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The effects of oxidation on bilayer membranes studied using giant unilamellar vesicles
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The effects of oxidation on bilayer membranes studied using giant unilamellar vesicles
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THE EFFECTS OF OXIDATION ON BILAYER MEMBRANES STUDIED USING GIANT UNILAMELLAR VESICLES A Dissertation Presented to the FACULTY OF THE USC GRADUATE SCHOOL UNIVERSITY OF SOUTHERN CALIFORNIA In Partial Fulfillment of the Requirements for the Degree DOCTOR OF PHILOSOPHY (CHEMICAL ENGINEERING) Shalene Sankhagowit May 2016 In honor of Mrs. Cathy Walker, my high school biology teacher, whose fascination in the science of life was contagious and inspirational. Acknowledgements I feel most fortunate and grateful to have been pursuing my Ph.D. degree under the advisement of Dr. Noah Malmstadt. With his guidance, my skills in conducting and communicating research work have undergone tremendous growths. His constructive teaching methods in encouraging questions and expressions of ideas have been instrumental in my training to become an independent thinker and pursuer of knowledge. I would also like to acknowledge Dr. Michelle L. Povinelli and Dr. C. Ted Lee for serving on my dissertation committee and more. Dr. Povinelli has provided me with the valuable opportunity to collaborate with her group during most of my time at USC. Through this collaboration, I have worked with her students: Dr. Roshni Biswas, Shao-Hua (Nick) Wu, Dr. Mehmet Solmaz, and Dr. Camilo Mejia. With Dr. Lee, I worked with nanoparticles while assisting him in teaching an undergraduate class; this knowledge is directly relevant to my postdoctoral appointment beginning in 2016. I would also like to thank my other collaborators: Dr. Ralf Langen, Dr. Natalie C. Kegulian, Dr. Gerard C. L. Wong, and Ernest Y. Lee. I am grateful to every staff member of the Ronald Tutor Hall Business Center, the Mork Family Department of Chemical Engineering and Materials Science, and the Viterbi School of Engineering who had provided me with assistance at USC. Dr. Malmstadt’s lab has been like another home for me and all members have been wonderful colleagues and friends: Dr. Su Li, Dr. Yasaman Dayani, Peichi Hu, ii Astro Yang, Dr. Carson T. Riche, Gary Newsom, Dr. Kristina A. Runas, Dr. James R. Thompson, Krisna C. Bhargava, Dr. Jesper S. Hansen, M. Gertrude Gutierrez, Bryant Thompson, Dr. Celine Billerit, Lu Wang, and Joshua Moore. I am also thankful for the efforts of my mentees, namely Shuyang Wu, Rebeca Thweatt, Jocelynda Salvador, and Robert Young. The USC Thai scholars have been my family in Los Angeles, and many have provided an essential foundation of support during my residence here. I have exceptional appreciation for Pattaramon Vuttipittayamongkol and Sucha Supittayapornpong for their friendship. I would also like to thank Chalayuth Aungmanee for his never-ending care and support, despite distance and time differences. Most importantly, I owe my successes to my family for their lifelong commitment to helping me achieve my goals. All of my accomplishments were made possible because my father, mother, stepfather, and sister have kept me moving forward. They have all been very supportive of my decisions and were the dependable constants through every change in my life. I am truly grateful. iii Contents Acknowledgements ............................................................................................................. i List of Figures .................................................................................................................... v Abstract ........................................................................................................................... viii Chapter 1: Molecular Perspectives in the Etiology and Pathogenesis of Lipid Oxidation-Associated Diseases ......................................................................................... 1 1.1. Introduction ............................................................................................................. 1 1.2. Reactions that lead to oxidative stress .................................................................. 2 1.2.1. Reactive oxygen species ..................................................................................... 2 1.2.2. Oxidation of lipids .............................................................................................. 3 1.2.3. Oxidation of proteins .......................................................................................... 5 1.2.4. Oxidation of nucleic acids .................................................................................. 8 1.3. Oxidation-associated diseases ................................................................................ 9 1.3.1. Atherosclerosis ................................................................................................... 9 1.3.2. Alzheimer’s disease .......................................................................................... 12 1.3.3. Age-related macular degeneration .................................................................... 14 1.4. Minimizing oxidative stress .................................................................................. 17 Chapter 2: Oxidation of Bilayer-Forming Lipids ........................................................ 18 2.1. Introduction ........................................................................................................... 18 2.2. Experimental ......................................................................................................... 21 2.2.1. Materials ........................................................................................................... 21 2.2.2. GUV preparations ............................................................................................. 22 2.2.3. Basic photooxidation experiments .................................................................... 23 2.2.4. Oxidation while stretching in a dual-beam optical trap (DBOT) ..................... 24 2.2.5. Chemical analysis of oxidized lipids ................................................................ 26 2.2.6. Data analysis ..................................................................................................... 26 2.3. Results and discussion .......................................................................................... 29 2.3.1. Time course of GUV deformation .................................................................... 29 2.3.2. DBOT stretching results ................................................................................... 34 2.3.3. Modeling oxidation kinetics ............................................................................. 36 2.3.4. Line tension of oxidized membrane pores ........................................................ 44 2.3.5. Controlling for oxidation with sodium azide and Trolox ................................. 46 2.3.6. Identifying oxidation products .......................................................................... 48 2.4. Conclusions ............................................................................................................ 50 Chapter 3: Oxidation of Non-Bilayer Forming Lipids ................................................ 52 3.1. Introduction ........................................................................................................... 52 3.2. Experimental ......................................................................................................... 56 3.2.1. Materials ........................................................................................................... 56 3.2.2. Oxidation of DOPE .......................................................................................... 57 3.2.3. Hydration of lipid films .................................................................................... 57 3.2.4. SAXS experiments ........................................................................................... 58 iv 3.2.5. Chemical analysis of oxidized DOPE .............................................................. 59 3.3. Results and discussions ......................................................................................... 59 3.3.1. Hydration of oxidized DOPE lipid films .......................................................... 59 3.3.2. Analysis of the cubic structure ......................................................................... 65 3.3.3. Characterization of lipid geometry ................................................................... 71 3.4. Conclusions ............................................................................................................ 76 Chapter 4: The Effects of Oxidation on Lipid Membrane Bending Rigidity ............ 78 4.1. Introduction ........................................................................................................... 78 4.2. Experimental ......................................................................................................... 80 4.2.1. Materials ........................................................................................................... 80 4.2.2. GUV preparations ............................................................................................. 80 4.2.3. Micropipette aspiration ..................................................................................... 81 4.2.4. Data analysis ..................................................................................................... 85 4.3. Future Work .......................................................................................................... 87 Conclusions ...................................................................................................................... 88 References ........................................................................................................................ 89 v List of Figures Figure 2.1. The microscopy arrangement ......................................................................... 24 Figure 2.2. The dual-beam optical trap configuration ...................................................... 25 Figure 2.3. Example of original and truncated data set for estimating line tension ......... 27 Figure 2.4. The GUV under minimum power (100 mW), maximum power (250 mW) and during RhDPPE excitation irradiation, and the extracted contour through image processing .......................................................................................... 28 Figure 2.5. Vesicle morphologies and radius profile during GUV oxidation at 0.72 mW rhodamine excitation intensity and 90 mol% DOPC-10 mol% RhDPPE ......... 30 Figure 2.6. Summary of averaged normalized radius and fluorescent intensity profiles ....................................................................................................................... 31 Figure 2.7. The mechanism of photo-induced oxidation .................................................. 32 Figure 2.8. Area storing morphologies during oxidation of a 90 mol% DOPC-10 mol% RhDPPE GUV irradiated at 0.72 mW rhodamine excitation intensity from the beginning of irradiation at 0 s ..................................................................... 34 Figure 2.9. Multiple peaks of GUV area strain from DBOT data .................................... 35 Figure 2.10. Distribution of time of irradiation (237.57±79.80 s) at 0.313 mW (561 nm) to arrive at and the percent increase (7.81±2.75%) at the maximum membrane area increased due to lipid oxidation on the DBOT apparatus ................ 35 Figure 2.11. Kinetic model fitted to data averaged from 20 GUVs of 90 mol% DOPC-10 mol% RhDPPE composition, irradiated at 2.85 mW for 15 minutes ....... 38 Figure 2.12. Area parameters A OX1 and A OX2 as functions of the irradiation intensity (I) and rate constants k 1 and k 2 as functions of I 1/2 , obtained through fitting the kinetic model to vesicle area time-course data averaged from ~20 90 mol% DOPC-10 mol% RhDPPE GUVs .............................................................................. 40 Figure 2.13. Kinetic model fit using k 1 = 0.0997 s -1 and k 1 = 0.0490 s -1 to the data averaged from 20 GUVs of 90 mol%-10 mol% RhDPPE composition, irradiated at 2.85 mW for 15 min .............................................................................................. 42 Figure 2.14. Fitting the natural logarithm of irradiation intensity from the basic oxidation experiment as a linear function of time ..................................................... 43 vi Figure 2.15. The distribution of A OX1 obtained through fitting the kinetic model to the DBOT data of 19 90 mol% DOPC-10 mol% RhDPPE GUVs irradiated at an estimated intensity of 0.02 mW (561 nm) ................................................................. 44 Figure 2.16. Distribution of measured line tension values (pN) for pure DOPC (6.10 ± 2.08 pN, n = 93), 7:1 DOPC/DPhPC (8.89 ± 2.66 pN, n = 54), and 3:1 DOPC/DPhPC (9.95 ± 2.56 pN, n = 42) GUVs, all with 10 mol% RhDPPE, in 1:1 v/v 200 mM sucrose solution/glycerol (μ = 10.3 mPa•s) .................................... 46 Figure 2.17. Oxidation of a 90 mol% DOPC-10 mol% RhDPPE GUV in the presence of 1 mM Trolox outside the vesicles at 0 min, 1 min, and 15 min of excitation light irradiation at 0.72 mW ..................................................................... 47 Figure 2.18. Vesicle adhesion of 90 mol% DOPC-10 mol% RhDPPE GUVs from the addition of 20 mM sodium azide outside the vesicles after 1 min and 15 min of excitation light irradiation at 0.72 mW ................................................................. 48 Figure 2.19. 1 H NMR spectra of oxidized and non-oxidized samples of 90%DOPC- 10%Rhodamine B ..................................................................................................... 49 Figure 2.20. 1 H NMR spectra of oxidized and non-oxidized samples of 90%DOPC- 10%Rhodamine B (0.7-3.45 ppm) ............................................................................ 50 Figure 3.1. Hydrated 24-hour oxidized DOPE lipid ........................................................ 62 Figure 3.2. Hydrated oxidized DOPE lipid (low-magnification) ..................................... 63 Figure 3.3. Comparison of lamellar and non-lamellar phases in water ............................ 64 Figure 3.4. Comparison of lamellar and non-lamellar phases in water (low magnification) ........................................................................................................... 65 Figure 3.5. Hydrated 24-hour oxidized DOPE showing two lattice orientations & two-dimensional fast Fourier transform of hexagonal and square patterns .............. 66 Figure 3.6. Section from Figure 3.1 showing the magnified lattice structure of oxidized DOPE & its two-dimensional fast Fourier transform ................................. 67 Figure 3.7. Cross sections of a three-dimensional image stack of the lattice structure of hydrated oxidized DOPE, spanning 10.1 µm x10.1 µm x 4.42 µm (x-y-z) & two-dimensional fast Fourier transforms at an x-y cross-section .............................. 68 Figure 3.8. SAXS results for oxidized DOPE showing the Im3m phase with a = 14.04 nm and non-oxidized DOPE showing the inverted hexagonal phase ............. 69 vii Figure 3.9. Lipid films of DOPE/lyso-PE mixtures showing lamellar phase structures (low magnification) ................................................................................... 73 Figure 3.10. Hydrated lipid films of 60 mol% DOPE/40 mol% lyso-PE, 5 mol% DOPE/95 mol% lyso-PE, and oxidized DOPE dried at the same lipid concentration (approximately 0.7 mg/mL), 5 min after addition of water ................ 74 Figure 3.11. Estimating the extent of oxidation with 1 H NMR ........................................ 75 Figure 4.1. Schematic of the pressure control system and manometer ............................ 83 Figure 4.2. Micropipette aspiration sample holder ........................................................... 84 viii Abstract Oxidation-associated conditions pose as major threats for the aging population. Beyond certain age milestones, these conditions are routinely screened for in order to be caught at earlier, more manageable stages. Their treatments are currently limited, as the understanding of mechanisms leading from biomolecule oxidation to cellular dysfunction and to manifestation of a disease is largely incomplete. While routine cell activities are usually convoluted with participation from many types of biomolecules, their failures can be unraveled by focusing on oxidative damage on lipid membranes that are the foundation of the cellular architecture. The bilayer membrane compartmentalizes the cell and its organelles and hosts a variety of membrane protein functions. Changes in its properties, such as lateral fluidity and continuity, can result in unsuccessful functional protein complex assembly and in leakage of vital nourishments. The research work presented in the following chapters focus on oxidation of membrane phospholipids. After an introduction with more details on how damages at the molecular level lead to well-studied aging-associated conditions, the dynamics of light sensitization-induced oxidation on a bilayer membrane composed of unsaturated lipids is analyzed, where the progress of lipid oxidation is monitored solely based on the overall membrane area changes. 1 Oxidation results in increased polarity of the formerly hydrophobic tails, which alters the geometry of the molecule. Recognizing that the molecular intrinsic curvature can thus be affected by oxidative damage, the structural changes of a species utilizing such property in supporting bilayer membrane bending is then explored. Finally, the last study aimed ix to quantify the holistic membrane bending rigidity changes that arise from previously discussed structural changes to individual lipid molecules. 1 Chapter 1: Molecular Perspectives in the Etiology and Pathogenesis of Lipid Oxidation-Associated Diseases a 1.1. Introduction Eukaryotic cells require oxygen for metabolic production of energy, 2 but this very element required for survival also accelerates our aging and breakdown. The deleterious effects of oxygen and various reactive species derived from it are necessary, as they directly participate in inflammatory responses to defend the body against external pathogens. 3 This ability to fight against infectious diseases has been favored upon by natural selection from early human civilizations. Individuals that possessed robust immune responses were more prone to survive to produce offspring that most likely shared this trait. In our early years, endogenous antioxidant defenses prevent reactive species from turning against our own cells, but their amount and potency gradually diminish through post-reproductive years. 