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Biophysical characterization of catonic liposome/plasmid DNA complexes for gene therapy Wasan, Ellen K. 1999

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B I O P H Y S I C A L C H A R A C T E R I Z A T I O N O F C A T I O N I C L I P O S O M E / P L A S M I D D N A C O M P L E X E S F O R G E N E T H E R A P Y B Y E L L E N K . W A S A N , R.Ph. B.S.Pharm., U N I V E R S I T Y O F H O U S T O N , 1992 A T H E S I S S U B M I T T E D I N P A R T I A L F U L F I L L M E N T O F T H E R E Q U I R E M E N T S F O R T H E D E G R E E O F D O C T O R O F P H I L O S O P H Y in T H E F A C U L T Y O F G R A D U A T E S T U D I E S (Department of Pathology and Laboratory Medicine) We accept this thesis as conforming to the required standard. T H E U N I V E R S I T Y O F B R I T I S H C O L U M B I A June, 1999 © Ellen Kathleen Wasan, 1999 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. Department of ?/?77V^&W f LrtPvtWTDl^ MM)rw The University of British Columbia Vancouver, Canada Date } DtU^n 1997 DE-6 (2/88) Abstract The goal of gene therapy is to achieve expression of an exogenous gene that results in a specific functional change. Cationic liposome-based carriers of plasmid D N A ("lipoplexes") are the most widely used method of nonviral D N A delivery. Lipoplexes form through electrostatic interactions between D N A and liposomes. This thesis investigates how lipid composition and ionic environment affect the biophysical behavior of lipoplexes and their transfection ability. A major concern regarding lipoplex development is their instability in a physiological environment or salt solutions. Lipoplexes containing a cationic l ipid and the transfection-helper l ipid dioleoylphosphatidylethanolamine (DOPE) exhibit heterogeneous morphology and variable activity. A charge ratio near neutrality or the presence of salts promotes super-aggregation (particle mean diameter > 1 um). Cryo-transmission electron microscopy showed that liposome morphology differs when plasmid, oligodeoxynucleotides or phosphate anions are added. In conjunction with l ipid mixing assays and particle size analysis, these observations demonstrate that charge ratio and charge density are critical for lipoplex structure. The extent of lipid mixing and aggregation, during or after lipoplex formation, is influenced by l ipid composition as well as the presence of salt or serum. A multi-step lipid-mixing assay to model in vitro transfection demonstrated that lipoplexes with relatively high in vitro transfection undergo lipid-mixing reactions after salt or serum interactions. Significantly, liposomal internal aqueous contents were retained in the lipoplexes. This may allow the codelivery of drugs within the lipoplexes during transfection of cells. Smaller, salt-stable lipoplexes are desirable for systemic in vivo use. It was also of great interest to pursue development of lipoplexes capable of trapping drugs for codelivery (e.g. transfection enhancers). Such lipoplexes must be transfection-competent, possess a large trapped volume and have the ability to maintain an ion gradient for drug loading. This was achieved by replacing D O P E with cholesterol and including poly(ethylene glycol) to stabilize the lipoplexes against aggregation. A model amine drug, vincristine, was loaded into lipoplexes via a p H gradient. This research introduces the novel concept that salt-stable lipoplexes can be generated 11 which can be loaded with secondary compounds for codelivery, adding a new functionality to cationic liposomes as carriers of DNA for gene therapy. Table of Contents Abstract i i Table of Contents iv List of Figures v i i i List of Tables x i Abbreviations x i i Acknowledgements xv Dedication xv i i i Chapter 1 Introduction 1 1.1 Effective D N A Delivery is Essential for Gene Therapy 1 1.1.1 Overview of Nonviral D N A Delivery Systems 3 1.1.2 Overview of Nonliposomal Gene Therapy 5 1.1.2.1 Polycat ion/DNA Complexes for Gene Transfer 5 1.1.3 Lipid-Based Gene Therapy 7 1.1.3.1 L i p i d / D N A Particles for Gene Transfer 10 1.1.4 Additional Formulation Strategies and Incorporation of Targeting Molecules 14 1.2 Liposomes as Carriers of D N A and Drugs 15 1.2.1 Liposome Preparation 16 1.2.2 Chemistry and Physics of Lipids 17 1.2.2.1 L ip id Polymorphism 17 1.2.2.2 L ip id Phase Behavior 20 1.2.2.3 Specific Lipids Used 21 1.2.2.3.1 Cationic Lipids 22 1.2.2.3.2 Phospholipids 23 1.2.2.3.3 Cholesterol 24 1.2.2.3.4 PEG-Lip ids 26 1.2.3 Entrapment of Drugs in Liposomes 28 1.2.3.1 Passive Drug Loading into Liposomes 28 1.2.3.2 Active Drug Loading into Liposomes 29 1.3 Liposome-DNA Complexes 35 1.3.1 Preparation of Lipoplexes 35 1.3.2 Physical Properties of Lipoplexes 36 1.4 Hypotheses of the Research and Specific Aims 36 Chapter 2 Materals and Methods 39 2.1. Introduction 39 2.2 Materials 39 2.2.1 Materials for Plasmid Preparation 39 2.2.2 Lipids 40 2.2.2.1 Cationic Lipids 40 2.2.2.2 Radiolabeled, Fluorescent and Anionic Lipids 40 2.2.3 Materials for Liposome Preparation. 41 2.2.4 Materials for Analysis of D N A Protection, Delivery and Transfection 41 2.2.5 Materials and Ce l l Lines for in Vitro Transfection ...42 iv 2.2.6 Other Mated als and Equipment 42 2.3. Methods for Preparation of Plasmid and Liposomes 43 2.3.1 Plasmids 43 2.3.1.1 Plasmid Isolation and Purification 43 2.3.1.2 Preparation of Radiolabeled Plasmid 45 2.3.2 Cationic Liposomes for Gene Transfer 46 2.3.2.1 Sizing of Liposomes and Lipoplexes 47 2.4 Formation of Lipoplexes 48 2.5 Characterization of Liposomal Carriers of D N A 49 2.5.1 Measuring Particle Size and Aggregation State 49 2.5.1.1 Sedimentation Assay 49 2.5.1.2 Salt-Induced Aggregation 50 2.5.2 Measurement of Liposomal Trapped Volume 50 2.5.2.1 Measurement of Trapped Volume by Filtration 51 2.5.2.2 Measurement of Trapped Volume by Dialysis 50 2.5.2.3 Determination of the Effect of Incubation with Tissue Culture Medium on Trapped Volume • 52 2.5.3 Cryo-Transmission Electron Microscopy to Study Lipoplex Morphology 52 2.5.3.1 Preparation of L iposome/Na 2 HP0 4 Mixture 52 2.5.3.2 C T E M Analysis 52 2.5.4 L ip id M i x i n g Assay 55 2.5.4.1 Multi-Step L ip id M i x i n g Assay 56 2.6 Characterization of D N A Protection 57 2.6.1 Sonication of Lipoplexes 57 2.6.2 DNase I Sensitivity 58 2.7 Expression of the Transgene 58 2.7.1 Measurement of Cellular Protein Concentration 59 2.7.2 fj-galactosidase 59 2.7.3 Chloramphenicol Acetyltransferase ( C A T ) 60 