation with 1 µM PB-probe, followed by two washes with PBS. These labeling parameters are more rapid than other labeling methods and are compatible with simultaneous immunocytochemistry techniques [134]. The selectivity of the PB-probe is supported by its colocalization with established detection methods (59 ± 1% overlap), the finding that cells are labeled much more effectively by the PB-probe compared to the PB-linker (almost three-fold increase), and the similarity of its labeling pattern to the subcellular localization of the aV and b5 subunits outlined in previous reports [50]. Preliminary results in zebrafish established that the PB-probe is suitable for in vivo applications. Finally, it was found that stimulated BV-2 microglia serve as a robust model for phagocytic activity in vitro. The engulfment of vitronectin-coated beads was shown to be integrin-dependent 86 and quantifiable through flow cytometry. Cilengitide, the PB-probe, and the Alexa FluorÒ 568 probe blocked phagocytosis in a concentration-dependent manner, all with similar potency in the nanomolar range. This effect was not due to a reduction in cell viability. Together, these data demonstrate that the probes retain their IC50 towards integrin aVb5-mediated signaling cascades, relative to cilengitide. This study confirms that the PB-probe is a powerful fluorescence imaging tool to track endogenous integrin aVb5 in live-cell systems. 7.1 Limitations While this study was successful in confirming the utility of a ligand-directed labeling molecule to track the subcellular localization of integrin aVb5 through fluorescence microscopy, it does have some limitations. First, various data are shown that represent single experiments. There are several shortcomings associated with reporting data in this matter; namely, if the observed results were due to an error or were an uncharacteristic phenomenon, this would not be discovered through subsequent experimentation. Another consideration is that data from single experiments are not suitable for statistical analysis. Second, the binding affinity of the PB-probe towards purified integrin αVβ5 protein was not measured. It is expected that the results from the Alexa FluorÒ 568 probe are translatable to the PB-probe, since these molecules share several structural features. However, the PB dye differs in its chemical structure from Alexa FluorÒ 568, and therefore may affect the binding affinity of the PB-probe. Third, there were unexpected observations using the PB-probe to label cell cultures in vitro. The subcellular localization and fluorescence intensity of PB-probe labeling varied between: live cells, fixed and permeabilized cells, and fixed but not permeabilized cells (Fig. 5.7). The implications of these observations are not fully understood. Additionally, two independent methods to determine the specificity of PB-probe binding to integrin αVβ5 showed that as much as 40% of PB-probe labeling might be nonspecific (Fig. 5.7 and 5.9). Another investigation of labeling specificity, competitive inhibition of PB-probe labeling, was tested in this study using an equimolar concentration of PB-probe and cilengitide (Fig. 5.6); however, a range of cilengitide concentrations would likely provide more information on the specificity of the PB-probe and its competition for the integrin αVβ5 RGD-binding domain. It is advised that such an investigation be conducted in a high-throughput manner, as analysis with confocal microscopy would be unnecessarily labour-intensive. 87 Additionally, the use of integrin knockout organisms or cell lines may help further elucidate the specificity of the PB-probe. Integrin β5 knockout models have been generated both in mice and in cell culture by previous research groups and these organisms exhibit good survivability [105, 177]. Contrarily, integrin αV knockout mice have an 80% mortality rate prior to delivery and display extensive malformations [178]. These findings complicate the use of true integrin αVβ5 knockout organisms as a research model; however, the use of integrin β5 knockout systems is suggested to provide several meaningful insights into the specificity of the PB-probe. A final limitation of this research project is