4 When reactive species frequently overwhelm antioxidants, human tissues are injured enough that chronic diseases result. 5 The most prevalent of these conditions are associated with the parts of the body with high exposure to and/or consumption of oxygen. The circulatory system provides direct transport routes of oxygen throughout the body. The nervous system is also a frequent target of oxidative attack, as it has high oxygen consumption to maintain its especially active metabolism. 6 The various oxidative stress-related diseases are inflicted by damage from several of the same reactive species that are produced through cellular metabolism and subsequent a The review presented in this chapter has been adapted from my research report under the same title, completed on April 29, 2015 for the course Gerontology 510: Physiology of Development and Aging offered by the Davis School of Gerontology at the University of Southern California. 2 reactions. This review therefore begins with oxidative damage at the molecular level, focusing on lipids but also extending to other biomolecules for completeness. The discussions will then expand towards particular medical conditions as motivation for the studies on lipid oxidation presented here. 1.2. Reactions that lead to oxidative stress 1.2.1. Reactive oxygen species The term oxidative damage refers to alteration(s) done to one molecule by another, more reactive molecule that involves both in a reduction-oxidation (redox) coupled reaction. 7 When a molecule is reduced, it gains one or more electrons from the other participating molecule that loses the electrons and is oxidized. Paired electrons are more stable than unpaired electrons. Free radicals are atoms, molecules, or ions that contain unpaired electrons. They are highly reactive and favorably participate in redox reactions, usually to gain or lose electrons to result in only paired electrons. A free radical can react with another free radical, or it can react with a non-free radical, in which one of several scenarios can occur. 6 A radical may add to a non-radical to create a new radical. It can donate or obtain a single electron, in which case the non- radical becomes a radical species. A free radical may also abstract a hydrogen atom (and with it, a single electron for its unpaired electron) from a carbon-hydrogen bond. The allylic hydrogen, which is the hydrogen atom attached to a carbon adjacent to a carbon-carbon double bond, is highly susceptible to abstraction by a free radical. 8 The likelihood for this process to occur increases significantly with the number of double bonds on the molecule. Conjugated double bonds in certain antioxidants, such as carotene and lycopene, make them very efficient in attracting 3 and scavenging reactive species to protect vital biomolecules from oxidative damage. 9,10 Free radicals may arise from exogenous and endogenous sources. 11 Incomplete combustion from exhaust gases and cigarette smokes yields a variety of harmful free radicals. Solar ultraviolet radiation indirectly provides free radicals by splitting a non-radical into free radical species with the high energy it carries. The primary free radicals generated in cells are superoxide (O 2 - ) and nitric oxide (NO ). The former usually results from electron leakage from the electron transport chain at the mitochondrial inner membrane during cellular respiration; 12 the latter is intentionally produced as a cardiovascular signaling molecule and is important to immune responses. 13,14 The danger from superoxide is with its potent capability to form a variety of other free radicals. 6,9 It reacts with nitric oxide to form peroxynitrite (ONOO - ) that becomes peroxynitrous acid (ONOOH) at physiological pH; not only does superoxide deplete the availability of the beneficial nitric oxide but it also initiates a pathway to form powerful oxidizing radicals. Peroxynitrous acid can break into nitrogen dioxide (NO 2 ) and hydroxyl (OH ) radicals. The latter can also be formed through metal-catalyzed cleavage of hydrogen peroxide (H 2 O 2 ), known as the Fenton reaction, 15 and through radiolysis of water with gamma and X- radiations. 16 Hydroxyl radicals are especially malicious as they attack everything around them. 1.2.2. Oxidation of lipids Essentially every biomolecules comprising the cell can be oxidized. 6 Examining from outside in, the first structure exposed to extracellular reactive species is the 4 plasma membrane that composes the boundary of a cell with a bilayer of lipid molecules. Phospholipids are a major class of lipids that comprise the plasma membrane bilayer and other cellular membranes forming boundaries of organelles. A phospholipid is amphiphilic and contains a hydrophilic head group and two hydrophobic hydrocarbon tails. Phospholipids self-assemble into the bilayer sheet such that the latter “water-hating” group is shielded from the aqueous environment. 17 The lipid bilayer acts as a barrier to maintain concentration gradients of ions for cell vitality; unintentional leakages can lead to osmotic swelling and cell death through lysis. The bilayer also serves as a platform for activities of membrane proteins such as receptors, enzymes, and ion channels. The two-dimensional lateral fluidity of the bilayer is necessary for assembly of many protein monomers into functional structures and is controlled by the spacing between lipid molecules as governed by their molecular structures and polarity. 18 Oxidation threatens lipid functionality with alterations to molecular structure that controls membrane fluidity, permeability, and topology. 1 Oxidation of a lipid molecule is initiated by the abstraction of a hydrogen atom. 19 An unsaturated lipid molecule has one or more double bonds on its fatty acid moiety, and the allylic hydrogen atoms on the molecule are especially attractive targets to reactive species. For this reason, lipid molecules are more easily oxidized the more double bonds they contain. The removal of a hydrogen atom from a lipid tail results in a lipid radical (LOO ) with the addition of oxygen. This chain of reactions propagates towards oxidation of adjacent lipid molecules, as a lipid radical abstracts a hydrogen atom from its neighbor and becomes a lipid peroxide (LOOH); 5 that neighbor oxidizes yet another neighbor, and so on. This series of lipid oxidation concludes when two radicals react with each other. Metal ions, of iron and copper, accelerate lipid oxidation in two ways. 6 As previously mentioned, they catalyze the conversion of hydrogen peroxide into hydroxyl radical with breakage of the oxygen- oxygen bond. Another way that they perform their destructive role is by turning lipid peroxides back into lipid radicals to sustain the chain of lipid peroxidation reactions. A lipid peroxide molecule is also reactive on its own and can be further oxidized to break the fatty acid moiety into fragments capped with an alcohol (hydroxyl, -OH) group and/or an aldehyde group (-C=O). In general, oxidation leaves hydrocarbon tail(s) decorated with a polar group (hydroperoxyl, hydroxyl, or aldehyde) and causes the formerly hydrophobic moiety to migrate towards the lipid-water interface. As a result, there is usually an increase in the area of the hydrophilic region and a decrease in that of the hydrophobic membrane core, drastically changing the lipid molecular structure and overall landscape of the bilayer. Oxidized membranes tend to have altered flexibility and increased permeability. 1 1.2.3. Oxidation of proteins Membrane proteins are also vulnerable to oxidative damages that affect lipid molecules. These proteins are a means through which the cell senses and interacts with its environment, receiving outside signals and regulating intracellular conditions. 20 In general, proteins serve many vital roles for the cell and for the body. They are important in the repair and maintenance of the cellular construction, from fixing DNA sequences to supporting structural integrity and controlling cellular movements and shape shifts. Enzymes are proteins that regulate chemical reactions 6 to the rates at which are needed. There are proteins that function inside the cell and many that serve their roles in the extracellular matrix. For the multitudes of purposes proteins provide, conformation is the principal determinant of functionality and is studied at four different levels. The primary structure of proteins is the sequence in how its monomers (amino acids) are arranged. The polarity of amino acids determines where local segments of the protein chain can form hydrogen bonds with each other to form a three-dimensional secondary structure such as alpha-helices and beta-sheets. Amino acid side chains of a single protein molecule can also interact, as in the case of sulfur-containing moieties forming a disulfide bridge, to determine tertiary structure domains. Many single-chain subunits can assemble, described as the quaternary structure, to form a fully functional complex. As in the case of lipids, oxidation of proteins threatens functionality with modifications to conformation. An amino acid contains a carbon center that is connected to a hydrogen atom, an amine group (-NH 2 ), a carboxylic acid (-COOH), and a side-chain specific to each amino acid. Amino acids are linearly jointed with the condensation reaction (producing water as a byproduct) between the carboxylic acid and amine groups that results in a peptide bond. The backbone of the amino acid chains can be oxidized in a similar manner as in lipid peroxidation. 16 First, the abstraction of a hydrogen atom, usually by a hydroxyl free radical, from the amino acid center carbon initiates the formation of a carbon-centered radical. The end result may be the original amino acid with the hydrogen atom replaced by a hydroxyl group, a protein-protein linked derivative from two carbon-centered radicals, or a peptide bond cleavage resulting in protein fragmentation. Many amino acid side- 7 chains are also prone to oxidative attacks. Sulfur-containing residues such as cysteine and methionine are particularly sensitive to oxidation by nearly all forms of reactive oxygen species, into disulfide and methionine sulfoxide (MeSOX), respectively. They are particularly vulnerable to attacks by peroxynitrite. However, these are probably the only known oxidative modification of proteins that can be repaired, by disulfide reductase and MeSOX reductase, respectively. Aromatic (benzene ring-containing) amino acid residues such as tryptophan, phenylalanine, and histidine are also targeted by reactive species, and oxidation of glutamine, aspartame, and proline can result in protein fragmentation. Lastly, oxidation of lysine, arginine, proline, and threonine results in carbonyl (-C=O) functional groups. 16 Proteins and aminophospholipids (phospholipids with an amine group in the hydrophilic moiety) can participate in the glycation process. 21 Here, the amine group (-NH 2 ) can react with an aldehyde in a condensation reaction to form an imine (- N=C-) called a Schiff base. This process is well studied in the food sciences field as the Maillard reaction or oxidative browning. 22 The aldehyde groups may be available as the reduced (open-chained) form of glucose, as well as from oxidized lipids and proteins. The imine group has a nitrogen-carbon double bond, and Schiff bases are fluorescent molecules that absorb light in the ultraviolet range. This results in a twofold of damages. First, the actual formation of the Schiff base results in a bridge between two molecules to produce one with a different structure and size from its precursors. Second, an electron excited during light absorption of fluorophores can transfer its high energy into a nearby oxygen molecule to create singlet oxygen ( 1 O 2 ), 8 a reactive oxygen species. 23 Unlike free radicals, singlet oxygen has paired electrons that spin in the same direction and so are also unstable. Singlet oxygen is commonly associated with lipid peroxidation. 1.2.4. Oxidation of nucleic acids While all biomolecules are susceptible to attacks by reactive oxygen species, oxidation of nucleic acids is more prevalent if the reactive oxygen species originate from inside the cell. 24 Nucleic acids include both DNA and RNA and consist of a sugar-phosphate backbone that connects to a nitrogen-base nucleotide that distinguishes each monomer base. The nitrogen bases in DNA are thymine (uracil in RNA), cytosine, guanine, and adenine. The first two are pyrimidines (6- carbon/nitrogen ring); guanine and adenine are purines, which contains a pyrimidine ring attached to an imidazole (5-carbon/nitrogen) ring. All nucleotides are susceptible to oxidation, but guanine has the most vulnerable site for damage: the C- 8 position in between two nitrogen atoms and not involved in the guanine-cytosine (G-C) base pair bond is easily exposed for an attack. 24 Adenine, which is the other purine, can be oxidized at the C-2 and C-8 locations, although not as readily. Pyrimidines can be oxidized at the C-5 position. 25 Just as with lipids and proteins, oxidation of nucleotides begins with the abstraction of a proton and ends with its replacement by the hydroxyl (-OH) group that can reversibly form a tautomeric isomer as a carboxyl group (-C=O) with an adjacent double bond. DNA oxidation occurs daily and this damage is repaired by removal and replacement of faulty bases, resulting in freed oxidized moieties (e.g. 8-hydroxy-guanine or 8-OH-G) within the cell. 26 9 Oxidized guanine that is not readily repaired is a common cause for somatic mutations, 26 which are mutations that can be passed to daughter cells formed through mitosis. 8-OH-G occasionally rotates on the sugar-phosphate backbone of DNA such that N-7 (the nitrogen atom next to the C-8 position where oxidation occurred) is turned towards the base-pairing side that makes the nucleotide appear as a thymine. During replication, it is then paired with adenine (that typically pairs with thymine), and then a new thymine monomer is matched with that adenine. Such mistake can be corrected, but cells that have dysfunctional DNA repair mechanisms can become cancerous. Oxidation of RNA guanine also leads to base pair mismatch that can get translated into abnormal proteins that are highly associated with neurodegeneration. 27,28 Accumulations of 8-OH-dG and 8-OH-G are associated with aging and with many conditions related to oxidative stress such as diabetes, atherosclerosis, cancer, inflammatory diseases, and neurological diseases. 24 1.3. Oxidation-associated diseases 1.3.1. Atherosclerosis Vascular diseases are the most prevalent causes of mortality in almost all parts of the world. 11 Factors contributing to these chronic inflammation disorders of the vasculatures include hyperlipidemia, hypertension, diabetes, obesity, infection, and smoking, most of which increase risks by promoting oxidative stress. Arteries are easily targeted by oxidative stress due to several factors. Nitric oxide is abundantly present as a vasodilator, but it is also a precursor for many reactive species that can initiate oxidation. Carried through the arteries are oxygenated red blood cells that provide oxygen and oxidation-promoting iron. 29 During an inflammatory response, 10 oxidative stress is heavily enhanced with leukocytic activity. Lipid abnormalities are the key feature in atherosclerosis, 30 where accumulation of plaques may limit or obstruct blood flow through an artery. Oxidized unsaturated, low-density lipids (LDLs) accumulate and further induce inflammatory responses inside the vessel wall. Inflammation also increases the number of reactive species that further promotes oxidation product buildups and can ultimately result in complete blockages that prevents oxygen delivery to vital tissues. This subsequently induces ischemic episodes such as acute coronary syndrome and cerebral infarction. The arterial wall consists of three layers. 31 The adventitia is the outermost layer that consists of fibroblasts, mast cells, microvessels, lymphatic vessels, and nerves, and it covers the media. The media contains layers of smooth muscle cells that provide mechanical support for the artery carrying blood pumped from the heart and contract to regulate blood pressure and flow to arterioles (connecting to capillaries). The intima is a single layer of endothelial cells that secrete a layer of the extracellular matrix (ECM) called the basal lamina, on which it rests. Intima endothelial cells are in direct contact with the arterial lumen and are the first to be affected by perturbations in the compositions of blood and interstitial fluids, such as hyperlipidemia, hyperglycemia, and inflammation. At elevated plasma LDL levels, the high concentration gradient drives lipids to cross the endothelial layer and become trapped in the protein-rich basal lamina and the ECM of the intima. Lipids and proteins can oxidize and proliferate to form an atheroma that disrupts cellular junctions between the endothelial and muscle cells and lead to endothelial cell dysfunction and stimulation of inflammatory responses. 11 Under normal conditions, the endothelial layer that is in contact with flowing blood resists adhesion of leukocytes. During an acute inflammatory response, however, stimulated endothelium cells increase expression of adhesive molecules to attract circulating white blood cells. 32 This induces localized production of monocytes, neutrophils, and lymphocytes that generate reactive oxygen species to combat against invading pathogens. Polyunsaturated fatty acids belonging to the host, here trapped in the intima, are also attractive targets for oxidative modification that can result in the formation of an aldehyde group. The aldehyde group then combines with an amine group of a protein or lipid head group that results in linkage of these molecules. In fact, the resulting Schiff bases formed through such reaction are stable markers of oxidative damage to tissues as they can be detected through their fluorescence. 33 Lipid oxidation also results in scission of the hydrocarbon tail that becomes more polar and moves to the water-lipid interface. Truncated phospholipid molecules are very similar in structure to platelet-activating factor that induces further inflammatory responses and platelet aggregation for clotting. 32 During this robust inflammatory reaction, monocytes in the intima differentiate into macrophages that later form foam cells that characterize an early atherosclerotic lesion. 