2.7.4 Luciferase 61 2.8 Methods of Loading Drugs into Liposomes and Lipoplexes 62 2.8.1 Measurement of p H Gradients 62 2.8.2 Loading of Ionizable Drugs into Cationic Liposomes or Lipoplexes 63 2.8.2.1 Chloroquine 63 2.8.2.2 Vincristine 64 Chapter 3 Cationic Liposomes Protect Plasmid DNA from Degradation by Ultrasonic Cavitation 65 3.1 Introduction .....65 3.3 Results 67 3.4 Discussion 76 Chapter 4 Cationic Liposome/DNA Complexes Retain a Significant Trapped Volume 81 4.1 Introduction 81 4.3 Results 82 4.4 Discussion 90 Chapter 5 The Role of Lipid Composition, Salts and Charge in Determining Lipoplex Structure 94 5.1 Introduction 94 5.2 Results 96 5.3 Discussion 108 Chapter 6 Lipid Mixing Behavior of Lipoplexes and Its Relation to In Vitro Transfection 113 6.1 Introduction 113 6.2 Results 116 6.3 Discussion 125 Chapter 7 Preparation of Salt-Stable Cationic Liposome/DNA Complexes and Coencapsulation of Drugs and DNA 132 7.1 Introduction 132 7.1.1 Theoretical Considerations on the Use of a Methylamine Gradient to Measure the p H Gradient 134 7.2 Results 137 7.3 Discussion 148 8. Summarizing Discussion 157 8.1 Summary of the Work 157 8.2 Physical Characterization of Lipoplexes Contributes to Advances in Gene Therapy 158 8.3 The Next Steps 159 8.4 The Future of Gene Therapy 161 8.5 Conclusions 163 9. References 165 Appendix: Analytical Methods for the Characterization of Lipid-Based DNA Carriers 190 A . l Introduction 190 A . 2 Materials 190 A.2.1 Additional Cationic Lipids 190 A . 3. The Active Components 191 A.3.1 Plasmids 191 A.3.1.1 Plasmid Isolation and Purification 191 A.3.1.2 Preparation of Radiolabeled Plasmid 196 A.3.1.3 Preparation of Fluorescently Labeled Plasmid 196 A.3.2 Cationic Liposomes for Gene Transfer 198 A.3.2.1 Preparation of Cationic Liposomes 198 A.3.2.2 Formation of Lipoplexes 199 vi A.3.3 Characterization of Lipid-Based Gene Transfer Formulations 200 A.3.3.1 Measuring Particle Size and Aggregation State 200 A.3.3.2 L ip id M i x i n g Potential 201 A.3.3,3 Carrier-Membrane L ip id M i x i n g 204 A.3.3.4 D N A Protection 205 A.3.3.4.1 Fluorescent Dye Labeling of Plasmid 206 A.3.3.4.2 DNase I Sensitivity 207 A.3.3.4.3 Serum Stability 207 A.3.4 Analysis of Ce l l Delivery 208 A.3.4.1 Plasmid and L ip id Delivery 210 A.3.4.1.1 Quantification of Plasmid from Transfected Cells 211 A.3.4.1.2 Analysis of Plasmid D N A Integrity 214 A.3.4.1.3 Analysis of L ip id Delivery 216 A.3.5 Expression of the Transgene 217 A.3.5.1 Evaluation of Transfection Using Reporter Genes 218 A.3.5.1.1 P-galactosidase 219 A.3.5.1.2 Chloramphenicol Acetyltransferase ( C A T ) 220 A.3.5.1.3 Luciferase 220 A.3.5.2 Gene vs. Message vs. Product 221 A.4 Summary 222 vii List of Figures Figure Title Page 1.1 In vitro transfection is a multi-step process 4 1.2 Chemical structures of cationic lipids used in lipid-based D N A carriers 10 1.2a D O D A C , D D A B and D O T M A 10 1.2b D M R I E , G A P - D L R I E , D C - C h o i and D O G S 11 1.2c D O T A P , D O T I M and D O S P A 12 1.3 Structures of phospholipids used in liposomes ( D O P C , D O P E and DOPS) 13 1.4 L ip id bilayer and liposomes 15 1.5 Shape concept of l ipid polymorphism 18 1.6 The hexagonal phase (HU) 19 1.7 Gel to liquid crystalline lipid phase transition 20 1.8 Chemical structure of cholesterol 25 1.9 Chemical structure of D S P E - P E G 26 1.10 Encapsulation of ionizable drugs by a p H gradient 31 1.11 Encapsulation of drugs in liposomes using an ammonium sulfate gradient 32 1.12 Encapsulation of drugs in liposomes using a redox system to generate a p H gradient 33 2.1 Chemical structures of rhodamine-PE and N B D - P E 40 2.2 Controlled environment vitrification chamber for C T E M sample preparation... .53 2.3 C T E M sample forms a film over the holey polymer matrix 52 2.4 The C T E M image is in 2D but the sample is in 3D 54 3.1 Effect of formulation and charge ratio on lipoplex size 68 3.2 Agarose gel electrophoresis for assessment of D N A integrity 73 viii 3.3 In vitro transfection of C H O cells with sonicated vs. unsonicated lipoplexes 75 4.1 The effect of binding to D N A on liposomal trap volume 83 4.2 Lipoplex aggregation and lipid mixing depends on charge ratio 85 4.3 Trapped volume of lipoplexes depends on l i p i d : D N A ratio 87 4.4 Lipoplex aggregation is associated with l ipid mixing 88 4.4A Sedimentation assay 88 4.4B L ip id mixing assay 88 4.5 The effect of tissue culture medium on trapped volume 90 5.1 Proposed model of l iposome/DNA complexes 95 5.2 The influence of polynucleotide concentration on lipoplex size 99 5.2A Liposomes and plasmid D N A . 99 5.2B Liposomes and oligodeoxynucleotides 99 5.2C Liposomes and Na 2 HP0 4 99 5.3 The influence of polynucleotide concentration on lipoplex sample turbidity.. ..100 5.3A Liposomes and plasmid D N A 100 5.23 Liposomes and oligodeoxynucleotides 100 5.2C Liposomes and N a 2 H P 0 4 100 5.4 C T E M of D O D A C / D O P E liposomes and lipoplexes in lactose 102 5.5 C T E M of D O D A C / D O P E liposomes and lipoplexes in H B S 104 5.6 C T E M of D O D A C / D O P C liposomes and lipoplexes in lactose 106 5.7 C T E M of D O D A C / D O P C liposomes and lipoplexes in H B S 107 5.8 Proposed mechanism for lipoplex polymorphism 110 6.1 L ip id mixing of cationic liposomes induced by polynucleotides of Na2HP04... 117 6.2 Multi-step l ipid mixing assay 121 ix 6.2A Multi-step lipid mixing assay without serum 121 6.2B Multi-step lipid mixing assay with serum 121 6.3 In vitro transfection of B16 cells for comparison to lipoplex lipid mixing properties 123 6.3A C A T assay 123 6.3B Luciferase assay 123 7.1 Cholesterol-containing cationic liposomes impart DNase I protection 138 7.2 Transfection of B16 cells with lipoplexes prepared in H B S 141 7.3 Ion gradients of DODAC/DOPC/choles te ro l /PEG lipoplexes 145 7.4 Loading of vincristine into liposomes via a p H gradient 147 7.5 Schematic illustration of three possible fates of liposomal entrapped contents after interaction with D N A 150 7.6 Chemical structures and properties of chloroquine and vincristine 154 8.1 Advances in molecular biology hold the key to gene therapy 162 x List of Tables Table Title Page 1 Nonviral D N A carriers in development 4 2 L ip id phase transition temperatures (Tc) 21 3 Drugs that have been encapsulated by active loading procedures 34 4 Effect of l ipid composition on lipoplex size 69 5 Effect o f sonication on lipoplex size 70 6 Semiquantitation of D N A integrity following lipoplex sonication 