the use of the BV-2 microglial cell line in place of primary brain cells. However, since the goal of this study was to evaluate the PB-probe as a tool to label and track endogenous integrin aVb5 in live cells, extension of the findings to the actual functioning of microglia was minimal. It is anticipated that applying the PB-probe to primary microglial cells (or cells of any other type) would return both a different labeling pattern and effect on endogenous integrin aVb5 signaling. However; this dynamism also underlies the utility of the PB-probe since changes in labeling patterns and signaling effects will provide unique insights into the role of integrin aVb5 in cells of different lineage and environment. 7.2 Future Directions The next phase of this research involves investigating the role of glia-mediated synaptic elimination in physiological and pathological conditions pertaining to neurodevelopment and neurodegenerative disease. It has been suggested that integrin aVb5-mediated phagocytic cascades directly contribute to disease-associated cognitive deficits [13-16, 21-24, 179-181], but methods to monitor integrin aVb5 in unperturbed live-cell systems have been unavailable until now. Since the subcellular localization of integrin aVb5 directly relates to its functional role, tracking its position with the PB-probe enables real-time evaluation of integrin-mediated phagocytosis [41]. Applied to cocultures of neurons and glia in vitro, or nervous tissue ex vivo, the PB-probe will serve as a useful diagnostic tool to investigate: 1) stimuli that precipitate phagocytic responses, and 2) how phagocytosis can be modulated by prospective therapeutics. Real-time evaluation of these processes will provide insights into the precise signaling mechanisms operating at the cellular level. More importantly, it will be possible to elucidate whether phagocytic responses by glia are beneficial or deleterious to neuron function throughout physiological and pathological conditions. Fully understanding the role of glia-mediated phagocytosis, and its contributions to both neural circuit refinement and irreversible neuron loss, 88 will aid in the development of therapeutics that modulate this function to optimize clinical outcomes. A current limitation of the PB-probe is that its binding to integrin aVb5 also inhibits signaling function. To circumvent this drawback, a second-generation labeling probe is being created that tethers cilengitide to a fluorophore through a reactive linker (Fig. 7.1A). This molecule is designed to covalently link the fluorophore to integrin aVb5, allowing cilengitide to dissociate from the RGD-binding domain (Fig. 7.1B). Through this process, the second-generation PB-probe is hypothesized to fluorescently label endogenous integrin aVb5 without inhibiting signaling. This tool will share the selective, rapidly applicable, and robust labeling properties of the PB-probe, enabling the study of integrin aVb5 function in effectively unaltered living cell systems. Figure 7.1 Second-generation PB-probe concept A) The second-generation PB-probe will be comprised of cilengitide coupled to PB with a cleavable linker at the D-phenylalanine meta position. B) Cartoon depiction of fluorophore delivery to integrin aVb5 by the second-generation PB-probe. 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Nat Med, 2018. 24(7): p. 931-8. 100 Appendices Appendix A: Drafted manuscript excerpt The synthesis of imaging probe 2 was carried in a total of 27 steps, with 19 steps for the longest linear sequence. The synthesis of a novel meta-substituted d-phenylalanine derivative 9 is outlined in Figure A.1A. m-Toluic acid (4) was converted to benzylic bromide 9 in 5 steps. The stereocenter of the amino acid precursor was installed via enantioselective alkylation of glycinyl imine 6 with a minute amount of Maruoka’s catalyst (7) under biphasic conditions (0.1 mol% catalyst loading). Enantioselectivity of the unnatural chiral center was determined to be 94% ee. Three additional steps were required to obtain the modified d-phenylalanine 9 bearing appropriate protecting groups for solid phase synthesis. Protected N-methyl valine 12 was prepared according to a reported procedure (88% yield over two steps). The pentapeptide was synthesized on trityl chloride resin 10, starting with Fmoc-protected glycine. HBTU was used as the principal coupling agent, with the exception of coupling of phenylalanine 9 to N-methylamine of 12, where more reactive HATU was used. The final pentapeptide 13 was cleaved from the resin with hexafluoroisopropanol and purified by chromatography to give 97:3 mixture of two diastereomers 14, corresponding to 94% ee of phenylalanine 9. Cyclization of pentapeptide was effected under high dilution conditions with diphenyl phosphoroazidate as the coupling reagent. Global deprotection in 95% TFA led to cyclic pentapeptide 15 in very good yield. Synthesis of the imaging probe 2 was completed by assembling the modified cyclic pseudopeptide 15 and a fluorophore via a six atoms linker chain (Figure A.1B). In short, Pacific Blue (16) was activated as a succinimidyl ester, coupled with 3-oxa-ω-aminovaleric acid, and converted to succinimidyl ester 17. Coupling of 15 with active ester 17 led to non-reactive fluorescent probe 2. The lower yield was caused by losses during purification by reverse-phase HPLC. Fluorescent probe 3 for fluorescence anisotropy assay was prepared from fully protected pentapeptide 18 (Figure A.1C). Greater reactivity of tert-butyl carbamate enabled selective deprotection and coupling with active ester of ω-azidobutyric acid. Following global deprotection afforded peptidic azide 19. Alexa 568 dye (20) was transformed into its propargyl glycinamide 21 by peptide coupling reagent HATU. Both fragments were then connected by copper-catalyzed azide-alkyne click chemistry to yield orange-fluorescent probe 3. 101 Figure A.1 Caption on following page 102 Figure A.1 Synthesis of a modified cyclo(-RGDf-MeV-) bearing a synthetic handle for molecular cargo attachment (A) Synthesis of meta substituted d-phenylalanine residue 9. (B) Synthesis of the imaging probe 2 by assembling 15 with a linker chain and coumarin fluorescent dye. Briefly, solid phase synthesis with a trityl resin was used to prepare the linear pentapeptide: the first two amino acids were assembled on the resin, followed by N-Fmoc protected N-methyl valine 12 , unnatural residue 9 and protected aspartate. Cyclization and global deprotection yielded cyclo-RGD(3’-aminomethylene)f(Me)V 15, which was acylated at the benzylic amine position with the activated NHS ester of fluorescent coumarin 17. Detailed conditions and experimental procedures are provided in the supporting information. (C) Synthesis of the orange imaging probe 3 by click chemistry of azide-containing peptide 19 and alkyne-Alexa 21. Appendix B: Microglia and neurodegenerative disease Neurodegenerative diseases include a broad range of pathologies that lead to the targeted death of neurons. Diseases in this category exhibit many unique pathological mechanisms that exert toxicity neurons of the CNS [182-185]. These diseases are commonly referred to as dementia, due to the gradual and irreversible loss of cognitive and motor function throughout pathogenesis. While several neurodegenerative diseases precipitate dementia, including Parkinson’s disease, dementia with Lewy bodies, and vascular dementia, the most common is AD [186]. Each of these malignancies have characteristic hallmarks that distinguish one from another. Such hallmarks typically include proteinaceous deposits; AD is characterized by extracellular Aβ plaques and intraneuronal neurofibrillary tangles while Lewy bodies present in both Parkinson’s disease and dementia with Lewy bodies [187, 188]. Vascular dementia is most adequately indicated by inadequate blood flow through the brain, commonly associated with stroke [189]. One aspect that is common to all of these diseases is inflammation of the nervous tissue, or neuroinflammation [190]. Neuroinflammation in the CNS mediated by glia is necessary for both the initiation and propagation of neurodegenerative disease pathology [191, 192]. Inflammatory mediators including complement, oxygen free radical species, cytokines, and other factors such as acute phase proteins have been suggested to contribute significantly to the development and progression of neurodegenerative disease [192, 193]. Even in the absence of traditional disease hallmarks such as Aβ deposits in Alzheimer’s disease, inflammatory stimuli are sufficient to accelerate neurodegeneration and cause cognitive decline [194-196]. The release of pro-inflammatory mediators also leads to the simultaneous recruitment and proliferation of immune cells that propagate and amplify the inflammatory milieu [187]. While the brain is considered to be immunologically privileged due its independence from peripheral immune responses, imbued by the blood brain barrier, the inflammatory mediators described above lead to a robust activation pattern of the microglia [197, 198]. These pro-inflammatory microglia are consistently localized in regions of the brain exhibiting high levels of 103 neurodegenerative disease pathology [187, 195]. More recently, it has been demonstrated that microglia can be directly neurotoxic through phagocytic processes [13]. Derived from the monocyte lineage, microglia are the main effectors of inflammation within the CNS and function in an innate immunity capacity to protect against infection and neurotoxic damage [6]. Some reports have indicated that microglial activation may have an overall beneficial effect on inflammatory resolution through the secretion of chemokines and growth factors [199]. These factors have been hypothesized to ameliorate the neurotoxic effects of cerebrovascular damage and inflammation [199, 200]. However, there is growing evidence that the altered secretory profile and enhanced phagocytic activity of microglia following activation may instead cause neuron damage and accelerate neurodegeneration [201]. Disease-associated regions of the brain characterized by necrotic neurons frequently exhibit elevated levels of the immune cytokines tumor necrosis factor a, interleukins 6 and 10, as well as monocyte chemoattractant protein 1; all of which are microglial-derived factors that have demonstrated neurotoxicity in some circumstances [202]. Additionally, microglia have been shown to exert control over astrocyte activation during neuroinflammation [203]. In support of the hypothesis that the inflammatory activation of glia during neuropathology has detrimental effects on neuron viability, the inhibition of astrocyte activation by microglia in Parkinson’s disease has a neuroprotective effect [204]. Appendix C: Confocal microscopy acquisition parameters General Scan Mode: XYZ File Version: 1.2.6.0 System Name: FLUOVIEW FV1000 System Version: 3.1.2.2 Dimensions X Dimension: 800, 0.0 - 211.735 [um], 0.265 [um/Pixel] Y Dimension: 800, 0.0 - 211.735 [um], 0.265 [um/Pixel] Channel Dimension: 2, 1.0 - 2.0 [Ch] Z Dimension: 2.0 [um/Slice] 104 Image Primary Dimensions: X * Y Image Size: 800 * 800 [Pixel] Image Size (Unit Converted): 211.735 [um] * 211.735 [um] Bits / Pixel: 12 [bits] Acquisition Device: FV1000 Observation Mode: LSM Objective Lens: PLAPON 60 x O SC NA:1.40 Objective Lens Mag.: 60 x Objective Lens NA: 1.4 Scan Mode: XYZ Scan Direction: Oneway Sampling Speed: 8.0 [us/Pixel] Confocal: ON C.A.: 80 [um] Sequential Mode: None Integration Type: Line Kalman Integration Count: 4 Region Mode: None Find Mode: x1 Rotation: 0.0 [deg] Pan X: 0 [um] Pan Y: 0 [um] Zoom: x1.0 Laser 1 Wavelength: 405 [nm] Laser 1 Transmissivity: 2.5 [%] Channel 1 Channel Name: CHS1 Dye Name: Alexa Fluor 405 Spectrum Separation: ON PMT Detection Mode: Analog PMT Voltage: 750 [V] 105 Excitation DM Name: DM405/488/559/635 Excitation DM Position: 4 Excitation Wavelength: 405 [nm] Emission DM Name: SDM490 Emission DM Position: 5 Emission Wavelength: 422 [nm] BF Position: 450 [nm] BF Range: 10 [nm] Channel 2 Channel Name: TD1 Dye Name: None Spectrum Separation: OFF PMT Detection Mode: Analog PMT Voltage: 234 [V] Excitation DM Name: DM405/488/559/635 Excitation DM Position: 4 Excitation Wavelength: 405 [nm] Emission Wavelength: 0 [nm]
Thesis/Dissertation
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Biochemistry and Molecular Biology
Vancouver : University of British Columbia Library
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Evaluation of a ligand-directed labeling molecule targeting integrin alphaVbeta5
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