34 Macrophages proliferate within the intima and secrete growth factors and cytokines that further the development of such lesion. Evolvement into complicated atheroma with fibrous plaque formation occurs with interactions with smooth muscle cells that are stimulated by the growth factor secreted by macrophages. Plaques are separated from the arterial lumen by a fibrous cap that is structurally supported by collagen. 12 Atherosclerosis becomes life threatening with plaque disruption. While the specific mechanism is unknown, it is related to the thinning of the fibrous cap due to excess inflammatory cytokines. Certain cytokines can inhibit smooth muscle production of collagen that provides mechanical integrity for the fibrous cap. Oxidative stress can also lead to over-expression of collagenase that accelerates collagen degradation. 34 Upon plaque rupture, blood comes into contact with atheroma contents. Oxidized lipid products from the lesion lyse red blood cells, eventually leading to the release of iron that promotes further oxidation reactions. Platelets play the major role in coagulation in late-stage atherosclerosis; they adhere to the ECM that is rich in pro-inflammatory factors, aggregate, and may lead to complete blockage of the artery and subsequent ischemia. 31 1.3.2. Alzheimer’s disease Many neurodegenerative diseases are associated with oxidative stress. The nervous system is especially sensitive to damage from reactive species for many reasons. 6 Principally, neurons are the site of high metabolic conversions for energy to regulate ion channels for propagation of the action potential in cell-to-cell communication. High oxygen consumption results in increased probability of reactive oxygen species formation. Some neurotransmitters such as dopamine, serotonin, and norepinephrine add to the reactive species population by reacting with oxygen to form superoxide. Many proteins contain oxidation-promoting iron; iron is also released when hemoglobin from blood is degraded upon exposure to excess hydrogen peroxide from metabolism of the nervous system. The neuronal membrane is also highly susceptible to oxidative damage because its lipids are rich in polyunsaturated tail 13 groups. Arachidonic acid and docosahexaenoic acid (DHA) have four and six double bonds, respectively, and account for 20% for total fatty acid contents in the brain. 35 The neurodegenerative diseases most studied for their associations with oxidative stress are Alzheimer and Parkinson’s diseases. 36,37 As of 2010, more than 35 million people worldwide (5.5 million in the US) have Alzheimer’s disease. 36 The condition can lead to short-term memory loss, dementia, and impairment in language, visuospatial function, and calculations. Accumulation of misfolded protein due to oxidative and inflammatory damage is characteristic in Alzheimer’s disease; other representations are inadequate neuronal energy production and synaptic dysfunction. Plaques of beta-amyloid peptide are an important feature in Alzheimer’s disease. The beta-amyloid peptide is a natural product of metabolism, but an imbalance between its production and clearance leads to the accumulation that drives their aggregation into oligomers and fibrils. Oligomers consist of a few coalesced peptide monomers. Amyloid fibrils are insoluble fibers that make up advanced plaques; however, the soluble, intermediate amyloids (dimers and trimers) are the more neurotoxic forms as they interfere with synapses. Despite the focus on beta-amyloid agglomerations in the etiology and pathogenesis of Alzheimer’s disease, the true underlying cause may be due to abnormalities in tau proteins that are the pathologic marker for severity of the disease. Tau proteins are responsible for the removal of beta-amyloid peptides, so without proper function of tau proteins, beta-amyloid peptides accumulate into plaques. Tau proteins are abundant in neuronal axons for formation of cytoskeleton 14 microtubules that support cellular structure. It was found that abnormally phosphorylated and glycated tau does not bind to microtubules and compromises neuronal structure. 38 Overall, oxidative stress leads to impaired protein functions and accumulations that affect cell-to-cell communication. Oxidation of the lipid membrane increases ion permeability leading to concentration imbalances and hindered action potential efficiency. Furthermore, leaky membranes lead to impaired glucose transport to fuel cellular respiration. Alzheimer’s disease seems to be primarily a synaptic failure disorder. 36 Approximately 25% of mild cases had reduction in the presynaptic protein synaptophysin. With progression of the disease, synapses decline more quickly than the number of neurons, leading to dementia. Late stage Alzheimer’s disease patients experience severe reduction of the growth factor protein neurotrophin that aids function and survival of neurons to moderate learning, memory, and behavior. 1.3.3. Age-related macular degeneration Polyunsaturated lipids are highly susceptible to oxidative damage, and they are present in particularly high concentrations in neuronal membranes. They have an important role in modulating membrane curvatures during synaptic cell-to-cell communication with neurotransmitter transfers. Another area with the need for an abundance of polyunsaturated lipids is in the membranes of photoreceptor (rod and cone) cells. To maximize the surface area-to-volume ratio, rods contain approximately 1000 membrane disks and cones have approximately 700 times folded membranes; 39 both kinds are decorated with rhodopsin, the light-sensing protein. As 15 introduced earlier, oxidative stress can arise from light-induced toxic effects in the formation of the reactive oxygen species singlet oxygen. Excessive exposure to light (particularly those with shorter wavelengths, carrying more energy) can lead to visual impairment conditions such as age-related macular degeneration (AMD) and cataracts. For this reason, lenses inserted after cataract extraction may have filters to remove ultraviolet and blue light. The central cause of AMD is oxidative stress, so while late-stage AMD is the most common cause of untreatable blindness in the Western world, its progress can be slowed with antioxidant-rich diet. The characteristic physical sign of AMD is the formation of drusens, which can be observed with an ophthalmoscope as white- yellow dots when they become larger than 25 µm. Drusens are present in early-stage AMD when visual impairment is not clearly presented; the larger the drusens are, the greater risk there is of development into late-stage AMD. Late-stage AMD presents with severe loss of vision and is highly associated with old age. The risk of developing it increases from 0.05% before 50 years to 11.8% after 80 years of age, likely due to lifetime accumulation of oxidative product debris. 39 The retinal pigment epithelium (RPE) is the central component in the pathogenesis of AMD. 39 It is a single layer of cells with very high metabolic rates, on which rod and cone cells rest. The tips of rods and cones with rhodopsin already exposed to light are shed from the photoreceptor cells and engulfed and degraded in RPE phagosomes. RPE supplies photoreceptor cells with nucleic acids, proteins, and lipids to balance this shedding, and also transports nourishing fluids and ions from a layer of capillaries called the choriocapillaris. 16 The high amount of lipids and proteins to be digested in the RPE are often oxidized into cross-linked forms. Mitochondrial metabolism supplies free radicals through electron leakage that then poses further oxidative stress on the mitochondria and leads to its degradation by autophagic endosomes and lysosomes. Polyunsaturated lipids can oxidize into products such as malondialdehyde (MDA) and 2-hydroxy-4-trans-nonanal (HNE) that are especially prone to cross-link with proteins. 11 Once such cross-links are formed, they cannot be degraded with the enzyme hydrolase; they also provide “initiation cores” for growth of lipofuscin clusters that accumulate in lysosomes and are expelled into the RPE cytosol after a critical concentration is reached 40 . Lipofuscins contain several fluorescent Schiff bases. One of the very few identified lipofuscin fluorophore is bis-retinoid N- retinylidene-N-retinylethanolamine (A2E), 41 which is synthesized from all-trans retinal forming a dimer with phospholipids containing an amine group. The photoxicity of A2E results in damages to DNA and leads to apoptosis of RPE cells. Lipofuscins accumulate from the age of 18 months. While it is unknown how drusens develop, they probably result from accumulations of numerous oxidized materials such as lipofuscins. Drusens contain glycoprotein cores and outer layers of proteins related to inflammation and RPE cell fragments. They appear from the age of 30 years, situated between the basement membranes of RPE and a collagenous layer of Bruch’s membrane. Bruch’s membrane consists of three layers (as a central thin, elastic layer sandwiched between two collagenous layers) and rests in between RPE basement membranes and the choriocapillaris. Initially, drusens at 30 years old is lipid-free, but they thicken with accumulations of collagen, lipids, and other debris 17 from then. The calcification and thickening of drusens limit fluid permeability and transport of nutrients from the choriocapillaris. This leads to the thinning and destruction of the RPE with increasing age and plays a central role in the development of AMD. 1.4. Minimizing oxidative stress Oxidative stress poses great danger towards cellular building blocks, and the risks increase with age as endogenous defense mechanisms subside. Maintaining healthy weight, blood pressure, blood sugar, and fat contents by consumption of antioxidants, green leafy vegetables, whole grain, fish, and nuts all contribute to lowered risks of oxidative damage. 42 Smoking is the recurring cause of increased risks for atherosclerosis, Alzheimer’s disease, and age-related macular degeneration, as well as for other oxidation-associated diseases. Antioxidant supplements such as vitamin C, vitamin E, beta-carotene, lycopenes, zinc oxide, and cupric oxide can reduce progression of diseases over a period of time and are useful for managing early stages of oxidation-associated conditions. While factors such as genetics and past environmental exposures cannot be prevented from predisposing one to oxidation-associated diseases, lifestyle and dietary modifications can slow down lifetime accumulation of damages that could make a difference between gradually declining health and rapid transition into disability. 18 Chapter 2: Oxidation of Bilayer-Forming Lipids b 2.1. Introduction Lipid bilayer membranes are the structural barriers that contain the cell and its compartments. They serve as the platforms on which membrane proteins are localized and play a central role in a host of physiological processes. The ability of lipid bilayers to perform their physiological role depends on the integrity of the membrane structure. 43 Lipid oxidation causes modification and/or loss of essential membrane functions 44 and has been identified in pathological conditions such as cancer 45,46 and aging-associated conditions such as atherosclerosis 47 and Alzheimer’s disease 48 . Despite the strong connection of membrane oxidation to human health, the specific molecular mechanism connecting lipid oxidation to the membrane’s roles in disease etiology and pathogenesis is not well understood. Here, we use a light- induced model of lipid oxidation to probe the kinetics of lipid oxidation by observing oxidation-linked changes to the morphology of giant unilamellar lipid vesicles (GUVs). b The work presented in this chapter has been published in Biochimica et Biophysica Acta (BBA) – Biomembranes under the title “The dynamics of giant unilamellar vesicle oxidation probed by morphological transitions” and used with permission under license number 3730370617426, obtained on 15 October 2015. This work was authored by Shalene Sankhagowit, Shao-Hua Wu, Roshni Biswas, Carson T. Riche, Michelle L. Povinelli, and Noah Malmstadt. S.S. and N.M. designed the study. S.S. designed and performed the optical microscopy experiments. S.S., S.-H.W., and R.B. designed and performed the DBOT experiments. C.T.R. designed and performed the 1 H NMR experiments. M.L.P. contributed the tools for the DBOT experiments. S.S., R.B., and N.M. wrote the paper. All authors reviewed the results and approved the final version of the manuscript. 19 The key oxidation process that occurs within the lipid bilayer involves oxidative species such as reactive oxygen species (ROS) attacking unsaturated lipids. The products of the oxidation reactions depend on the type of lipids involved (including mono- and polyunsaturated) and the particular oxidative species mediating the attack. 49 However, the most commonly observed fundamental process can be viewed as a series of key chemical events. First, the oxidation of an unsaturated lipid molecule is initiated by the abstraction of the allylic hydrogen adjacent to the double bond and the reaction with molecular oxygen to form a carbon-centered peroxyl radical. This in turns initiates oxidation of neighboring lipid molecules, and the lipid tails are left modified with a hydroperoxy group, 19,50 and further oxidation leads to lipid tail scission into a truncated lipid molecule and tail fragment. 51 Singlet oxygen ( 1 O 2 ) is an ROS frequently encountered during visualization of the lipid membrane by fluorescence microscopy. When irradiated, photosensitive molecules such as porphyrin derivatives and fluorescent rhodamine dyes can transfer their energy to molecular oxygen (O 2 ) to generate the more reactive 1 O 2 . 52,53 Biomedically, single oxygen production by photosensitization is utilized by photodynamic therapy (PDT) for targeted destruction of malignant tissues. 54 Singlet oxygen is also naturally-occurring, resulting when UVA rays photosensitize endogenous porphyrins. 55 Singlet oxygen-induced effects have been noted to alter membrane phase behavior. 56,57 Other reports have noted more drastic changes to the lipid membrane morphology induced by singlet oxygen-related reactions. For the general case of 20 phospholipid membrane oxidation, molecular dynamics simulations of a 1,2- dioleoyl-sn-glycero-3-phosphocholine (DOPC) membrane reveal membrane bending and pore formation within nanoseconds of oxidation, especially with scission of both acyl tails. 58 Investigations adding photosensitizers to 1-palmitoyl-2-oleoyl-sn- glycero-3-phosphocholine (POPC) and DOPC GUVs reported changes to the membrane bending modulus 59 and area expansion modulus 60 with lipid oxidation. Others observed increases in the membrane surface area accompanied by vesicle shape fluctuations when irradiating porphyrin-labeled pure POPC or POPC- containing GUVs. 57,61 In addition, a study using DOPC GUVs in different concentrations of methylene blue solution also observed post-expansion membrane area contraction accompanied by loss of optical contrast across the membrane; this was interpreted as the result of pore formation. 62 Similar effects have also been reported in polymersomes composed of polyethyleneoxide-b-polybutadiene (PEO-b- PBD) diblock copolymer with the chromophore chlorin e6, where the vesicles grew larger over three minutes but also shrunk afterwards. 63 A baseline understanding of the dynamics of lipid oxidation is of broad utility. Rhodamine-based dyes are broadly used in imaging studies; understanding the precise conditions under which they can be expected to lead to oxidative damage—and potential artifactual results—is therefore important. Rhodamine dyes have also been considered as potential photosensitizers in photodynamic therapy (PDT). 52,64 Understanding the kinetics of oxidation produced by the photosensitizer is key to designing such therapies, and the framework of our kinetic model is extensible beyond rhodamine fluorophores to other photosensitizers, such as 21 porphyrin chromophores. 65 Despite many observations of light-induced oxidation of the lipid membrane, the dynamics of the underlying chemical events is largely unexplored, so the main goal of this study is to provide a kinetic model applicable to the complex process of oxidation while retaining simplicity by constraining to rate- limiting processes. Based on the morphological changes observed by others 57,61-63 and in this study, the process is apparently a two-step process involving membrane area expansion followed by contraction. Bateman and Gee 66 described the rate of light-induced oxidation of non-conjugated olefins to be first-order in which the effective rate constant varies proportionally with light absorption, which directed the focus of our study towards the dependence of oxidation dynamics on the intensity of the excitation light. 2.2. Experimental 2.2.1. Materials The lipids 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-dipalmitoyl-sn- glycero-3-phosphoethanolamine (DPPE), 1,2-diphytanoyl-sn-glycero-3- phosphocholine (DPhPC), and 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine- N-(lissamine rhodamine B sulfonyl) (RhDPPE) were purchased from Avanti Polar Lipids. Sucrose, glycerol, 6-hydroxy-2,5,7,8-tetrametylchromane-2-carboxylic acid (Trolox), and 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) were purchased from Sigma-Aldrich. Glucose and rhodamine B were purchased from Alfa Aesar, chloroform was from Macron Fine Chemicals, and sodium azide was from BDH Chemicals. Deuterated chloroform was from Cambridge Isotope Laboratories, Inc. 22 2.2.2. GUV preparations Our standard GUV composition consisted of 9:1 molar ratios of DOPC to RhDPPE. In reducing DOPC concentrations to 85 and 75%, the unsaturated lipid was substituted with DPhPC such that RhDPPE was maintained at 10% of the total composition; DPhPC was selected as a substitute that would have a fluidity and phase behavior similar to that of an unsaturated lipid without susceptibility to oxidation. To investigate rhodamine content dependence, RhDPPE was substituted with unlabeled DPPE. The GUVs were formed using the electroformation method pioneered by Angelova and coworkers. 67 A lipid solution (in chloroform) was deposited on an indium-tin oxide (ITO)-coated side of a glass slide (Delta Technologies), inside the perimeter enclosed by a silicone o-ring (13/16” ID, 1” OD, Sterling Seal & Supply) attached to the slide by silicone vacuum grease (Dow Corning). After drying under vacuum overnight, the lipid film was hydrated with a 200 mM sucrose solution in 10 mM HEPES buffer at pH 7.40 such that the final lipid concentration was 30-40 µg/mL. The electroformation apparatus was completed by attaching another ITO- coated slide to the o-ring with vacuum grease, with the conductive side facing towards the lipid film. Electrodes connecting each of the ITO-coated slides to a function generator (Hewlett-Packard/Agilent Technologies) allowed for the application of an AC field at 10 Hz and 1.3 V for 1 hour at room temperature. Vesicles were used within a day of electroformation. Lipid mixtures containing rhodamine dye were always shielded from ambient light to avoid photobleaching and other unintended oxidation effects. 