74 7 Effect of N a C l on liposome size with and without added plasmid 96 8 Cationic liposomes and lipoplexes containing cholesterol and P E G are stable in 150 m M N a C l 138 9 Trapped volume of liposomes and lipoplexes in H B S 143 10 Comparison of liposome uptake between drugs with differing physico-chemical properties 155 Abbreviations B C A , bicinchoninic acid P-gal, beta galactosidase C A T , chloramphenicol acetyltransferase C H E , cholesteryl hexadecyl ether C H O cells, Chinese hamster ovary cells Choi, cholesterol C M V , cytomegalovirus C o A , coenzyme A cpm, counts per minute C T E M , cryo-transmission electron microscopy D C - C h o i , cholesteryl 3p-[N-dimethylaminoethyl] carbamate D D A B , dimethyldioctadecylammonium bromide D M P E - P E G , monomethoxypoly(ethylene glycol)2ooosuccinate-dimyristoylphosphatidylethanolamine D M R J E , dimyristyloxypropyl-3-dimethyl-hydroxyethyl ammonium D M S O , dimethylsulfoxide D N A , deoxyribonucleic acid DNase, deoxyribonuclease D O D A C , dioleyldimethylammonium chloride D O G S , dioctadecylamidoglycyl spermidine D O P C , dioleoylphosphatidylcholine D O P E , dioleoylphosphatidylethanolamine D O P S , dioleoylphosphatidylserine D O S P A , 2,3-dioleyloxy-N[2-(sperminecarboxyamido)ethyl]-N,N-dimethyl-l-propanaminium trifluoroacetate D O T A P , dioleoyl-l,2-diacyl-3-trimethylammonium propane D O T M A , N-[l-(2,3-dioleoyloxy)propyl]-N, N , N-trimethylammonium chloride dpm, disintegrations per minute D S P E - P E G , monomethoxypoly(ethylene glycol)2ooosuccinate-distearoylphosphatidylethanolamine EDTA, ethylenediamine tetraacetic acid E P C , egg phosphatidylcholine F A C S , fluorescence-activated cell sorting F B S , fetal bovine serum g, gravity G M P , Good Manufacturing Procedures HIT, hexagonal phase H B S , N-(2-hydroxy-ethyl)piper-azine-N'-(2-ethane-sulfonic acid)("HEPES")-buffered saline L B , Luria broth Lipoplex, cationic liposome/plasmid D N A complex L M , l ipid mixing L U V , large unilamellar vesicle L a , liquid crystalline phase L p , gel phase max, maximum m R N A , messenger ribonucleic acid M L V , multilamellar vesicle mol%, molar percentage M W , molecular weight NJBD-PE, N-7-nitrobenz-2-oxa-l,3-diazol-4-yl)-l,2-dihexadecanoyl-5«-glycero-3-phosphoethanolamine O D , optical density O G P , octyl glucopyranoside Oligo, oligodeoxynucleotide(s) P C , phosphatidylcholine p C M V p , plasmid encoding Eschericia coli P-galactosidase P C R , polymerase chain reaction P E , phosphatidylethanolamine P E G , poly(ethylene glycol) P G , phosphatidylglycerol PS, phosphatidylserine psi, pounds per square inch Q E L S , quasi-elastic light scattering xiii R E S , reticuloendothelial system R E T , resonance energy transfer technique R N A , ribonucleic acid RNase, ribonuclease Rh-PE, N-rhodamine phosphatidylethanolamine rpm, revolutions per minute R T , room temperature R V t , residual liposomal trapped volume SDS, sodium dodecylsulfate S U V , small unilamellar vesicle SV40, simian virus 40 T O - P R O , quinolinium, 4-[(3-methyl-2(3H)-benzothiazolylidene)methyl]-1 -[3-(trimethylammonio) propyl]-, diiodide U , units of activity Vt , liposomal internal aqueous trapped volume Y O - Y O , quinolinium, 1, V-[ 1,3-propanediylbis[(dimethyliminio)-3,1 -propanediyl]]bis[4-[(3-methyl-2(3H)-benzoxazolylidene)methyl]]-,tetraiodide xiv Acknowledgements I would like to thank my supervisor, Dr. Bal ly , for all his guidance, extreme patience, and for allowing me a great deal of independence in the direction of my research project. Sometimes that wasn't the quickest way to get to the answer we were looking for, but it was a valuable part of the process of becoming a Ph.D. Marcel 's generosity to all o f us working in the lab was certainly much appreciated. He and senior scientist Dr. Lawrence Mayer set the tone of Advanced Therapeutics to be a friendly and fun yet productive place to work. Somehow Marcel transmitted his high expectations without appearing to pressure the students. Sending me to Sweden to learn C T E M was an amazing experience for which I w i l l always be grateful. With his support, I was able to present my research at international gene therapy and pharmaceutical meetings every year. I guess it was the Texas connection that helped us get along so well , although I never did get those tortillas... I would also like to acknowledge support from Inex Pharmaceuticals, for materials, technical assistance, and Dr. Michael Hope of Inex for participating as the industrial collaborator for the Science Council of B C Graduate Research and Engineering Training fellowship. The provision of lipids and plasmid made the work significantly easier. Many thanks also, to former postdoctoral fellows Drs. Dody Reimer and Y . P . Zhang. Their assistance in the lab and inspiration through frequent discussions was invaluable. Their suggestions were constructive, helping me to stay on track or dig a little deeper. Thanks to former graduate students Howard L i m , Troy Harasym and Shane Longman for teaching me how to make liposomes and Rajesh Krishna for listening. Dr. Mayer was always wil l ing to give advice for troubleshooting experiments. I appreciated his open door policy and that both his group and ours worked together on so many projects. The personnel of the Drug Development Unit - Jean Heggie, Norma Hudon, Ginette St. Onge, Daria Hartley and Carole Dedhar were invaluable resources for analytical methodology and procedural advice. Spencer Kong was also a tremendous font of know-how on a seemingly endless number of subjects, an essential part of the Advanced Therapeutics team. I would also like to thank Dr. Stanton Gerson of Case Western Reserve University, Departments of Hematology/Oncology and Pharmacology, Cleveland, Ohio. Dr. Gerson was my first supervisor as a graduate student, who provided a wonderful example of a successful clinician-scientist and who always made time for in-depth scientific discussions with his students. Even earlier than that, while in pharmacy school, I first learned about the research world from Drs. Gayle Brazeau of the University of Houston, Department of Pharmaceutics and XV Gabriel Lopez-Berestein of U . T. M . D . Anderson Cancer Center in Houston, T X , Department of Medical Oncology. In their laboratories I first experienced the thrills of science. They both always believed in me and encouraged me to go as far as I could, which meant a great deal to me. Finally, the B C Cancer Agency was a unique working environment, at which I observed how basic science, clinical science and industry can come together to advance the treatment of human disease. Many other individuals are not listed by name here, but in many ways influenced my experience of graduate school or inspired my efforts to become a scientist. No one can do scientific research without the involvement of other people on a variety of levels. Several of the chapters of this thesis note that other individuals merited authorship on the publications derived from the work presented in those chapters besides myself and my supervisor, Dr. Marcel Bal ly . While I wrote the papers myself (with revisions from Dr. Bally), the other authors provided valuable input also. For the manuscript acknowledged in Chapter 3, Dr. Dorothy Reimer established cationic liposome-based transfection assays in our laboratory and as a post-doctoral fellow helped start the gene therapy program in the Department of Advanced Therapeutics. For trapped volume studies described in Chapter 4, A lysa Fairchild was a diligent summer student who performed many of the trapped volume measurements under my direction. Chapter 5 acknowledges Dr. Katarina Edwards o f Uppsala University in Uppsala, Sweden as wel l as her technician M r . Goran Karlsson. I spent a month in Dr. Edward's laboratory learning about cryo-transmission electron microscopy under their excellent tutelage (tak sa mickett!). They incurred considerable expense in time spent with me and in materials for operating the equipment, in exchange only for an academic collaboration and a publication. Dr. Edwards contributed intellectually to the interpretation of the images and in the manuscript development. The other two authors on that manuscript are post-doctoral fellow Dr. Pierrot Harvie and graduate student Frances Wong, who taught me how to make their special l i p i d / D N A particles and prepared materials for the C T E M . Both of them reviewed the manuscript, which is the same one acknowledged in Chapter 6 of this thesis. Dr. Harvie also kindly performed the in vitro C A T assays described in Chapter 6 using lipoplexes I prepared. We had a number of interesting discussions regarding the development and interpretation of the l ipid mixing assay for l i p i d / D N A delivery systems. Chapter 7 has not been transformed into a publication as of this writing, however, I wish to acknowledge the contribution of postdoctoral follow Dr. Margaret Wong, who performed the vincristine uptake studies in Figure 7.4. She used liposomes and protocols I developed for the salt-stable formulations and standard operating procedures of the xvi Drug Development Unit of the B C Cancer Agency for the controls. She also did several additional related experiments for me (with M s . Dana Masin), not included in this thesis but important for the eventual publication of the work. Finally, the Appendix is taken in part from a review article I wrote for Methods in Molecular Medicine with contributions from a number of individuals already mentioned above. They either wrote short segments on topics relevant to their work, or provided me with technical information, opinions or references. A l l authors reviewed the manuscripts in detail. XVII Dedication To my husband, Kishor, for his unwavering love and support. To my Dad, for instilling in me a strong work ethic and the sense that anything worth doing is worth doing right. To my M o m , for always encouraging me to go as far as I can and for her delight in my successes at every stage of my life. Good, better, best never let it rest until your good is better and your better, best. (a phrase often recited to me as a child by my grandfather, George V . Laco, which to this day rings through my head whenever I am tempted to give something less than my best effort) I feel that the more we learn about the amazing power of life processes and the intricate way the physical world is constructed, the more God's glory is revealed to us. Praise to our Creator! xviii Chapter 1 Introduction 1.1 Effective DNA Delivery is Essential for Gene Therapy The overall goal of gene therapy is to cure or stabilize a disease process that results from the over- or underexpression of a specific gene. We can achieve this goal by replacing the defective gene, or by reducing the overexpression of the target gene using an antisense strategy (Sokol and Gewirtz, 1996, Roth and Cristiano, 1991). It is critical to transfer D N A into target cells in the appropriate conformation and concentration to be effective in modifying the disease. D N A must be delivered to the desired cell population in an intact state. For gene therapy the D N A must be efficiently transcribed and ultimately translated appropriately in the host cell. For antisense therapy, the D N A must reach the host target complementary R N A or D N A sequences to block gene expression. The method of gene transfer must be highly efficient, nontoxic, relatively easy to prepare and to administer (Bertling et al, 1991). There is a great deal of optimism surrounding the development of gene therapy as an effective strategy for management of many different human diseases. The active agent used to procure gene therapy is likely to consist of oligonucleotides, ribozymes or a D N A sequence that can be transcribed into a message capable of eliciting a therapeutic response. Unlike conventional small molecule therapeutics however, gene therapy requires the use of a carrier system to deliver the active agent directly into the target cell population. Viruses are the most common gene therapy vectors. Viruses have the ability to infect cells containing the appropriate receptors very efficiently. Most clinical gene therapy trials to date utilize viruses to deliver D N A (Ledley et al, 1995, Roth and Cristiano, 1997). Nonviral D N A delivery methods, however, are growing in popularity due to several distinct advantages. The D N A used for nonviral-mediated gene transfer is usually in plasmid form, for which the vector size restrictions are much less limiting than for most viral-based systems. Plasmids are 1 closed, circular, double-stranded D N A of bacterial origin that can be modified to encode the gene of interest and to allow transcription in mammalian cells. Some plasmids do not usually integrate into the host genome but rather remain episomal, greatly reducing the potential for insertional mutagenesis or modification of the transgene by host cell factors (such as