23 For dual-beam optical trap (DBOT) experiments, which require a higher density of GUVs in solution, GUVs were formed by hydrating the lipid film dried on a layer of agarose hydrogel. 68-70 The GUVs were formed in 500 mM sucrose solution in 20 mM HEPES buffer at pH 7.00, a condition used in previous work with a DBOT. 71,72 2.2.3. Basic photooxidation experiments Electroformed RhDPPE-labeled GUVs were observed with epifluorescence microscopy on a Nikon TI-E inverted microscope, using illumination filtered through a green excitation filter (528-553 nm bandpass, 540 nm cut-on wavelength). Excitation light was provided by a 130W mercury lamp (Intensilight, Nikon). The maximum irradiation intensity through the objective (Apo TIRF 60X Oil/NA 1.49, Nikon) was measured by a laser power meter (Thorlabs, Inc.) to be 5.93 mW at 561 nm. Intensity was reduced in binary ratios using neutral density filters. We utilized asymmetry in aqueous solutions to facilitate microscopy observations: the GUVs formed in sucrose were transferred to an isoosmotic glucose solution (200 mM in 10 mM HEPES at pH 7.40). Since sucrose solutions are denser than glucose solutions at the same concentration, the GUVs sedimented to rest on the glass coverslip, minimizing their distances to the objective and their mobility. Figure 2.1 illustrates the described arrangement. 24 Figure 2.1. The microscopy arrangement. To estimate line tension in the pore-forming regime of DOPC GUV oxidation, pore closure was delayed by performing the experiments in ten-fold elevated aqueous viscosity. Both 200 mM glucose and sucrose solutions were formed in a 1:1 v/v glycerol-water mixture. The viscosity of the resulting sucrose solution was measured with an Ubbelohde viscometer (Cannon Instrument Company) to be 10.3 mPa•s. 2.2.4. Oxidation while stretching in a dual-beam optical trap (DBOT) Rapid GUV fluctuation in early stages of oxidation made it difficult to accurately measure membrane surface area. For accurate area measurements, we repeated a set of the oxidation experiments while GUVs were captured and stretched in a dual- beam optical trap (DBOT). In a DBOT, counter-propagating laser beams emitted from two single mode optical fibers form a trap at the geometric center of the two fiber cores, owing to the Gaussian intensity profile of the beams, 73,74 as illustrated in Figure 2.2. In our apparatus, described in detail elsewhere, 72 the optical fibers are Glass Coverslip Microscope Objective Glucose Sucrose GUV Irradiation Source Fluorescence Emission 25 placed on grooves etched on silicon which ensure the fiber core alignment. Perpendicular to the fiber grooves runs another groove that houses the microfluidic glass channel. The GUVs were flowed through this glass channel using a peristaltic pump and were brought close to the trapping region. The vesicles were initially trapped using a minimal laser power (50 mW from each fiber operating at a wavelength of 808 nm). At this point, we started capturing images of a GUV at the equatorial plane, using a CCD camera, through a 50X (NA 0.55, Nikon) objective at a frame rate of 62 fps. The rhodamine excitation light was emitted through the same objective, using the same green excitation filter described above but with a weaker illumination source (Nikon TE2-PS100W), in order to initiate the oxidation process. Maintaining the GUV in the trap, we then increased the laser power to 250 mW (from each laser) and caused the GUV membrane to stretch. The GUV was kept stretched for 5 s and the laser power then dropped to zero. Keeping the illumination on, the stretching experiment was repeated 15-20 times in order to capture data at several points over the course of oxidation. Figure 2.2. The dual-beam optical trap configuration. (OF = optical fiber) 26 2.2.5. Chemical analysis of oxidized lipids In preparation for nuclear magnetic resonance (NMR) spectroscopy, lipid samples (in chloroform) were dried onto the bottom of a glass vial under an argon stream and placed under vacuum for at least 1 hour. The samples were irradiated with the green excitation light for 1 hour. Samples were redissolved in ~800 µL of deuterated chloroform and scanned on a Varian VNMRS-500 2-channel NMR spectrometer at 25°C. Samples were scanned prior to irradiation and after each of four subsequent cycles of green excitation light exposure. 2.2.6. Data analysis The time-lapse image sequence observed by epifluorescence microscopy for the oxidation of each GUV was recorded by Nikon NIS Elements software in the ND2 file format, which was then imported into MATLAB for data processing using the Bio-Formats m-file package available through the University of Wisconsin-Madison Laboratory for Optical and Computational Instrumentation (LOCI). The GUV edge was traced using built-in MATLAB functions to apply a Gaussian filter and Canny edge detection. For images in which pores were evident, the portion of the GUV perimeter occupied by the pore was traced by fitting observable edge coordinates to an ellipse. With knowledge of the coordinate of the vesicle center and the lengths of the major and minor axes, the angle of the arc from the dark void in the membrane was then used to calculate the arc chord corresponding to pore diameter. At the quasi-static leak-out limit where pore radius and surface tension are assumed constant with respect to time, Karatekin and coworkers simplified the 27 hydrodynamic model of pore closure from a differential equation to the linear relationship € R 2 lnr =− 2τ 3πη 0 t +C (2.2.6-1) where R and r are the GUV and pore radii respectively and t denotes time. 75 The aqueous viscosity is denoted by η 0 , and C is a constant of integration. By plotting R 2 ln r as a function of time, the measured slope of the linear region was then used to calculate τ, the line tension. Figure 2.3 shows an example of such plot, which reflects a sudden rise at pore opening, the linear decrease characteristic of the quasi-static leak-out regime, and a rapid drop towards pore closure. Figure 2.3. Example of original (squares) and truncated (circles) data set for estimating line tension. In order to reproducibly and objectively select the linear region from the full pore closure data set, an adaptation of a truncation algorithm was used. 76 First, data points preceding the occurrence of the maximum pore radius as well as those 28 showing total pore closure (r = 0) were removed. The remaining pore radii comprise data set A. Another data set consists of points from data set A, minus the end point. If the difference of the root mean squares (RMS) of the residuals from linear fittings of the two sets is equal to or exceeds 10% of the RMS of set A, then the removal of such point was considered to increase linearity. This step was iterated until the relative RMS change was less than 10% and/or there were fewer than four data points remaining. An example of an original and truncated data set is shown in Figure 2.3. In the case of the DBOT experiment, an in-house image-processing algorithm was applied in order to trace the GUV edge from the two-dimensional (2D) micrographs. The calculation of the three-dimensional (3D) vesicle surface area employed a method similar to one described by Milner and Safran. 77 The micrographs such as in Figure 2.4b represent the equatorial cross-section of a stretched GUV having the ellipsoid shape. Figure 2.4. The GUV under (a) minimum power (100 mW), (b) maximum power (250 mW) and during RhDPPE excitation irradiation, and (c) the extracted contour through image processing. Scale bar = 5 µm. With data from the first time-point, the volume of this ellipsoid was approximated from the estimated major and minor axes of the cross-sectional ellipse, (a) (b) (c) 29 based on traced GUV (Figure 2.4c) edge coordinates. The radius of the equivalent sphere (R) of the same volume was then obtained and used in € r θ ( ) =R + 1 2 3cos 2 θ −1 ( ) $ % & ' ( ) u 2 . (2.2.6-2) We assumed conserved volume of the GUV throughout the course of the experiment, so R is constant (note: DBOT experiments were terminated prior to the onset of pore formation). The second term represents the second-order Legendre polynomial associated with the gross shape change of the GUV under stretching tension. The 2- D contours were fitted to Equation 2.2.6-2 to relate the vesicle radius (r) as a function of the azimuthal angle (θ). Assuming the GUVs to be symmetric about the optical beam axis, we performed a solid angle of revolution integral on the fitted contour to obtain the surface area of the GUV. 2.3. Results and discussion 2.3.1. Time course of GUV deformation The light-induced oxidation of DOPC GUVs is apparently a two-stage process: a typical example is shown in Figure 2.5. GUVs are initially spherical and relaxed, with low-amplitude fluctuation observed (Figure 2.5a). Within the first 3 min of irradiation, the vesicle collapses towards the coverslip, resulting in an apparently greatly increased radius, and high-amplitude fluctuations are seen (Figures 2.5b and 2.8b-e). As irradiation continues, membrane tension increases, the GUV returns to a rigid spherical shape (Figure 2.5c), and micron-scale pores begin to regularly form (Figure 2.5d-f). The two phases of decreased tension and collapse followed by increased tension and pore formation can be seen clearly by plotting the apparent 30 average radius of the GUV as a function of time (Figure 2.5g). The radius plotted here represents the average radius of the traced vesicle contour (i.e. distance of the contour from the center point averaged across all angles) obtained by image analysis of epifluorescent micrographs. Note the initial rapid increase (collapse and fluctuations) to a peak radius followed by a decrease (tension increase) that is initially smooth but at long times becomes stepwise. Each step here corresponds to a single pore formation event. The time-course radius changes for other experimental conditions (varying excitation light intensity, DOPC concentration, and RhDPPE concentration) are available in Figure 2.6. Figure 2.5. Vesicle morphologies and radius profile during GUV oxidation at 0.72 mW rhodamine excitation intensity and 90 mol% DOPC-10 mol% RhDPPE. The epifluorescence micrographs in (a)-(f) shows the same GUV (a) at the beginning of irradiation at 0 s, (b) during area expansion at 63 s, and with increased rigidity with (c) continuous membrane at 216 s and with pores at (d) 217 s, (e) 243 s, and (f) 289 s. Scale bars = 10 µm. The radius values normalized to initial GUV dimension over 900 s for another GUV from the same electroformation batch is shown in (g). 0 200 400 600 800 Time / s 0.8 0.9 1 1.1 1.2 Normalized Radius (a) (b) (c) (d) (e) (f) (g) 31 Figure 2.6. Summary of averaged normalized radius and fluorescent intensity profiles. Conditions varied include irradiation intensity (0.19 to 2.85 mW) at 90 mol% DOPC-10 mol% RhDPPE, the concentration of RhDPPE (from 0.5% to 10%, by substituting in DPPE) at 90 mol% DOPC and 0.72 mW irradiation intensity, and the concentration of DOPC (from 67.5% to 90%, by substituting in DPhPE) at 10 mol% RhDPPE and 0.72 mW irradiation intensity. As introduced earlier, oxidation of unsaturated phospholipids result in the addition of the hydroperoxy group adjacent to the double bond on the acyl tail, following allylic hydrogen abstraction and carbon-centered radical formation. 78 The presence of this newly added functional group increases the lateral area in the membrane that the lipid molecule occupies. Collectively, this leads to membrane area increase that we observed and is represented by the data in the 0 to 175 s interval of Figure 2.5g. Further oxidation results in acyl chain scission and loss of 32 lipid material from the membrane, 79,80 resulting in lipid area and membrane area contraction. The described oxidation steps are briefly summarized in Figure 2.7. Figure 2.7. The mechanism of photo-induced oxidation. (a) The allylic hydrogen is abstracted by 1 O 2 and with the addition of molecular oxygen, the peroxyl radical is added in its place. (b) The peroxyl radical can initiate the oxidation of neighboring lipid molecules to result in a hydroperoxy (-OOH) group adjacent to the double bond. (c) Oxidation of the - OOH group results in lipid tail scission into a shortened acyl chain capped with a hydroxyl (- OH) group and a tail fragment capped with the aldehyde (-CHO) group. (d) Further oxidation of -OH on the shortened lipid molecule results in its replacement with -CHO. The kinetic data produced by these experiments are time series of images showing changing morphology. The molecular parameter that changes with progress of oxidation is the area per lipid, so we needed to first measure vesicle surface area from each of these images in order to model the kinetics. In measuring surface area from these two-dimensional micrographs, we utilized image processing in MATLAB for membrane edge detection. The coordinates of the pixels comprising the vesicle edge were averaged to approximate the coordinates of the vesicle center, and the distances from the center to all edge points were then averaged to yield vesicle radius. We assumed that the GUVs were always spherical in calculating vesicle surface area from this radius. In observation of GUVs resting on the coverslip surface, several experimental artifacts persisted and increased the uncertainty of the time course data during the membrane area expansion phase of oxidation when the vesicle morphology O 2 O 2 1 OO . H + . OO . + A A+dA O 2 OOH OH + O O (a) (b) (c) (d) 33 significantly deviated from a spherical shape. With collapsed GUVs, as seen in Figure 2.5b, the measured vesicle dimensions were overestimated, and adding high amplitude fluctuations (Figure 2.8b) meant that the average radius data could not be used to accurately estimate changes in the membrane area that resulted in decreased tension. Furthermore, the GUVs mitigated increased surface area-to-volume ratios by storing excess membrane as tubules and buds, and these were later stretched out as further oxidation resulted in products with decreased surface area. While the morphological transitions of GUVs are more accurately represented by Figure 1, a few which formed tubules and buds to mitigate increased surface area (Figure 2.8) experienced several brief interruptions during the membrane area-decrease phase. First, the surface tension of the collapsed GUVs increased such that the spherical shape was returned (Figure 2.8f). Excess membrane stored away as tubules were stretched back to the main GUV body and the sudden return of membrane area was accompanied by more fluctuations (Figure 2.8g). The time course radius data reflects this competition between the increasing membrane tension above the threshold necessary for spontaneous pore formation and the returning of excess membrane with cyclic rapid increase and step-decreases around 200 s in Figure 2.5g. It should also be noted that the collapse and swell of GUVs occurred rapidly and were not always accurately followed during epifluorescence microscopy so images were sometimes out of focus during these transitions. For the purpose of fitting the kinetic model, the data points representing these behaviors were not considered. 34 Figure 2.8. Area storing morphologies during oxidation of a 90 mol% DOPC-10 mol% RhDPPE GUV irradiated at 0.72 mW rhodamine excitation intensity from (a) the beginning of irradiation at 0 s. The GUV underwent the area expansion phase of oxidation and formed tubules and connected buds from (b) 51 s, (c) 55 s, (d) 61s, and (e) 71 s. During the transition phase, the GUV membrane area was reduced and rigidity increased at (f) 169 s before increased surface tension stretched a tubule and returned excess membrane area at (g) 177 s. The GUV became fully rigid by (h) 207 s. Scale bars = 10 µm. 2.3.2. DBOT stretching results Since the surface area of GUVs collapsed on the glass surface cannot be accurately measured, oxidation experiments were repeated in a DBOT, which stretches the vesicles to allow accurate surface area measurements. We observed both the time at which peak area was achieved and the maximum area increase at this peak. The distributions of peak time and maximum area strain, confirmed to be normal by the Kolmogorov-Smirnov test, are summarized in Figure 2.10. The formation of membrane tubules led to multiple peaks in the time-course measurement of vesicle size (Figure 2.9); the vesicles that exhibit such behavior were excluded from the distribution of the maximum percent area increase (Figure 2.9b). (a) (b) (c) (d) (e) (f) (g) (h) 35 Figure 2.9. Multiple peaks of GUV area strain from DBOT data. Figure 2.10. Distribution of (a) time of irradiation (237.57±79.80 s) at 0.313 mW (561 nm) to arrive at and (b) the percent increase (7.81±2.75%) at the maximum membrane area increased due to lipid oxidation on the DBOT apparatus. The GUV membrane area increased by 7.81 ± 2.75% on the DBOT apparatus. Note that the light source used was significantly weaker than that used in the main setup, as shown by the lengthened time to maximum area increase (Figure 2.9a). Mertins and coworkers reported that the maximum area increase depends on the rate of singlet oxygen formation, 62 which can be controlled by changing photosensitizer concentration and irradiation intensity. The time-course surface area calculations based on data collected with the DBOT apparatus were also used for fitting to the dynamical model described next. Count 4 8 12 0 3 6 9 12 15 Maximum Strain (%) Count 5 10 15 0 80 160 240 320 400 Peak Time (s) (a)! (b) 36 2.3.3. Modeling oxidation kinetics Based on the two distinct stages of oxidation observed, we modeled DOPC oxidation as having two rate-limiting steps in series; each step was simplified to an irreversible reaction with one collective species each for reactant and product. The product of the first reaction is an oxidized species (called OX1) that occupies more membrane area than non-oxidized DOPC does. The product of the second reaction is called OX2, which occupies less area than DOPC. Bateman and Gee 66 observed that the kinetics of photochemical oxidation of non-conjugated olefins have first-order dependence on the concentration of the species to be oxidized. We therefore assume first-order kinetics for both steps. The dependence of reaction rate on illumination intensity will be taken into account via modifications of the effective rate constant of both reactions, which are € DOPC k 1 " → " OX1, r 1 =k 1 C DOPC (2.3.3-1a) € OX1 k 2 " → " OX2, r 2 =k 2 C OX1 (2.3.3-1b) where k 1 and k 2 are rate constants for the first and second steps, respectively, and r 1 and r 2 are reaction rates. The material balances for the three species are € dC DOPC dt =−k 1 C DOPC (2.3.3-2a) € dC OX1 dt =k 1 C DOPC −k 2 C OX1 (2.3.3-2b) € dC OX2 dt =k 2 C OX1 (2.3.3-2c) Using the initial conditions € C DOPC t =0 ( ) =1 (2.3.3-3a) 37 € C OX1 t =0 ( ) =C OX2 t =0 ( ) =0 (2.3.3-3b) this system can be analytically solved in order to obtain expressions for the relative concentrations as a function of time (t), which are € C DOPC t ( ) =e −k 1 t (2.3.3-4a) € C OX1 t ( ) = k 1 k 1 −k 2 e −k 1 t −e −k 2 t ( ) (2.3.3-4b) € C OX2 t ( ) =1− k 2 k 2 −k 1 e −k 1 t + k 1 k 2 −k 1 e −k 2 t (2.3.3-4c) We also assumed that the GUV surface area (S) is a simple linear combination of the surface areas of the three species (DOPC, OX1, and OX2) € S t ( ) =A DOPC C DOPC t ( ) +A OX1 C OX1 t ( ) +A OX2 C OX2 t ( ) (2.3.3-5) where A OX1 and A OX2 are areas per lipid of OX1 and OX2, respectively, relative to that of DOPC, A DOPC (i.e. A DOPC = 1, A OX1 > 1, A OX2 < 1). The rate constants (k 1 and k 2 ) and area parameters (A OX1 and A OX2 ) were obtained through fitting the above model (Equation 7.3.3-5) to normalized area data averaged from approximately twenty GUVs for each experimental condition. Figure 2.11 shows model results for 90%DOPC-10%RhDPPE GUVs irradiated at 2.85 mW over 15 minutes. Note that the area expansion regime only spanned an interval of 80 seconds, and more time points were considered during this interval in which surface area changed rapidly. 