methylation). Plasmids are not potentially infectious agents, unlike viruses, which could become infectious i f complementation occurs in the host (Cooper et al, 1996, Ledley et al, 1995, Tomlinson and Rolland, 1996). For effective uptake into most tissues, plasmid requires the use of carrier molecules, such as cationic lipids (Nabel, E . et al, 1992, Nabel, G . et al, 1992, Canonico et al, 1994) or cationic polymers (Bogdanov et al, 1993, Chancerelle et al, 1993). These have the advantage of low antigenicity (permitting multiple administrations i f necessary) and low toxicity. Targeting molecules possibly can be incorporated into the DNA/carrier complex to improve specificity of delivery. Viruses, on the other hand, may have limited trophism which is determined by the presence of specific viral receptors on the cell surface. Numerous approaches are available to prepare effective lipid-based D N A delivery systems. In general, however, the formulations usually involve cationic (positively charged) liposomes (lipid vesicles), or sometimes monomelic cationic lipids, bound to the D N A ("lipoplex"). The charged components bind D N A directly (such as cationic l ipid binding to the anionic D N A molecule) or in some cases indirectly (anionic l ipid binding to a cationic peptide/DNA complex). Lipid-based carriers are proving to be a versatile and pharmaceutical^ viable technology. The process of D N A transfer into cells is facilitated by the physical and chemical attributes of the lipoplex, which insure that D N A is maintained in a form that is protected from enzymatic hydrolysis and that it is readily taken up by the cells. However, many D N A binding molecules can fulfill these rather non-specific functions such as agents which "condense" (compact) the D N A . A s a flexible loop, plasmids can adopt many different conformations. The ones most commonly described are supercoiled, relaxed (one strand nicked) 2 and linear (both strands cut). In addition, certain cations, particularly divalent cations such as C a 2 + and M g 2 + , can disrupt hydrogen bonding between the D N A molecule and water molecules, resulting in a "condensation" of the D N A . The molecule coils tightly in upon itself in a characteristic toroid manner (Bloomfield et al, 1990). It has been suggested that cationic lipid may also condense D N A , but lipids may play additional roles in the gene transfer process. Besides D N A protection during delivery and the role of lipids in membrane fusion, lipoplex components may also need to possess the ability to destabilize condensed DNA/carrier structures once arriving at the cellular destination. Without a means of unpackaging the D N A from the carrier after cellular delivery, the D N A w i l l not be transcribed and no product w i l l result. Regardless of the strategy, it is imperative to understand the mechanism(s) involved in the delivery process. Targeting of D N A to a specific cell population to achieve gene therapy by any method can only be optimized when the components of the delivery system are fully characterized. This w i l l ultimately enable researchers to effectively assess the efficiency of D N A delivery and cellular uptake, critical parameters in gene transfection. This thesis addresses many of the current issues related to the development and analysis o f cationic l iposome/DNA complexes, the most promising nonviral D N A delivery method to date. 1.1.1 An Overview of Nonviral DNA Delivery Systems While the specific physico-chemical properties of nonviral plasmid carriers w i l l vary, the goal is the same: to produce stable particles in a highly reproducible manner, which promote efficient cellular uptake of the D N A . Table 1 lists the major classes of nonviral D N A carriers and their primary attributes. Small particle size (<200 nm) is required for effective systemic biodistribution. The DNA/carrier complex must be stable during sample preparation and dilution into standard buffers. The D N A also must be protected from rapid degradation, either in vitro or in vivo, yet must be released from the carrier at the appropriate time within the cell for it to be actively transcribed (Figure 1.1). 3 Class Examples Features Lipids Cationic liposomes Easy to prepare, but low and variable activity, heterogeneous particles, must be optimized empirically Lipid/DNA particles Activity equivalent to cationic liposomes, may be suitable for systemic use due to small particle size Polymers Lipopolyamines, polylysine May have excellent DNA-condensing properties, but still requires empirical optimization for transfection. Dendrimers Surface chemistry controllable, excellent D N A condensing ability, variable activity Peptides Polylysine derivatives Low activity, potential for immunogenicity Nuclear localization peptides as adjuvant Appears to aid activity but mechanism unclear pH-sensitive peptides as adjuvant for lipid- or polymer-based carriers Promote endosomal release of D N A . Increase in activity minimal Combinations D N A condensed by polymers, bound to cationic lipid and attached to targeting molecules or antibodies Preparation complicated, increase in activity minimal Table 1 Classes of nonviral D N A carriers presently under development Figure 1.1 Schematic representation o f the multi-step transfection process A . Lipoplex Preparation B. Dilution I 0 n o X o « °— - ° o o ° Plasmid D N A V Liposomes Lipoplex 1. lipoplex delivery ( J . Cellular Interactions ^-0 3 . degradation O in lysosomes *> 2. endocytic uptake SB. translation 3. release from endosomes jiranscriptiom wcfeus $4 nuclear entry cytosol lipoplex plasmid DNA messenger RNA protein product 4 The inclusion of targeting moieties on the surface of the carrier is another possibility to improve activity. While potentially improving the specificity of delivery, this approach also increases the complexity. The physiological attributes of the tissue in which one wishes to achieve D N A expression are important parameters when considering in vivo administration. A number of pharmaceutical factors such as carrier aggregation state and stability, reproducibility of preparation, sterilization and scale-up o f the production for manufacturing purposes are also ongoing challenges. 1.1.2 Overview of Nonliposomal Gene Therapy The carriers for D N A in the nonviral delivery systems currently include not only cationic liposomes (Feigner et al, 1987, Farhood et al, 1994, Caplen et al, 1995), but also nonliposomal l i p i d / D N A particles (Gao and Huang, 1996, Zhang et al, 1996), lipopolyamines (Behr et al, 1989) and other polycations (Barthel et al, 1993, Kabanov and Kabanov, 1995, Wolfert et al, 1996). These polycations including polylysine (Cotton et al, 1990, Wagner et al, 1990, Zenke et al, 1990, Zhou et al, 1991), other polyvalent polymers such as the dendrimers (Haensler and Szoka, 1993, Tang et al, 1996, Bielinksa et al, 1996), cationic or amphipathic peptides (Kato et al, 1996, Legendre et al, 1997), and combinations of these (Hong et al, 1997) (Table 1). 1.1.2.1 Polycation/DNA Complexes for Gene Transfer Polyvalent cations used in D N A delivery can cause a type o f condensation o f D N A , in which the D N A becomes inaccessible to D N A intercalating fluorescent dyes like ethidium bromide. Polymers such as polylysine and dendrimers have been shown to condense D N A , and in the case of dendrimers, to promote transfection at least as well as the cationic l ipid delivery systems. Recently this class o f D N A carriers has become known as "polyplexes, or "lipopolyplexes" i f they contain both lipids and polymers. It has been observed in some studies that D N A condensation by the carrier lipids or polymers results in improved transfection (Wu, G . and W u , C , 1988, Wagner et al, 1991). Condensation may impart a protective effect against nucleases and possibly improve its eventual 5 activity within the cell. Polylysine can be covalently coupled to targeting peptides, as discussed later, to achieve improved specificity of uptake. Antigenicity o f polylysine is not anticipated to be a concern. Polylysine is used as a component of the microencapsulation system used to protect live cells in allogeneic transplantation from immune attack (L iu et al, 1993, Halle et al, 1993, Okada et al, 1997). Its use as a D N A carrier should not induce an immune response. Dendrimers, highly branched synthetic polymers with a spherical shape, are currently under development for a variety of applications. The size, shape and surface features of these macromolecules can be readily controlled (Tomalia et al, 1990, Tomalia et al, 1993). Polyamidoamines, also known as Starburst™ dendrimers (Dendritech, Detroit MI) or P A M A M , in particular have been used as gene transfer vehicles (Haensler and Szoka, 1993). They have a large charge density due to their terminal amine groups, which makes them amenable to D N A complexation. Dendrimers have been demonstrated to be at least as efficient as cationic liposomes in enhancing transfection of a variety of cell types, and they have been reported to be less toxic. Transfection efficiency is affected by the size of the polymer sphere and the ratio of polymer to D N A . The current hypothesis about their mode of action is that, following endocytosis, the titratable amines of the polymer buffer the acidic environment of the endosome, preventing degradation of the plasmid by pH-dependent nucleases. Furthermore, i f enough flexibility is present in the polymer molecule, such as when it is pre-treated with heat, it w i l l swell in the endosome, disrupting the organelle and promoting D N A escape (Tang et al, 1996). This swelling may be due to an increase in hydrodynamic radius as more terminal amines become protonated in the low p H environment and thus increase hydrogen bonding with water. Cationic peptides have also been used as D N A carriers. In a strategy developed by Legendre and Szoka (1993), the cationic peptides gramicidin S and tyrocidine were combined with D O P E (5:1 lipid:peptide ratio). Complex formation was promoted by simple mixing of the peptide with D N A followed by addition of D O P E . For transfection, the optimal molar ratio of 6 DOPE:peptide was approximately 4.5:1. The efficiency of the peptide compared to lipoplexes varies by cell type, but the low toxicity is similar. In another study, cationic peptides were made more lipophilic by chemically attaching a fatty acyl group to one of the peptide amino groups. This molecule was used to prepare peptide/DNA particles similar to l iposome/DNA complexes (Ohmori et al, 1997). Modified peptides have also been used like cationic lipids; N , N -dihexadecyl-N a-[6-(trimethylamino)-hexanoyl]-L-alaninamide bromide was used to make sonicated liposomes that formed a complex with plasmid D N A . COS-7 cells were transfected with this complex in vitro with greater efficiency and less toxicity than conventional cationic l iposome/DNA complexes (Kato et al, 1996). Other types of cationic peptides have also been utilized for gene transfer. They are most effective in combination with molecules that exhibit pH-dependent membrane-perturbation effects (Legendre, et al, 1997, Ohmori et al, 1997). Presumably, these helper components promote endocytic escape after cellular uptake. Cationic peptide-type carriers are not in wide usage at this time, particularly in vivo. It w i l l be interesting to see i f these peptides induce any immune response when administered to animals. 1.1.3 Lipid-Based Gene Therapy The most frequently cited method of nonviral D N A transfer is that employing cationic lipid carriers. This idea was based originally on experience with liposomal delivery of conventional small molecule drugs, and on the need to improve the interaction of the D N A with the cell (Feigner et al, 1997). In the late 1970s, several investigators managed to encapsulate D N A into anionic or neutral liposomes and achieved delivery of the D N A into cells (Fraley et al, 1980, Wong et al, 1980). Liposomes and their physical properties, including drug encapsulation, are discussed in Section 1.2. Plasmids, however, do not encapsulate well into liposomes because of their size and charge, and neutral liposomes are not taken up into cells avidly. The major advance in this field was the introduction by P. Feigner in 1987 of cationic liposomes that bind D N A through electrostatic interactions, 7 forming a complex ("lipoplex"). Lipoplexes interact readily with the negatively charged plasma membrane of eukaryotic cells and thus improve cellular uptake o f the D N A . Since that breakthrough, lipoplexes have been used for gene transfer for a variety of tissues and cell types. They have been administered intravenously (Lesoon-Wood et al, 1995, Lew et al, 1995), intraperitoneally (Yu et al, 1995, X i n g et al, 1996, Reimer et al, 1999), intratracheally (Hyde et al, 1993, Meyer et al, 1995, McLachlan et al, 1996) and intraocularly (Jones et al, 