38 Figure 2.11. Kinetic model (blue, solid line) fitted to data (black, dots) averaged from 20 GUVs of 90 mol% DOPC-10 mol% RhDPPE composition, irradiated at 2.85 mW for 15 minutes. Parameters obtained from this fit are A OX1 = 1.1165±0.0148, A OX2 = 0.4867±0.0021, k 1 = 0.0977±0.0037 s -1 , and k 2 = 0.0061±0.0001 s -1 . We studied the same GUV composition (90 mol% DOPC-10 mol% RhDPPE) excited by five different power levels measured to be 2.85, 1.40, 0.72, 0.38, and 0.19 mW. Each condition was represented by time-course data averaged from approximately 20 GUVs. The fitting of the kinetic model (Equation 2.3.3-5) to the normalized area data (Y i ) utilizes Nelder and Mead’s simplex method 81 executed on MATLAB to minimize the objective function € F = w i Y i −S t i ( ) ( ) i=1 n ∑ 2 (2.3.3-6) which sums for n data points the product of the square of the residuals and the weighting function € w i = 1 σ y,i 2 (2.3.3-7) with σ y,I being the uncertainty of Y i . These fitting parameters α = [k 1 , k 2 , b, c] have the uncertainty 82 0 200 400 600 800 Time / s 0.5 0.6 0.7 0.8 0.9 1 1.1 Normalized Surface Area 39 € σ a i = F n− p C kk −1 (2.3.3-8) for p number of parameters, and where C is a p-by-p matrix and its elements are defined as € C jk = w i ∂S ∂a j ∂S ∂a k i=1 n ∑ (2.3.3-9) The data points (Y i ) contain 20-second intervals, with the exception of the area expansion region (during the first 40 seconds for the data set with 2.2 mW irradiation shown in Figure 2.11), where 1-second interval data points were used to emphasize this dynamical region for the least squares minimization. Similar treatment was done for other irradiation strength conditions: more points were added to span the membrane-expansion period without extending into the transitional region in between membrane expansion and contraction phases. This was done to minimize the influence of experimental artifacts on the fitting parameters. The kinetic expression given by Bateman and Gee 66 shows that oxidation rate scales as the square root of light absorption. Since the rate of light absorption linearly varies with the incidental light intensity, we plotted the rate constants as a function of the square root of intensity and observed approximately linear behavior. The results of four fitting parameters are summarized below in Figure 2.12. 40 Figure 2.12. (a) Area parameters A OX1 (green, filled circles) and A OX2 (orange, opened circles) as functions of the irradiation intensity (I) and (b) rate constants k 1 (blue, opened squares) and k 2 (red, filled squares) as functions of I 1/2 , obtained through fitting the kinetic model to vesicle area time-course data averaged from ~20 90 mol% DOPC-10 mol% RhDPPE GUVs. Note that some error bars are smaller than the markers and are not visible. The area parameter A OX1 represents the relative area of the oxidized species resulting from the first rate-limiting step of DOPC oxidation, compared to the lateral area that a DOPC molecule occupies: 0.68 nm 2 . 58 To estimate this parameter, we averaged the best-fit value from experiments at various illumination intensities. Our estimate of the molecular area increase of oxidized DOPC at 16.9% is within the ranges reported by Weber (15-20%) 60 and predicted by molecular dynamics simulation. 44 Figure 2.12a also depicts diminishing values of A OX2 , the relative area of the final oxidized species OX2, with higher irradiation intensity. During this second area-contracting regime, the GUVs assumed rigid spherical geometry and the area measurement artifacts associated with the first oxidation phase were eliminated. Smith and coworkers 80 demonstrated that lipid tail scission from oxidation resulted in loss of lipid materials as fragments from the membrane into the aqueous environment. Additionally, the molecular dynamics simulation work of Cwiklik and Jungwirth 58 predicted micellization of the shortened oxidation products. These 0 0.5 1 1.5 2 Intensity 0 0.02 0.04 0.06 0.08 0.1 0.12 k / s 0 0.001 0.002 0.003 0.004 0.005 0.006 0.007 k k 1 k / s 2 1/2 1 2 (b) A A OX1 OX2 0 0.5 1 1.5 2 2.5 3 Irradiation Intensity (mW) 0.4 0.6 0.8 1 1.2 Area Parameter (a) 41 results support our finding, shown by the behavior of A OX2 , that increased irradiation intensity (and thus increasing singlet oxygen formation and lipid oxidation rates) causes a collective decrease in the membrane area resulting from the loss of lipid mass from the membrane. The rate constants k 1 and k 2 were expected to vary directly with the square root of irradiation intensity, following the model by Bateman and Gee. 66 At the highest irradiation intensity, k 1 appears to not conform to this model. Note that k 1 is related to the shape of the area-expansion peak. The rapid onset of this peak at high irradiation intensities presents an observational challenge that may have yielded an inaccurate result. Extrapolating the value of k 1 at 2.85 mW based on the linear fit of the other four points yielded k 1 = 0.0490 s -1 , which also resulted in a reasonable fit to the surface area time course data (Figure 2.13). With accurate area measurement in the area-contraction regime, the results of k 2 more clearly display its linear dependence on I 1/2 . Despite maintaining constant irradiation intensity throughout the span of each 15-minute experiment, the first oxidation step (to OX1) took place more rapidly than when oxidizing from OX1 to OX2. This could be visualized in Figure 2.5g and quantitatively confirmed by the difference between k 1 and k 2 . The fact that k 1 is one order of magnitude higher than k 2 is largely due to the autocatalytic nature of the first phase, where the allylic hydrogen abstraction of one lipid molecule resulted in a radical that can initiate oxidation of neighboring lipid molecules in the membrane. 42 Figure 2.13. Kinetic model fit using k 1 = 0.0997 s -1 (blue) and k 1 = 0.0490 s -1 to the data (black) averaged from 20 GUVs of 90 mol%-10 mol% RhDPPE composition, irradiated at 2.85 mW for 15 min. The other parameters used are b = 1.1165, c = 0.4867, and k 2 = 0.0061 s -1 . With increased accuracy in membrane area measurements, we were able to use the data collected with the DBOT apparatus for fitting to the dynamical model. However, the GUV surface area could only be calculated until the maximum surface area increase was reached. Without certainty on whether membrane pores formed any time afterwards, we could neither assume constant volume for surface area calculation nor maintain the optical contrast required for vesicle stretching. The model requires data spanning over both membrane expansion and contraction phases, so there were not enough data points collected with the DBOT apparatus to constrain a four-parameter fit. With the parameters shown in Figure 2.12 for 0.19, 0.39, 0.73, and 1.4 mW intensities, we assumed linear dependence of A OX2 on the irradiation intensity and of k 1 and k 2 on the square root of the intensity to extrapolate these three parameters for the irradiation intensity used in the DBOT experiments. Due to 43 differences in light sources and microscope objective specifications between the basic setup shown in Figure 2.1 and in the DBOT setup, this irradiation intensity (0.02 mW) was estimated by extrapolation from the linear fit as a function of the natural logarithm of time at the maximum area increase on the basic experimental setup, shown in Figure 2.14. Figure 2.14. Fitting the natural logarithm of irradiation intensity from the basic oxidation experiment as a linear function of time. This linear fit (solid line) of ln(I) = -0.0219t + 1.2793 was used to extrapolate at t = 237 s to 0.02 mW irradiation intensity used in the DBOT setup. Figure 2.14 shows how the irradiation intensity was extrapolated for the oxidation experiment on the DBOT apparatus. At 0.02 mW, the parameters A OX2 , k 1 , and k 2 were estimated to simplify the model to fit with the data for one parameter, A OX1 , which was estimated to be 1.1076 ± 0.0272. We have considered how strongly the result of A OX1 depended on the extrapolated intensity value: at one order of magnitude higher and lower, the fits to the data were visually inaccurate while the resulting range of A OX1 values (1.1113 ± 0.0270 and 1.1381 ± 0.0349, respectively) overlapped with the reported range of uncertainty. 44 The resulting one-parameter fit of the kinetic model to the DBOT data yielded A OX1 of 1.1076 ± 0.0272; the distribution of A OX1 is shown in Figure 2.15. We have acknowledged that the data from the basic experimental apparatus resulted in overestimated values of GUV surface area, so it is reasonable that the DBOT data would suggest a 10.7% increase in lipid area, which is less than the value obtained from the basic experiment data of 16.9%. However, this value is less than the lower limit of the ranges reported by Weber and predicted by Wong-Ekkabut. 44,60 Figure 2.15. The distribution of A OX1 obtained through fitting the kinetic model to the DBOT data of 19 90 mol% DOPC-10 mol% RhDPPE GUVs irradiated at an estimated intensity of 0.02 mW (561 nm); the Kolmogorov-Smirnov test confirmed normal distribution of the values averaged at 1.1076 ± 0.0272. 2.3.4. Line tension of oxidized membrane pores In addition to vesicle surface area changes, the excitation of the fluorescent dye labeling the phospholipid bilayer also resulted in experimentally observed pores in the membrane (Figure 2.5d-f), such as those seen by Sandre and coworkers. 83 Pores often form at tensions beyond a certain threshold; above this threshold, vesicle membrane stress is mitigated by the ejection of liquid through pores. Energetically, pore opening competes with the line tension that arises from the energetic cost to arrange lipid molecules at the pore edge, which is why lipid pores are often small and short-lived and are not typically visible in standard microscopy experiments. 83 Count 4 8 1 1.04 1.08 1.12 1.16 1.2 A OX1 0 12 45 Sandre and coworkers 83 were able to observe pores by prolonging their durations. They formed GUVs in a mixture of water and glycerol, increasing the viscosity to 32.1 ± 0.4 mPa•s and slowing down the leakage process to delay pore closure. The pores formed could be resolved by optical microscopy and remained opened for seconds. Another way pore duration can be modified is altering membrane line tension by adding molecules with non-zero spontaneous curvatures. Surfactants and phospholipids with relatively large head groups can more easily be organized at the pore edge than cylindrical lipids can. Treatment of DOPC GUVs with the surfactant triton X-100 resulted in pores that could last tens of seconds. 84 The opposite effect is observed for cholesterol, which has a cone shape. 85 Using glycerol to raise the viscosity of the aqueous phase ten-fold to 10.3 mPa•s, we increased the duration of pore opening from less than a second to 3-4 seconds, giving us 20-40 frames of epifluorescent micrographs for each pore occurrence. We observed pore formation for three different compositions of decreasing DOPC concentration. As DOPC concentration decreased from 90% to 68%, membrane line tension increased from ~6 pN to ~10 pN and pore occurrence became less frequent with shorter duration. This indicates that a DOPC oxidation product resulting from lipid tail scission during the membrane-contracting phase decreases membrane line tension. This result suggests that these lipid products have edge-stabilizing inverted cone geometry. The distributions, confirmed to be normal by the Kolmogorov-Smirnov test, of the line tension values for all three conditions are summarized in Figure 2.16. 46 Figure 2.16. Distribution of measured line tension values (pN) for (a) pure DOPC (6.10 ± 2.08 pN, n = 93), (b) 7:1 DOPC/DPhPC (8.89 ± 2.66 pN, n = 54), and (c) 3:1 DOPC/DPhPC (9.95 ± 2.56 pN, n = 42) GUVs, all with 10 mol% RhDPPE, in 1:1 v/v 200 mM sucrose solution/glycerol (μ = 10.3 mPa•s). Smith and coworkers 80 studied lipid tail scission with oxidation and demonstrated that some tail fragments were lost to the aqueous environment. This is depicted in our study by shrinking GUV size following initial membrane-expansion. The contribution of lipid oxidation towards pore formation is two-fold. First, the decreasing vesicle membrane area encompassing an incompressible fluid accumulates surface tension required to drive pore opening. Second, the shortened lipid oxidation products were shown by us to reduce line tension with a pore edge- stabilizing geometry. Surface tension promotes pore formation while line tension acts against it, and lipid membrane oxidation alters both properties in favor of pore opening. In cells, increased membrane permeability due to pores then leads to solute imbalances triggering cell death. 86 2.3.5. Controlling for oxidation with sodium azide and Trolox To demonstrate that oxidation was due to singlet oxygen, we performed the basic photooxidation experiment described in section 2.3 in the presence of sodium azide, a specific quencher for singlet oxygen, 87 and of Trolox, which scavenges free radicals. 88 With 1 mM Trolox present in the glucose solution outside the 90 mol% DOPC-10 mol% RhDPPE GUV, irradiation with green excitation light at 0.72 mW Count 10 20 30 0 2 4 6 8 10 12 14 16 18 20 (a) 5 10 15 0 2 4 6 8 10 12 14 16 18 20 Line Tension (pN) (b) 5 10 0 2 4 6 8 10 12 14 16 18 20 (c) 47 resulted in the same vesicle morphologies as in the condition without Trolox (Figure 2.17). Figure 2.17. Oxidation of a 90 mol% DOPC-10 mol% RhDPPE GUV in the presence of 1 mM Trolox outside the vesicles at (a) 0 min, (b) 1 min, and (c) 15 min of excitation light irradiation at 0.72 mW. Scale bars = 10 µm. The addition of 20 mM sodium azide yielded more interesting results. The presence of sodium azide induced vesicle adhesions, as shown in Figure 2.18a. During the course of irradiation for photosensitization of the rhodamine dye, the GUV membrane area did not increase but the vesicles eventually became smaller, as seen in the comparison between six GUVs in Figure 2.18. We speculated that azide ions were gradually depleted during the 15 minutes of irradiation, as seen by restored vesicle mobility and separation (Figure 2.18b), so milder lipid oxidation did occur. Nevertheless, as sodium azide specifically quenches singlet oxygen, this control experiment demonstrated that this ROS, formed through photosensitization, was in fact responsible for the oxidation of DOPC lipid molecules. 48 Figure 2.18. Vesicle adhesion of 90 mol% DOPC-10 mol% RhDPPE GUVs from the addition of 20 mM sodium azide outside the vesicles after (a) 1 min and (b) 15 min of excitation light irradiation at 0.72 mW. Vesicle movement resulted in decreased numbers of GUVs; the remaining ones are labeled (1-6). Scale bars = 10 µm. 2.3.6. Identifying oxidation products We analyzed the chemical transformation of DOPC upon photooxidation. A 10 mol% mixture of rhodamine B and DOPC was probed by 1 H NMR and successively oxidized as a thin film. The rhodamine B signal did not interfere with the DOPC signal. We observed the evolution of three non-native DOPC peaks that are attributed to hydroperoxy, alcohol, and aldehyde groups (Figure 2.19). 49 Figure 2.19. 1 H NMR spectra of oxidized (left) and non-oxidized (right) samples of 90%DOPC-10%Rhodamine B. The colored dots indicate the oxidation products: (blue) hydroperoxy, (green) alcohol, and (red) aldehyde. The bottom spectra are of stock samples and each successive spectrum was taken after a cycle of evaporating chloroform (both oxidized and control), irradiating to excite Rhodamine B for 1 h (oxidized batch only), and redissolving in chloroform for 1 H NMR spectroscopy (both oxidized and control). Singlet oxygen reacts with the olefin in the DOPC tail according to standard “ene” chemistry. In a concerted reaction, singlet oxygen reacts with an allylic hydrogen (with respect to the double bond) and a hydroperoxy group adds to the tail, shifting the double bond. 89-91 The hydroperoxy peak appears in the spectra after the first round of oxidation at around 4.4 ppm. Subsequent reduction of the hydroperoxide results in the formation of an alcohol. Finally the allylic alcohol undergoes a Hock rearrangement and the tail cleaves, resulting in an alcohol (4.25 ppm) and an aldehyde (9.75 ppm). In the end, the alcohol is converted to an aldehyde. 51,92,93 These 50 aldehydes are present after the third round of oxidation. The head group was chemically stable as indicated in the consistency of spectra for all rounds of photooxidation in the 0.7-3.45 ppm range (Figure 2.20). The control sample, which underwent the same drying and scanning protocol, did not exhibit any of the oxidation products. Figure 2.20. 1 H NMR spectra of oxidized (left) and non-oxidized (right) samples of 90%DOPC-10%Rhodamine B. The bottom spectra are of stock samples and each successive spectrum was taken after a cycle of evaporating chloroform (both oxidized and control), irradiating to excite Rhodamine B for 1 h(oxidized batch only), and redissolving in chloroform for 1 H NMR spectroscopy (both oxidized and control). 