1994). Lipoplexes have been injected locally into tumors (Parker et al, 1996, Egilmez 1996) or solid organs (Schmid et al, 1994, Hickman et al, 1994) and in various animal species (Canonico et al, 1994, Leibiger et al, 1990, Thierry et al, 1997). They have also been administered to humans for the treatment of cystic fibrosis (Caplen et al, 1995) and melanoma (Nabel et al, 1996). Numerous clinical trials are ongoing worldwide employing cationic l ipid-based D N A carriers for gene therapy. Transfection with lipoplexes tends to be less efficient than with viral vectors, and shares with many viral vectors a transient and variable level o f expression for most tissues. Transfection with lipoplexes is considerably greater than plasmid in the absence of the carrier. To improve the efficiency of cationic liposome-mediated transfection, many groups have spent considerable effort developing new cationic lipids (Feigner, J. et al, 1994, Feigner, P. et al, 1995, Balasubramaniam et al, 1996, Wheeler et al, 1996, Bennett et al, 1998). Others have explored the effect o f "helper lipids," such as dioleoylphosphatidylethanolamine (DOPE) (Farhood et al, 1995). D O P E enhances the transfection ability o f some cationic lipids, supposedly by improving the fusogenicity or endosomolytic activity o f the liposome component. These functions would allow D N A to escape into the cytoplasm. In spite o f their usefulness for transfection in vitro or for local administration in vivo, conventional cationic l iposome/DNA complexes may not be suitable for systemic (intravenous) use. One of the major reasons is because they have a tendency to undergo aggregation, resulting 8 in large, heterogeneous particle sizes (> 1 um) (Wasan et al, 1996, Wasan et al, 1999). This is particularly true in ionic solutions, an observation characterized in detail throughout this thesis. More recently, small and homogeneous l i p i d / D N A particles have been produced by detergent dialysis methods. L i p i d / D N A particles have different physical properties than cationic l iposome/DNA complexes, such as improved stability, but no better transfection ability (Zhang et al, 1996, Harvie et al, 1998). These particles are discussed further in section 1.1.3.1. For preparation of gene transfer vehicles the liposomes are pre-formed, then mixed in a defined ratio with the nucleic acid for rapid, spontaneous formation of l iposome/DNA complexes ("lipoplexes"). Cationic lipids that are typically used derive their positive charge from a modified amine or amide headgroup. Those useful for transfection include but are not limited to: D O T M A , D O S P A , D O T A P , D D A B , D M R I E , D C - C h o i , D O G S , D O D A C (see Figure 1.2 for chemical names and structures, and the Appendix for sources) and an ever-increasing number of variants of these older lipids. Helper lipids such as D O P E (Figure 1.3) are often included with certain cationic lipids in liposome formulations, generally at 50/50 m o l % with the cationic lipid. A s mentioned above, D O P E is thought to act by promoting endosomolysis upon transfection through its ability to promote membrane fusion (Farhood et al, 1995, H u i et al, 1996) (see Chapter 5), although the exact mechanism remains controversial. Some cationic lipids, such as D O G S , do not seem to require D O P E to achieve transfection (Balasubramaniam et al, 1996, Wheeler et al, 1996). The lipid to be used for a given transfection experiment remains somewhat an empirical choice at this time because the difference between the effective cationic lipids overall appears to be slight. Derivatives of the older cationic lipids continue to be synthesized, with reports of improved transfection efficiency in a few cell lines. Further investigation is required to determine i f these derivatives offer any significant benefit over the parent compounds in a variety of cell types and experimental conditions. 9 1.1.3.1 Lipid/DNA Particles for Gene Transfer L i p i d / D N A particles represent a non-liposomal but lipid-based delivery system for gene transfer. Monomeric or micellar lipids interact with D N A in the presence of detergent or some other surface-active agent that is later removed by dialysis. A s the surface-active agent diffuses out, solid, condensed particles of l ipid and D N A form (Zhang et al., 1996). These can be prepared such that they are smaller and more homogeneous than l iposome/DNA complexes, however, transfection is no better (Harvie et al., 1998). In one method of making l i p i d / D N A particles a lipid film is first made consisting of cationic and neutral lipids. Hydration is performed with a nonionic detergent solution, such as 30 m M octyl- glucopyranoside (OGP). Further characterization of l i p i d / D N A particles w i l l be required to optimize their transfection ability. dioleyldimethylammonium chloride (DODAC) ci-dimethyl dioctadecylammonium bromide (DDAB) \ B r N[l-(2,3-dioleyloxy)propyl]-N,N,N-trimethyl-ammonium chloride (DOTMA) Cl-+ N Figure 1.2a Chemical structures of cationic lipids used in lipid-based D N A carriers 10 l^-dimyristyloxypropyl-3-dimethyl-hydroxy ethyl ammonium bromide (DMRIE) N-(3-aminopropyl)-N,N-dimethyl-2,3-bis(dodecyloxy)-1-propanimium bromide (GAP-DLRIE) 3b- [N-(Dimethylaminoethane)-carbamoyl] -+NH3 Figure 1.2b Chemical structures of cationic lipids used in lipid-based D N A carriers 11 2-(octadecenoyloxy)ethyl-2-(heptadecenyl)-3 -[hydroxyethyljimidazolinium chloride (DOTIM) O ci-Figure 1.2c Chemical structures of cationic lipids used in lipid-based D N A carriers 12 dioleoylphosphatidylcholine (DOPC) o—p-i Fatty Acyl Chain Glycerol Phosphate Backbone Headgroup dioleoylphosphatidylethanolamine (DOPE) O dioloeoylphosphatidylserine (DOP S) Figure 1.3 Structures of phospholipids used in liposomes When preparing l i p i d / D N A particles it is important that the detergent concentration be above the critical micelle concentration ( C M C ) . Equal volumes of plasmid D N A and the mixed detergent/lipid micelles are mixed by pipetting one into the other, followed by extensive dialysis of the mixture to remove the detergent. Particle formation w i l l occur spontaneously under these conditions. Typically, these l i p i d / D N A particles have a mean diameter of 110-120 nm, but particles as small as 50 nm can also be prepared by this method, depending on the l ipid composition and the detergent used. Aggregation can be a problem i f the lipids are not fully dispersed in the O G P initially, the detergent concentration is too low or i f the charge ratio of l i p i d : D N A is approximately 1. 