2.4. Conclusions We demonstrated that the complex and multistep chemical processes of lipid oxidation could be simplified to rate-limiting steps in the formation of quasistable oxidation products, in order to follow the depletion of the original unsaturated lipid species without taking any chemical measurements. This kinetic model can be used as a tool for estimating the lateral molecular area of the resulting oxidation products, as well as the rate constants of the steps to form them. Applications beyond the fundamental studies may extend to fine-tuning the designs of photodynamic therapy 51 treatments and controlled drug delivery vehicles through controlled lipid membrane deterioration with oxidation. 52 Chapter 3: Oxidation of Non-Bilayer Forming Lipids c 3.1. Introduction Lipids in cellular membranes can exist in a fluid lamellar (L α ) phase, forming a planar bilayer morphology as depicted in the fluid mosaic model. 17 Non-lamellar lipid structures can exist, and are often associated with dynamic cellular processes. In the inverted hexagonal (H II ) phase, lipid molecules arrange into cylinders with the head groups oriented inward in contact with the aqueous phase core. 94 Lipids that form the H II phase have been observed at higher concentrations in intermediate structures during membrane fusion in model membranes and in cells. 95-97 The inverted cubic (Q II ) phase consists of lipids arranged into minimal surfaces. 98 It is periodic in three dimensions and creates two discrete networks of water channels. 99 The Q II phase has been observed during plasma membrane folding through non- clathrin mediated endocytosis. 100 Cubic phases have been previously associated with smooth endoplasmic reticulum (ER) and certain states of inner mitochondrial (MT) membranes. 100,101 Under numerous nicknames, including “undulating membranes” 102 and “tubuloreticular structures”, 100,103 cubic phase structures are regarded as potential c The work presented in this chapter has been published in Langmuir under the title “Oxidation of membrane curvature-regulating phosphatidylethanolamine lipid results in formation of bilayer and cubic structures” and used with permission from the American Chemical Society. This work was authored by Shalene Sankhagowit, Ernest Y. Lee, Gerard C. L. Wong, and Noah Malmstadt. S.S. and N.M. designed the study. S.S. designed and performed the optical microscopy experiments. E.Y.L. designed and performed the SAXS experiments. S.S., E.Y.L, G.C.L.W., and N.M. wrote the paper. All authors reviewed the results and approved the final version of the manuscript. 53 ultrastructural markers for cellular stresses. 104 They have been observed in smooth ER membranes of severe acute respiratory syndrome (SARS)-infected cells 105 and tumor cells in various organs. 102 One of the most well characterized in vivo cases of cubic phase formation results from the morphological transition of the inner MT membrane upon food depletion in the giant amoeba Chaos carolinensis. 104 The progressive formation of the cubic phase was accompanied by increasing levels of the reactive oxygen species (ROS) superoxide and hydrogen peroxide, resulting from changes in oxidative metabolism of the organism upon starvation. 106 It is important to note that these studies reported cubic phase formation in smooth ER and inner MT membranes that contain highly folded membranes with high concentrations of lipids with non-zero intrinsic curvature. 101,107-109 Approximately half of the phospholipids comprising the highly folded and compartmentalized inner MT membrane in mammalian cells are curvature-regulating phosphatidylethanolamine (PE) and cardiolipin species. 109 These lipids are found in particularly high concentrations at the contact sites between inner and outer mitochondrial membranes. They likely play a role in protein import and phospholipid translocation between the two layers of the organelle. 110,111 It has also been shown that PE lipids facilitate formation of membrane fusion intermediate structures 95,96 and support cytokinesis during cell division. 112 The ability of PE lipids to support high membrane curvatures for routine cell functions and division arises from their molecular structures. PE lipids have a smaller head group relative to the cross-sectional area of the hydrocarbon tails than phosphatidylcholine (PC) lipids that comprise the majority of phospholipids. Lipid 54 molecules that support formation of the lamellar phases in bilayer membrane, such as PC lipids, have comparable lateral areas in the head and tail regions and have an overall cylindrical molecular geometry. For phospholipids in general, the ratio of head-to-tail area is temperature dependent, with the area occupied by the tail groups increasing with temperature. 113 Above the phase transition temperature (T h ), a PE lipid can have an anisotropic wedge shape: it looks fan-shaped (hydrophobic region wider that the head group) in the plane that contains the centers of mass of the two alkyl chains; however, perpendicular to this plane, the shape is much less anisotropic. 114 This wedge shape supports the formation of the H II phase. Our study focuses on the lipid 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE) that transforms from the bilayer-forming fluid lamellar phase (L α ) to the H II phase as temperature increases across T h = 13º C. 115 Oxidation alters lipid molecular structures and can disrupt PE lipid functionality. While many products can form upon oxidation of the lipid bilayer, the typical result involves scission of an unsaturated hydrocarbon tail and/or the addition of a polar group to the same chain. 116 The common consequence is the loss of a portion of the hydrophobic moiety, either to the hydrophilic region or by removal from the bilayer into the aqueous phase. 1 This results in a subpopulation with a reduction of the negative intrinsic curvature. In this study, we used optical microscopy to observe the hydration of oxidized DOPE lipid films at room temperature. With the addition of water, the lipid film swelled to form micron-scale lamellar phase structures that included elongated tubules and spherical vesicles. In contrast, the non-oxidized, H II phase DOPE shows 55 no apparent changes upon hydration to the lipid film at this length scale. Using the same viewing technique, we saw formation of sub-micron-scale lattice structures that are stable for up to 3 hours. Characterization of 2-D projections in optical micrographs with a 2-D discrete fast Fourier transform (FFT) revealed that such lattices resemble body-centered cubic structures, with the lattice parameter on the order of hundreds of nanometers. Small-angle X-ray Scattering (SAXS) in general cannot access micron scale ordering in weakly scattering structures, such as those depicted in the micrographs here. However, SAXS can assay unambiguously the existence of cubic phases. Using SAXS, we find that oxidized DOPE can form a 3-D Im3m cubic phase with a lattice parameter of 14.04 nm, which indicates that the DOPE oxidation process has enhanced the system’s ability to form negative Gaussian curvature, the type of curvature topologically required for membrane permeation processes. Lipid membranes in Q II phases can be formed in vitro with monoacylglycerides to provide 3-D periodic structures for protein crystallization. 117,118 Cubic phases have also been shown to emerge after temperature cycling DOPE (as well as its monomethylated analog, DOPE-Me) across its phase transition temperature hundreds of times. 119,120 The lattice parameters of these cubic phase structures range between tens and hundreds of angstroms and are tunable with either lipid or aqueous phase composition. 118,121,122 The largest reported stable cubic phase lattice parameter so far is 470 Å, and transient cubic structures have been formed at 300-400 Å. 123 Although chain packing does not permit the formation of cubic phases based on minimal surfaces at these large length-scales, it is interesting 56 that DOPE oxidation apparently drives the generation of lipid structures with cubic symmetry, and do so at length-scales comparable to those of cubic structures observed in vivo. 102,104,124 It has been previously hypothesized that membrane lipids can be structured into the cubic phase by oxidative damage. 104 We quantified the extent of oxidation with nuclear magnetic resonance (NMR) spectroscopy in order to estimate the effective molecular shape of the collective oxidation product species that could support bilayer formation. We then observed hydrated films of non-oxidized DOPE with various concentrations of a lyso-PE lipid in order to mimic the process of tail group area reduction that occurs with oxidation. None of the combinations between H II phase and micelle formation yielded lattice structures that are observable optical microscopy. Our results suggest that the cubic structure formation in oxidized DOPE results from molecular alterations that affect lipid packing beyond changes to heat- to-tail group area ratios. 3.2. Experimental 3.2.1. Materials The lipids 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE), 1,2-dioleoyl-sn- glycero-3-phosphocholine (DOPC), and 1-oleoyl-2-hydroxy-sn-glycero-3- phosphoethanolamine (lyso-PE) were purchased from Avanti Polar Lipids. Chloroform was from Macron Fine Chemicals, and deuterated chloroform was from Cambridge Isotope Laboratories, Inc. 57 3.2.2. Oxidation of DOPE DOPE was oxidized while dried and exposed to air. Heat was applied to accelerate the oxidation process. For each DOPE sample, 1 mL of the lipid solution at 10 mg/mL in chloroform was dried on a glass petri dish (60 x 15 mm, VWR) with an argon stream and stored under vacuum for 90 minutes. For each 4-hour interval of oxidation, the lipid film-lined petri dish was then placed on a hot plate (C-MAG HS 7, IKA) set to 50º C. Three samples were oxidized in parallel. The petri dishes were covered to avoid falling contaminants but air exposure was allowed. Outside the 90- minute dry time under vacuum and 4-hour oxidation time on the hot plate, the lipid samples were sealed under an argon environment and stored dissolved in chloroform at -20º C to avoid unintentional oxidation. 3.2.3. Hydration of lipid films To observe the morphology of a hydrated oxidized DOPE sample, lipid films were prepared by drying 5 µL x 10 mg/mL of the lipid in chloroform as approximately 0.5 µL spots on a round glass coverslip (#1, 25 mm, Chemglass) with a 25 µL glass syringe (Hamilton). The lipids were allowed to dry for at least 90 minutes under vacuum before the coverslip was placed in a Sykes-Moore viewing chamber (Bellco Glass). Note that non-oxidized DOPE, DOPC, and monoolein samples were also prepared using this protocol, dried with 0.05 mg of lipids on each coverslip. For the hydration of lyso-PE/DOPE mixtures, 5 µL x 10 mg/mL DOPE was combined with different volumes of 5 or 10 mg/mL lyso-PE solutions to yield 10, 20, 30, 40, 50, 60, 70, 80, 90, and 95 mol% lyso-PE. These solutions were diluted with different amounts of chloroform such that the final concentration of DOPE in 58 each was approximately 0.7 mg/mL. For each mixture, 5 µL was then used to dry approximately 0.5 µL spots on a glass coverslip as described above. For observation, the edge of a spot of dried lipid film close to the center of the coverslip was located using a Zeiss Axio Observer inverted microscope operating under the differential interference contrast (DIC) mode, with a Plan-Apochromat 20x/0.8 NA objective or a Plan-Apochromat 63x/1.40 NA oil objective with 1.6x optovar magnification. The lipid film was hydrated with 500 µL Milli-Q water (Millipore). Note that this high volume was used to maintain hydration at the center of the coverslip while the added water wetted the wall of the viewing chamber. 3.2.4. SAXS experiments Small-angle X-ray scattering experiments were conducted to investigate the structure of oxidized DOPE in solution. Samples were equilibrated for two days before measurement. 24-hour oxidized DOPE and regular DOPE were solubilized in chloroform at 12.5 mg/mL and 20 mg/mL, respectively. Individual samples were aliquoted into smaller volumes in glass vials. The chloroform was evaporated from each vial under dry nitrogen gas and the sample was further desiccated overnight under vacuum. After 24 hours, Milli-Q water was added for resuspension to 20 mg/mL. Samples were prepared for SAXS by diluting hydrated regular DOPE or oxidized DOPE to 10 mg/mL in Milli-Q water to a total volume of 40 µL. Quartz glass capillaries (Hilgenberg) were loaded with the 40 uL samples and hermetically sealed. Measurements of the samples were made at the Stanford Synchrotron Radiation Lightsource, beamline 4-2. A monochromatic 9 keV X-ray with a 1.7 m 59 path length was used, and samples were exposed for 10 seconds. Scattered radiation was collected using a Rayonix MX225-HE detector (pixel size, 73.2 µm). 2-D diffraction images were processed and integrated with the Nika 1.68 package for Igor Pro 6.37 and FIT2D. For all samples, several measurements were taken to ensure consistency of data. 3.2.5. Chemical analysis of oxidized DOPE To approximate the extent of oxidation of DOPE tails, the ratio between the amounts of vinyl and glycerol protons obtained from nuclear magnetic resonance (NMR) spectroscopy was determined. The lipid samples (approximately 10 mg) in chloroform were dried onto the bottom of a glass test tube under an argon stream and further dried under vacuum for at least 90 minutes before being dissolved in 800 µL deuterated chloroform. If the lipid were already dried on a glass petri dish (after an oxidation step), then it was directly dissolved into deuterated chloroform. Samples were scanned on a Varian Mercury 400 2-channel NMR spectrometer at 25º C. Manual phase and baseline correction, as well as integration, were performed using the MNova NMR processing software (Mestrelab Research). 3.3. Results and discussions 3.3.1. Hydration of oxidized DOPE lipid films Lipid morphologies were observed using optical microscopy in differential interference contrast (DIC) mode to enhance the edges of structures. The lipid samples were deposited directly onto the glass coverslips used in microscopy for time-course observations of lipid hydration from the dried state. Oxidized DOPE was 60 prepared in 10 mg batches, and oxidation was induced during 4-hour intervals for a total of 24 hours. To optimize the oxidation process, the lipid was dried onto a glass petri dish and heated to 50º C while exposed to ambient air during each 4-hour interval. Additionally, each batch was dissolved in chloroform and, again, dried with an argon stream and placed under vacuum for 90 min in between each oxidation interval. This was done to improve product uniformity as oxidation more readily occurred on the surface of the dried lipid film. To avoid unintentional oxidation of the samples, the lipids were stored dissolved in chloroform under an argon environment at -20º C outside of the 4-hour oxidation and 90-min vacuum-drying periods. For each hydration experiment, a lipid sample in chloroform at 10 mg/mL was applied as approximately 0.5 µL spots with a glass syringe onto the coverslip and placed under vacuum for at least 90 minutes for solvent removal. For uniformity, the results presented here are of 90-minute dried lipid films, although the same results were achieved if the lipid were dried overnight. The coverslip was then placed in a Sykes-Moore viewing chamber for microscopy. A series of micrographs from a time-course observation of oxidized DOPE film hydration is shown in Figure 3.1. Image capturing was initiated seconds prior to the addition of Milli-Q water to record the dried lipid film (Figure 3.1A). The lipid film edges are ideal for observation due to higher film thickness made by the coffee ring effect when drying. Note that the area of the film close to the left edge of the micrograph was thicker than in the rest of the image. Within 20 seconds of hydration, oxidized DOPE formed vesicles, as well as loose and densely packed tubules (Figure 3.1B, right and 61 left sides, respectively). The latter quickly organized into lattice structures with clusters of periodic domains that spanned tens of microns (Figure 3.1C, left half). Figure 3.1C shows coexistence of the L α phase (represented by giant vesicles) and this lattice structure. The lattice was stable for minutes before its transition back into densely packed tubules (Figure 3.1D, E) that dispersed in both lateral and vertical directions (Figure 3.1F). Note that structures appearing at higher contrast in Figure 3.1F were further away from the coverslip and the focal plane was not changed. The duration at which the lattice structure appears, from minutes to an hour, varied based on the concentration of the lipid solution in chloroform during film preparation. Higher concentrations yielded thicker lipid films and resulted in longer-lived latticed structures. 62 Figure 3.1. Hydrated 24-hour oxidized DOPE lipid. (A) Dried lipid film. Hydrated lipid film, (B) 20 s and (C) 4 min after addition of water, showing a lattice structure (left side) and the lamellar phase as vesicles and tubules (right side), followed by transition from the lattice structure into tubules (D) 7 min and (E) 8 min after hydration. (F) At 10 min after hydration, tubules lifted upwards and therefore appeared at higher contrast. Scale bars = 10 µm. Figure 3.2 compares another oxidized DOPE film in the dried state and at 5 minutes after hydration, observed under lower magnification. At this length scale, the lattice structure appears as nebulous clusters with indistinct boundaries. Additionally, high contrast globular structures were present from the beginning of the hydration period (Figure 3.1B-F and Figure 3.2B) and increased in numbers upon the transition of the lattice structure to tubules and vesicles (Figure 3.1F). A B D E C F 63 Figure 3.2. Hydrated oxidized DOPE lipid (low-magnification). (A) Dried lipid film and (B) hydrated lipid film 5 min after addition of water. Scale bars = 50 µm. For comparison with L α and H II phases, the hydration experiment was repeated using DOPC and non-oxidized DOPE, respectively (Figure 3.3 at higher magnification and Figure 3.4 at lower magnification). DOPC is a phospholipid commonly used in the laboratory to form L α phase structures. 125,126 After 5 minutes of hydration, the DOPC lipid molecules at the surface of the film had swelled into vesicles and tubules (Figure 3.3B). A B 64 Figure 3.3. Comparison of lamellar and non-lamellar phases in water. DOPC (L α phase): (A) dried and (B) 5 min after hydration. DOPE (H II phase at room temperature): (C) dried and (D) 5 min after hydration. Scale bars = 10 µm. Hydrated DOPE is expected to form the H II phase in the aqueous environment, 115 which is not optically resolvable. No formation of lamellar phase structures was observed 5 minutes after hydration (Figure 3.3D). An exception is the presence of several tubules formed at the edge of the DOPE film in Figure 3.4D (arrow and inset). The chemical analysis of lipid samples, presented later in this work, shows low levels of oxidation prior to the first round of exposure to an air stream. Immediately upon hydration, rearrangements within the DOPE film occurred that resulted in an apparently more compact film with receded edges. Under the same observation resolution used to view images in Figure 3.1, the lattice structure observed in hydrated oxidized DOPE was not seen in either of the DOPC and non- oxidized DOPE samples. A B C D 65 Figure 3.4. Comparison of lamellar and non-lamellar phases in water (low magnification). DOPC (lamellar phase): (A) dried and (B) 5 min after hydration. DOPE (hexagonal phase at room temperature): (C) dried and (D) 5 min after hydration. Note that a small fraction of DOPE was oxidized due to air exposure and formed tubules along the lipid film edge (arrow and inset). Scale bars = 50 µm (10 µm for inset). 3.3.2. Analysis of the cubic structure To characterize the lattice structure formed with hydrated oxidized DOPE, we applied a 2-D discrete fast Fourier transform (FFT) to visualize the 2-D projections of micrograph images in the frequency domain. A section cropped from a larger micrograph was used for FFT analysis in order to extract information only pertaining to the lattice structure. Due to the large number of structured domains that can be captured in the images, we assume that we have data at different orientations of the same structure. The micrograph in Figure 3.5A shows the lattice structure under two different orientations (in square boxes). A B C D 66 Figure 3.5. (A) Hydrated 24-hour oxidized DOPE showing two lattice orientations (boxed). Scale bar = 10 µm. Two-dimensional fast Fourier transform of (B) hexagonal and (C) square patterns; the indicated reflections in the latter correspond to reciprocal lattice vectors a* and b*. The angle between the reflections is 90º, and the lattice parameters are a = 628 ± 202 nm and b = 628 ± 217 nm. The FFT of an image cropped from each of these areas produced frequency domain images shown in Figure 3.5B-C. The reciprocal lattice vectors a* and b* in Figure 3.5C correspond to lattice parameters a = 625 ± 202 nm (standard deviation of the Gaussian function fitted to the reflection signal) and b = 628 ± 217 nm, and the angle between the means of the reflection signal is 90º. The reflection in between vectors a* and b* corresponds to a = 444 ± 72 nm period length, equivalent to the distance between the center and a corner of a unit cubic cell with 628 nm side widths. Thus, Figure 3.5C indicates that hydrated oxidized DOPE forms a transitional body- B C A a* b* 67 centered cubic system with lattice parameters close to half of a micron. The hexagonal reflections pattern in Figure 3.5B then corresponds to the same body- centered cubic system of a = 830 ± 150 nm lattice parameter, oriented close to the [111] direction. Similarly, the structure in Figure 3.1C has lattice parameters a = 304 ± 35 nm and b = 309 ± 68 nm (Figure 3.6). The reflections corresponding to the periodicity of the spaces in between cubic body centers and corners could not be clearly resolved in this case. Figure 3.6. (A) Section from Figure 3.1C showing the magnified lattice structure of oxidized DOPE. Scale bar = 2 µm. (B) Two-dimensional fast Fourier transform of (A); the indicated reflections correspond to reciprocal lattice vectors a* and b*. The angle between the reflections is approximately 92º, and the lattice parameters are a = 304 ± 35 nm and b = 309 ± 68 nm. We attempted to characterize the lattice structure in the z-dimension and collected image stacks of the same x-y location at multiple focal planes set to 0.01 µm intervals, spanning at least 4 µm. By reconstructing the image as x-z and y-z cross-sections of the image stack, the lattice intervals (a = 574 ± 135 nm and b = 574 ± 178 nm) seen in the x-y plane continued in the z-direction (Figure 3.7A). However, the lattice spacing in the z-direction could not be resolved. Taken together, these data a* b* A B 68 suggest that large structures with cubic symmetry can be made in the oxidation process. Figure 3.7. (A) Cross sections of a three-dimensional image stack of the lattice structure of hydrated oxidized DOPE, spanning 10.1 µm x10.1 µm x 4.42 µm (x-y-z). Scale bar = 2 µm. (B) Two-dimensional fast Fourier transforms at an x-y cross-section; the indicated reflections correspond to reciprocal lattice vectors a* and b*. The angle between reflections is 90º, and the lattice parameters are a = 574 ± 135 nm and b = 574 ± 178 nm. To determine whether oxidized DOPE can form a stable cubic phase structure, we utilized SAXS. 127,128 We find that oxidized DOPE forms an Im3m body-centered cubic lattice with a lattice parameter of a = 14.04 nm (Figure 3.8). Three peaks are observed at q = 0.063, 0.090, and 0.110 Å -1 , which index to a ratio of √2:√4:√6. Higher order reflections are also observed at √10:√12:√14, corresponding to q = 0.142, 0.154, 0.16. The magnitude of negative Gaussian curvature of this phase is <k> = 2πχ/A 0 d 2 = -5.4 x 10 -4 Å -2 where χ = -4 and A 0 = 2.345. For comparison, the sample prepared with non-oxidized DOPE showed inverted hexagonal phase characteristics with a lattice constant of a = 7.46 nm, which is consistent with values found in the literature. 129 y x z a* b* A B 69 Figure 3.8. SAXS results for oxidized DOPE showing the Im3m phase with a = 14.04 nm (yellow) and non-oxidized DOPE showing the inverted hexagonal phase (blue). Three inverted cubic phase supported by lipid species have been observed: primitive (Im3m), double diamond (Pn3m), and gyroid (Ia3d). A commonly fabricated lipidic cubic phase structure is composed of 1-oleoyl-rac-glycerol 70 (monoolein, MO) that forms the Pn3m structure with 40% (w/w) hydration at room temperature and supports the gyroid type with lower hydration conditions. 118 The Pn3m and Ia3d cubic phases are preferred by lipid species that support highly negative curvatures, similar to those forming the H II phase. 130 Monoolein-based lipid cubic phase structures that are formed in vitro usually have unit cell sizes of hundreds of angstroms, an order of magnitude lower than the cubic structures observed in nature. 123 While the double diamond and gyroid cubic phases contain tetrahedral (109.5º) and three-way (120º) water channel junctions, respectively, the Im3m phase contains 90º 6-way junctions and requires less negative curvature. 131 For this reason, Im3m structures are capable of more drastic cell size expansion compared to the other two cubic phases. 130 Anionic lipids and poly(ethylene glycol)- conjugated PE, both allowing for higher levels of head group hydration, induce formation of the Im3m phase and drastically increase the lattice parameter of monoolein-based lipid mixtures by approximately 70 Å, from 106 Å for pure monoolein. 118 These additive species form the L α phase in pure form and thus reduce the magnitude of the negative curvature when mixed with monoolein. 123 Similarly, DOPE acyl tails obtain polar functional groups through oxidation that causes the formerly hydrophobic moiety to migrate closer to the lipid-water interface and results in lipid head group area expansion. 1 Certain ternary mixtures of surfactants, water, and oil (similar to oxidation-scission lipid tail fragments) have been shown to support cubic lipid phases. 132,133 We speculate that the emergence of mixed subpopulations with different intrinsic curvatures, with the presence of negative curvature species (e.g. DOPE and monoolein) and L α phase-forming species 71 together, is significant in forming the Im3m phase and lattice structures with near- micron spacing. 3.3.3. Characterization of lipid geometry The oxidation of DOPE results in a mixture of products with increased head group areas at the expense of hydrophobic moieties. While DOPE supports the H II phase in water, the hydration of oxidized samples resulted in the formation of Q II (Im3m, confirmed by SAXS data) and L α phase structures that are supported by more cylindrical molecules. The deviation of a lipid molecular structure from resembling a cylindrical volume can be described and used to predict the morphological phase it can support. Israelachvili and coworkers 134-136 studied lipid packing of various shapes of molecules and quantified lipid structure with the lipid packing parameter, γ = V/Al, where V is the volume occupied by the lipid tails, A is the lateral area of the head group, and l is the length of the lipid tail region. Cylindrical lipids that support lamellar bilayer formation have γ close to unity, while lipids with small head groups (γ > 1) such as PE lipids form the H II phase. DOPE has the packing parameter γ DOPE = 1.38, while its PC counterpart more closely resembles a cylindrical volume (γ DOPC = 1.09). 137 Here, we aim to estimate the effective packing parameter of the collective oxidized DOPE species from the observed L α phase morphologies. Kumar 138 has demonstrated that packing parameter is additive in lipids with long chain lengths (10 or more carbon atoms per acyl chain): the effective packing parameter for a mixture of lipid species is the linear combination of the packing parameter of each individual species, weighted by mole fraction. To demonstrate that the calculated effective 72 packing parameter can predict hydrated lipid morphologies, we performed the hydration experiment using samples with varying ratios of DOPE to lyso-PE (γ lyso-PE = 0.5γ DOPE ). An 18-carbon, monounsaturated tail lyso-PE species was selected to mimic a truncated-tail DOPE molecule. The increasing molar fractions of lyso-PE were studied to simulate the increase in the extent of oxidation of the DOPE population. The lyso-PE/DOPE films were hydrated with Milli-Q water with 5 x 10 -7 M lyso-PE (estimated as the critical micelle concentration based on information from Marsh 139 ) in solution to minimize dissolution of the surfactant-like lipid from the film. At 100 mol% DOPE (γ = 1.38), the lipid film shows no significant changes when hydrated. As the fraction of lyso-PE increases, the calculated effective packing parameter decreases. From 40 mol% lyso-PE (γ = 1.10) upwards, tubules and vesicles were present within 5 minutes into hydration (Figure 3.9). The calculated packing parameter for this mixture is comparable to that of DOPC that forms the lamellar phase. Thus, we are using this packing parameter value for the transition point where the L α phase structures appear. 73 Figure 3.9. Lipid films of DOPE/lyso-PE mixtures showing lamellar phase structures (low magnification). (A) Dried and (B) 5 min after hydration of 60 mol% DOPE/40 mol% lyso- PE and (C) dried and (D) 5 min after hydration of 5 mol% DOPE/95 mol% lyso-PE. Scale bars = 50 µm. Note that the DOPE/lyso-PE mixtures were diluted with chloroform so that the final concentration before drying on the coverslip was uniformly 0.7 mg DOPE/mL. This is much lower than the 10 mg/mL concentration of the oxidized DOPE samples. We repeated the hydration experiment with oxidized DOPE using 0.7 mg/mL (Figure 3.10). At this lower lipid amount, the cubic lattice-to-lamellar phase transition occurred more quickly due to increased hydration rates. A B C D 74 Figure 3.10. Hydrated lipid films of (A) 60 mol% DOPE/40 mol% lyso-PE, (B) 5 mol% DOPE/95 mol% lyso-PE, and (C) oxidized DOPE dried at the same lipid concentration (approximately 0.7 mg/mL), 5 min after addition of water. Scale bars = 10 µm. As previously described, DOPE was oxidized in 4-hour intervals. After each oxidation period, we also analyzed the samples with proton nuclear magnetic resonance ( 1 H NMR) spectroscopy and performed the hydration experiment using 0.05 mg of the original 10 mg samples. The progress of oxidation was measured with the remaining amounts of vinyl protons, estimated by integrating the vinyl proton peak normalized to the area under the glycerol proton peak (highlighted blue and red, respectively, in Figure 3.11A). The normalized amount of vinyl protons for an ideal non-oxidized DOPE sample is 4. During the oxidation of a monounsaturated lipid such as DOPE, an allylic hydrogen is first abstracted from one of the lipid tails by a reactive oxygen species. The lipid molecule is left as a carbon-centered radical that reacts with molecular oxygen and another lipid molecule and results with a hydroperoxy group. While the peak for hydroperoxy proton is difficult to isolate in the NMR spectrum, the functional group being adjacent to the double bond causes a downshift of the vinyl peak (from 5.34 ppm to approximately 5.7 ppm, Figure 3.11B). The scission of the lipid tail with further oxidation eliminates the vinyl peak A B C 75 completely. Thus, integrating the vinyl proton peak of the 1 H NMR spectrum is a reasonable method for determining the fraction of DOPE lipids that is not oxidized. Figure 3.11. Estimating the extent of oxidation with 1 H NMR. (A) Molecular structure of DOPE showing vinyl protons (blue) and the glycerol proton (red). (B) 1 H NMR spectra of oxidized samples of DOPE showing the glycerol (5.22 ppm) and vinyl (5.34 ppm) proton peaks, as well as the emergence of the shifted vinyl peak (~5.7 ppm) due to the substitution of an allylic proton with a hydroperoxy group. The bottom spectra (red) are of the stock, non-oxidized sample and each successive spectrum was taken after a cycle of evaporating chloroform, heating at 50º C for 4 h, and redissolving in deuterated chloroform for 1 H NMR spectroscopy. (C) Progression of DOPE oxidation with air and heat exposure duration, measured as the integration of the vinyl proton peak with respect to the glycerol proton peak from the 1 H NMR spectrum obtained at each time point. Each data point is averaged from three samples oxidized in parallel, with the error bar representing the standard deviation. The progress of oxidation, as followed via the loss of vinyl protons, is shown in Figure 3.11C. Accompanying hydration experiments of samples after each set of NMR scans showed that oxidized DOPE films resembled that of non-oxidized DOPE, after up to 20 hours of air/heat treatment. However, the lamellar phase and cubic structures emerged after a total of 24 hours of oxidation (e.g. Figure 3.1), where (59.92 ± 1.66)% (standard deviation from 3 samples) of the vinyl protons were present. We noted that the stock DOPE sample (with 0 hours of intentional oxidation) started with (96.83 ± 1.50)% vinyl protons, and this was factored in for 0 5 10 15 20 Oxidation Duration / h 0.5 0.6 0.7 0.8 0.9 1 Fraction Vinyl Protons Present H P O - O O NH 3 + O O O O O H H H H B A C Cubic/Lamellar Inverted Hexagonal 5.4 5.3 5.2 5.8 5.6 Oxidation ppm 5.7 / ppm * not to scale between sets of spectra 76 the calculation of the effective packing parameter. This readjusted the effective packing parameter of the 60 mol% DOPE/40 mol% lyso-PE sample to 1.090 ± 0.014. In order for the 24-hour oxidized DOPE sample to have the same effective packing parameter, the average packing parameter value for the oxidized lipid portion was calculated to be 0.657 ± 0.069. This suggests that the inverted wedge or cone characteristic of an “average” oxidized DOPE molecule therefore has greater deviation from cylindrical than the original wedge shape, and in the opposite direction, of non-oxidized DOPE (i.e. γ oxDOPE < 1/ γ DOPE ). 3.4. Conclusions Oxidative stress is a common mechanism of cellular damage that leads to pathology in a whole host of human diseases, including cancer, atherosclerosis, and neurodegeneration. 6,140 Disruption of normal metabolic processes in the mitochondria and endoplasmic reticulum lead to formation of byproducts that either contain or lead to downstream production of reactive oxygen species. 6 Cellular membranes are especially susceptible to oxidative damage due to predominance of fatty acid chains. As a result, it is essential to understand the effects of lipid oxidation on the topology of cellular membranes. The molecular structure of PE lipids is central to its role in dynamic cellular processes where high curvature is involved. It serves as an ideal prototypical lipid for studies of oxidation-induced changes in molecular shape and intrinsic curvature. 95,96,112 In this study, we demonstrate that molecular alterations resulting from oxidation of PE lipids lead to drastic changes in the phase behavior of lipid membranes. The oxidation of a lipid molecule changes the ratio between its head and tail group areas. For DOPE, we 77 have estimated that the resulting oxidation products incur an intrinsic curvature sign change and, on average, contain a higher magnitude of curvature than the non- oxidized species. SAXS results show that the presence of oxidized DOPE induced structures rich in negative Gaussian curvature, which is a necessary ingredient in many membrane permeation events. 141 PE lipids are unique when compared to the more predominant PC phospholipids. For any lipid acyl tail, oxidation can result in the addition of a hydroperoxy group and the formation of an aldehyde group with tail scission. The amine group of the PE lipid head group can participate in a condensation reaction with this aldehyde group to form a fluorescent Schiff base product. 142-144 We speculate that it is this last, high-molecular weight product that provides the rigidity required to minimize thermal fluctuations for stabilizing such large lattice spacing. 78 Chapter 4: The Effects of Oxidation on Lipid Membrane Bending Rigidity d 4.1. Introduction Oxidation alters the geometry, and thus intrinsic curvature, of lipid molecules. We have observed cylindrical DOPC oxidize into inverted cone-shaped, pore edge- stabilizing products and DOPE being oxidized into a mixture of species that facilitates the formation of bilayer-forming lamellar phases and even transitional periodic cubic structures. This change can be more rigorously quantified by measuring the bending modulus the lipid membrane containing controlled amounts oxidized lipid species. The membrane bending rigidity is characterized with the bending modulus that describes the energy required to change the curvature of the bilayer from its intrinsic curvature governed by the geometry of the lipid molecules comprising the membrane. 145 Therefore, it is dependent upon the composition of the membrane 146 and changes with lipid oxidation. Membrane bending rigidity has been experimentally characterized through methods that analyze membrane thermal fluctuations or the stress-strain behavior through deformation of the membrane. The former is achieved through observation of the GUV membrane using optical microscopy 147-150 and of a lipid bilayer stack through scattering techniques. 151,152 d The data collection phase of the work presented in this chapter is still ongoing; the full work is expected to be completed in the next few months. This work will be authored by Shalene Sankhagowit, Shuyang Wu, and Noah Malmstadt. S.S. and N.M. designed the study. S.S. built the micropipette aspiration apparatus and wrote the analysis program. S.S. and S.W. performed the micropipette aspiration experiments. S.S., S.W., and N.M. will write the paper. 79 Deformation of GUV membrane can be incurred by using hydrodynamic, 153,154 electric, 155 magnetic, 156 and optical forces. 71,72,157,158 Existing work investigating the change in bending modulus with lipid peroxidation was based on red blood cell membrane thermal fluctuations and concluded that the membrane became more resistive to curvature changes with oxidation. 159 Bouvrais and coworkers also observed membrane undulations to compute changes in the bending modulus with the addition of 2 mol% of various fluorescent probes to POPC GUVs. 59 Despite dye sensitization being able to induce lipid oxidation, the main focus of their study was not to directly investigate the effects of lipid oxidation on the membrane mechanical properties. For the characterization of membrane bending rigidity, micropipette aspiration is a well-established technique and has long been used to measure the bending modulus of both cells and GUV membranes. During the experiment, a fraction of an enclosed bilayer membrane is drawn into the glass micropipette opening with a mild negative hydrostatic pressure (up to 10 2 Pa). 160 The resulting areal strain can be compared with the calculated membrane tension (from known vesicle and micropipette geometries and suction pressure) to obtain the value of the bending modulus. Fluorophore sensitization-induced lipid oxidation is a complex system in which to study the effects of lipid oxidation on the membrane bending rigidity. As shown in Chapter 2, oxidation leads to area expansion of individual lipid molecules, and this can be difficult to decouple from increased area due to aspiration through the micropipette. Furthermore, we have observed that applied pressure induces 80 budding and tubule formation from the aspirated membrane during oxidation. Thus, the study on the effects of oxidation on the membrane bending rigidity will be of GUVs with predetermined compositions containing the polyunsaturated lipid 1- palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (PLinPC, 16:0-18:1 PC) and its known oxidation product, 1-palmitoyl-2-(9’-oxo-nonanoyl)-sn-glycero-3- phosphocholine (POxnoPC). 161 4.2. Experimental 4.2.1. Materials The lipids 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1-palmitoyl-2-oleoyl- sn-glycero-3-phosphocholine (POPC), 1-palmitoyl-2-oleoyl-sn-glycero-3- phosphocholine (PLinPC), 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC), 1-palmitoyl-2-(9’-oxo-nonanoyl)-sn-glycero-3-phoshpcholine (POxnoPC), and cholesterol (plant derived) Chol were purchased from Avanti Polar Lipids. Sucrose, glucose, and alpha-tocopherol (α-Toc) were purchased from Sigma-Aldrich, and chloroform was from Macron Fine Chemicals. 4.2.2. GUV preparations For the control experiments to compare bending modulus values obtained from our micropipette aspiration apparatus to those reported in literature, the GUVs were composed of a single lipid species of DOPC, POPC, or DMPC. To study the effects of varying the amount of oxidized lipid in the GUV membrane composition, the mixture will be based on the PLinPC/DMPC/Chol mixture 161 or of pure PLinPC. Note that since PLinPC contains an easily oxidized polyunsaturated tail group, a 81 small amount of α-Toc would be added to the stock PLinPC solution to minimize unintentional oxidation. The known oxidation product of PLinPC, POxnoPC, would be substituted into fractions of PLinPC to achieve final contributions as 2.5, 5, 7.5, and 10 mol%. The fabrication of GUVs was done using the electroformation method 67 previously described in Section 2.2.2, with ITO-coated slides to conduct 10 Hz AC field at 1.3 V. In preparation for the electrofromation step, the lipid mixture was dried overnight under vacuum and hydrated with the 200 mM sucrose solution in 10 mM HEPES at pH 7.40. 4.2.3. Micropipette aspiration The micropipette aspiration apparatus was assembled based on the design described by Longo and Ly. 162 It consists of the micropipette that is placed in contact with the GUV within the sample chamber and through which suction pressure is applied by control of a hydrostatic manometer system. The micropipette was fabricated from a thin-walled borosilicate glass capillary (1 mm O.D., 0.75 mm I.D, Sutter Instrument) with the Narishige PE-21 glass microelectrode puller that uses gravitational force of the puller and adjustable magnet weights. In order to achieve gradual tapering of the pipette tip, relatively high heat and magnet weight settings were used. The pulled pipettes typically have a tip of approximately 1-3 µm, and the tip was cut and heat-polished to be 5-10 µm with the Narishige MF-900 microforge. Micropipettes were fabricated on the same day as aspiration experiments as the narrow glass tip easily attracted dust and debris that may clog the channel. Immediately prior to its usage to avoid evaporation and 82 introduction of air bubbles, each pipette was back-filled by attaching the blunt end to a piece of Tygon tubing (0.89 mm I.D.) that is attached on the other end to a 3-mL syringe (BD) filled with a 200 mM glucose solution in 10 mM HEPES buffer at pH 7.40. Once a back-filled micropipette has been prepared, it is attached to the tubing (1/32” I.D., 1/16” O.D., Tygon) that connects to the manometer system that is summarized in Figure 4.1. The manometer system consists of two water reservoirs, each filled with Mili-Q water (Milli-Pore) and connected to a differential pressure transducer (DP103-12-N-1-S-4-D, Validyne). One reservoir was used as reference, while the other was connected via tubing to the micropipette and its vertical position could be adjusted to induce pressure change. Note that two pressure transducers are available (for detection of separate ranges of differential pressure), but the current configuration only allows installation of one at a time; the DP103 model currently in use was labeled as P1. Each pressure transducer was connected to a carrier demodulator (CD17, Validyne), through which transducer readings could be received via a micro-USB connection and recorded in the comma-separate value format using data collection software from Validyne. 83 Figure 4.1. Schematic of the pressure control system and manometer. Shown are the placements of water reservoirs (R1 and R2), valves (V1, V2, V3, and V4), the pressure transducer (P1) and tubing. The sample holder frame was designed to fit on the Zeiss Axio Observer inverted microscope stage and was fabricated using a 3-D printer (MakerBot). The sample holder and its components are shown in Figure 4.2. The frame contains two parallel slits for placement of glass coverslips that were cut into proper lengths with a ceramic tile cutter (Sutter Instrument) from the 24 x 60 mm, No. 1 rectangular coverslips (VWR). The upper coverslip has a shorter length than the lower one to allow clearance for the micropipette to tilt downwards and submerge into the glucose solution. The glucose solution was held in place by adhesion to the two glass coverslips. When adding the GUVs to the sample chamber, 60 µL of the glucose solution (200 mM in 10 mM HEPES, pH 7.40) was first dispensed to the space in between the two coverslip by touching the micropipettor tip to the bottom of the upper coverslip to form a hanging water drop. Then, 30 µL of the GUV solution in sucrose (isoosmotic to glucose) was added to the glucose volume. The drop may touch the R1 R2 P1 V2 V3 V4 V1 to pipette 84 bottom glass coverslip to form a column of liquid at the center of the sample chamber, either before or after the addition of the GUV solution. Additionally, the top surface of the lower glass coverslip was scored gently around the center to prevent this column of liquid from moving, in case the sample chamber was tilted during transport to the microscope stage. Currently, this design is still flawed and incomplete, as significant evaporation of the liquid from the column occurred that severely affected the osmotic gradient across the GUV membrane. Figure 4.2. Micropipette aspiration sample holder. A 3-D printed frame (blue, dashed) with two slits hold rectangular coverslips (black and gray, thin lines); the upper coverslip (gray) has shorter length to allow room for the micropipette (black arrow, thick line) to tilt downwards from the micromanipulator. The GUVs are situated in the glucose solution (blue, circle) held in place by adhesion to both glass coverslips. During operation, valve V2 should first be opened while adjusting the height of both reservoirs together to match that of the micropipette opening. This was achieved when vesicles were no longer pulled into or pushed away from the micropipette tip. After the GUV was located and in focus and maintaining the micropipette as parallel to the coverslips as possible, the micropipette tip was located and brought close to the target GUV. Valve V2 was then closed to separate the two reservoirs and that the height difference between the two could then be detected by the differential pressure transducer. An applied suction of approximately 1 mm H 2 O was used to capture the GUV. Once both the GUV and micropipette tip were in focus, the micrograph was recorded. Note that since the micropipette was attached to 85 the tip of a holder that was held on the opposite end by a dovetail of the micromanipulator, it was prone to vibration. Thus, the micrograph data was recorded in series of 5 images at 2 ms exposure time. Successive recordings were made after lowering the reservoir R1 every 1 mm until close to the 15 mm H 2 O detection limit of the pressure transducer. 4.2.4. Data analysis The data obtained from micropipette aspiration experiments are series of micrographs; for one GUV, data is collected for at least four aspiration pressures (the first to be used as reference for calculation of relative areal strain and three more to use in linear fitting for the bending modulus). At each pressure, a series of five micrographs were recorded consecutively and examined for at least one that clearly showed the GUV membrane and inner micropipette edges. For the selected representative image at each imposed suction pressure, image processing was carried out using an in-house MATLAB program. First, the image of the GUV body (without the portion aspirated into the micropipette) was isolated by manually cropping the image to only show portions of the membrane edge. The built-in Canny edge detection function was then utilized to locate pixels corresponding to the GUV edge; these points were then used in conics fitting to extract the radius and center coordinates of the corresponding circle they comprise. For the determination of the micropipette inner radius, another section of the micrograph containing the portion of the micropipette with the aspirated membrane front was manually selected. The same edge detection function was used to locate the micropipette 2-D inner edges; user graphical input was utilized by allowing 86 manual selection of points on these upper and lower edges. Finally, the point corresponding to the depth of the aspiration apex was manually selected. The values obtained from the described image-processing program included the GUV and micropipette radii (R v and R P , respectively) and the aspiration length (L) (determined as the magnitude of the line connecting the aspiration apex with the predicted GUV surface with the slope averaged from those fitted to the upper and lower edges of the micropipette). The areal strain can then be calculated as α = 2πR P ΔL A 0 1- R P R V 4.2.4-1 where ΔL is the difference between the aspiration length of at the lowest (trapping) pressure and at a pressure value thereafter, and A 0 is the vesicle area of the GUV at this initial, lowest pressure value. With known applied pressure (detected by the pressure transducer), the induced membrane tension can then be calculated as τ = ΔPR P 2 1- R P R V 4.2.4-2 where ΔP is the pressure difference at the same conditions as described for ΔL. With calculated membrane tension and areal strain, the bending modulus (K C ) is determined using ln τ = 8πK C k B T α 4.2.4-3 where k B is the Boltzmann constant, and T is temperature in Kelvin. 162 With the natural logarithm of the tension values plotted as a function of strain, the bending modulus (as a multiple of k B T) is obtained from the slope of the fitted linear function. 87 4.3. Future Work In order to test the behavior of the newly built micropipette aspiration apparatus, control experiments were performed with DOPC and POPC for comparison of bending modulus values with available values from literature. The values of both compositions tend to be similar, so measurements with a saturated lipid, such as DMPC, should also be carried out. Prior to that, the problem of water evaporation from the glucose solution should be fixed for reproducible results between vesicles. Afterwards, the actual micropipette aspiration experiments with GUVs containing oxidized lipids will be performed. Since a usable population of GUVs can only be formed with up to 10 mol% POxnoPC, 161 this will be our limit of the maximum oxidized lipid concentration the bilayer. 88 Conclusions The studies presented in this dissertation demonstrate how oxidation alters the geometry of lipid molecules, which then affects holistic properties and disrupt the functionality of organized lipid phases. By understanding the impacts of oxidative damage done to individual molecules on the membrane, and consequently on organelles and the cell, we can begin to trace events that then lead to clinical presentations of aging-associated conditions. In this work, the studies of oxidative damage on lipid molecules emphasize on the simple and elegant relationship between form and function. The approach aids in visualization of the results of complex chemical processes and is rather suitable as orientation to the problems of aging and accumulation of oxidative stresses. This field is still largely unexplored, and humankind has so much to gain from mitigating oxidation-induced conditions. From the business side, the advertisement of products to reverse signs of aging and the marketing of a “better” antioxidant agent have never fallen out of favor with the general public consumers. However, my ultimate desire is for solutions to be readily available, such that treatments against conditions such as cancer would no longer be uphill battles with harmful side effects and less favorable chances of success. 89 References 1. Sankhagowit S, Wu S-H, Biswas R, Riche CT, Povinelli ML, Malmstadt N. 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Abstract (if available)
Abstract
Oxidation-associated conditions pose as major threats for the aging population. Beyond certain age milestones, these conditions are routinely screened for in order to be caught at earlier, more manageable stages. Their treatments are currently limited, as the understanding of mechanisms leading from biomolecule oxidation to cellular dysfunction and to manifestation of a disease is largely incomplete. While routine cell activities are usually convoluted with participation from many types of biomolecules, their failures can be unraveled by focusing on oxidative damage on lipid membranes that are the foundation of the cellular architecture. The bilayer membrane compartmentalizes the cell and its organelles and hosts a variety of membrane protein functions. Changes in its properties, such as lateral fluidity and continuity, can result in unsuccessful functional protein complex assembly and in leakage of vital nourishments. ❧ The research work presented in the following chapters focus on oxidation of membrane phospholipids. After an introduction with more details on how damages at the molecular level lead to well-studied aging-associated conditions, the dynamics of light sensitization-induced oxidation on a bilayer membrane composed of unsaturated lipids is analyzed, where the progress of lipid oxidation is monitored solely based on the overall membrane area changes. Oxidation results in increased polarity of the formerly hydrophobic tails, which alters the geometry of the molecule. Recognizing that the molecular intrinsic curvature can thus be affected by oxidative damage, the structural changes of a species utilizing such property in supporting bilayer membrane bending is then explored. Finally, the last study aimed to quantify the holistic membrane bending rigidity changes that arise from previously discussed structural changes to individual lipid molecules.
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Creator
Sankhagowit, Shalene
(author)
Core Title
The effects of oxidation on bilayer membranes studied using giant unilamellar vesicles
School
Viterbi School of Engineering
Degree
Doctor of Philosophy
Degree Program
Chemical Engineering
Defense Date
11/04/2015
Publisher
University of Southern California
(original),
University of Southern California. Libraries
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Tag
aging,giant unilamellar vesicles,lipid cubic phase,lipid packing parameter,membrane bending modulus,membrane curvature,micropipette aspiration,OAI-PMH Harvest,oxidation,oxidation kinetics,phospholipid bilayer
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English
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Electronically uploaded by the author
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Malmstadt, Noah (
committee chair
), Lee, C. Ted (
committee member
), Povinelli, Michelle L. (
committee member
)
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sankhagowit@gmail.com,shalene_sankhagowit@hotmail.com
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https://doi.org/10.25549/usctheses-c40-218428
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Sankhagowit, Shalene
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University of Southern California Dissertations and Theses
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Tags
giant unilamellar vesicles
lipid cubic phase
lipid packing parameter
membrane bending modulus
membrane curvature
micropipette aspiration
oxidation
oxidation kinetics
phospholipid bilayer