1.1.4 Additional Formulation Strategies and Incorporation of Targeting Molecules In order to incorporate targeting molecules in the DNA/carrier complex, usually polylysine is used as the D N A binding agent. The targeting ligand is chemically coupled to polylysine via a disulfide linkage, followed by purification by column chromatography and dialysis (Buschle et al, 1995). Polylysine of 300 repeat units has been found the most useful for coupling. Residual uncoupled polylysine can reduce the efficiency of specific receptor-mediated transfection in the final preparation unless extensive purification is done (McKee et al, 1994). The ligand-polylysine complex may then be mixed with plasmid D N A (1:1 wt:wt), or with adenoviral particles then plasmid D N A . Simple mixing of the components results in rapid and spontaneous formation of the macro-complex (Cristiano et al, 1993). Carbohydrates have also been used as targeting ligands, particularly to hepatocytes via receptor-mediated uptake (Thurnher et al, 1994, Erbacher et al, 1996). A great deal of effort has been directed towards development of variously targeted liposomes and it can be suggested that many of the approaches involving surface modifications can be applied to lipid-based carriers. It is wel l established, however, that current approaches involving covalent attachment of proteins onto conventional liposomes 14 can result in liposome-liposome crosslinking (Erbacher et al, 1996). It is anticipated therefore that such procedures w i l l not be suitable for use with lipoplexes. Coupling techniques may be useful for attaching protein-based targeting ligands onto l i p i d / D N A particles, however, provided that the ligand can be incorporated stably in the outermost l ipid monolayer. Targeting strategies based on incorporation of low molecular weight ligands covalently attached to appropriate lipid anchors may be best for achieving cell targeting. A n example of this approach is that described by Lee and L o w (1995). involving folate-modified lipid. Folate-modified P E G has also been used in the preparation of polylysine/DNA/cationic l ip id /PEG complexes for transfection, which showed folate-receptor specific uptake at a high l ipid to D N A ratio (Lee and Huang, 1996). 1.2 Liposomes as Carriers of DNA and Drugs The remainder of this thesis deals exclusively with cationic liposome-based polynucleotide transfer vehicles. The following sections describe the rationale for their use, methods of preparation and physical properties. Lipid Bilayer Liposome Figure 1.4 Liposomes are vesicles composed of bilayer of amphipathic lipid surrounding an aqueous core. Compounds can be trapped within the liposomes, for example, certain chemotherapeutic drugs, such that the pharmacokinetic properties and biodistribution of drugs can be favorably altered. Drug delivery using liposomes, after more than two decades of basic research, is finally reaching 15 the stage of clinical trials and marketable products (reviewed in Chonn and Cullis , 1995). The alteration in drug disposition achieved by liposome encapsulation can have a favorable effect on the therapeutic index of the drug, ideally minimizing its toxic side effects and maximizing its delivery to sites of inflammation, certain organs, or to tumors. The liposomes can also protect the drug from rapid chemical or metabolic degradation after administration. This same principle was applied to D N A as a macromolecular drug that requires protection from degradation prior to its reaching the site of action. The liposome-based D N A carriers used in this thesis work do not involve entrapping the D N A within the aqueous core of the vesicles, but rather with forming a charged complex. However, the starting material (the cationic liposomes) is prepared in the same way as for conventional liposomal drugs. In addition, several chapters of this thesis deal with the ability of the cationic liposomes to retain entrapped aqueous contents within the aqueous core. The following section therefore w i l l discuss the fundamentals of liposome formation, preparation and lipid behavior under typical experimental conditions. 1.2.1 Liposome Preparation Liposomes form spontaneously when bilayer-forming lipids, such as many phospholipids, are hydrated from the dry state with an aqueous solution and agitated. Multilayered structures, known as multilamellar vesicles ( M L V s ) typically form under these conditions. Specific preparation techniques can be used to influence the size of the vesicles, the internal volume, permeability and lamellarity ( M L V s vs. unilamellar vesicles). These methods include but are not limited to: reverse phase evaporation (Szoka and Papahadjopoulos, 1978), sonication (Mayhew et al, 1994), French press (Barenholz et al, 1979) and extrusion (Szoka et al, 1980, Hope et al, 1985). Liposomes <60 nm in diameter are called small unilamellar vesicles (SUVs), which are usually produced by sonicating the suspension of M L V s . S U V s are relatively unstable and have low internal aqueous volumes (trapped volume), making them unsuitable for drug loading purposes. Liposomes with a mean diameter of 60 to 16 approximately 400 nm are termed large unilamellar vesicles ( L U V s ) . These liposomes are typically produced by extrusion through filters, or by techniques involving the use of solvents or detergents (Cullis, 1987). In this thesis only the extrusion method of preparing liposomes was used, because of its ease of use and ability to generate stable, homogeneous L U V s of defined size with reproducible trap volumes (Szoka et al, 1980, Hope et al, 1985, Mayer et al, 1986). This procedure is described in more detail in Chapter 2. 1.2.2 Chemistry and Physics of Lipids It is important to discuss how liposomes form so that their biophysical behavior may be understood. The basic unit is the l ipid bilayer, with its hydrophilic and hydrophobic regions (Figure 1.4). However, under certain conditions, some lipids w i l l adopt non-bilayer structures, such as hexagonal phase lipid. The stability of the bilayer and the ability to form the type of nonbilayer structures involved in membrane fusion are critical properties for cationic liposomes used for gene transfer, a topic addressed in Chapter 6. Certain conditions such as temperature, p H , or the presence of electrolytes or charged polymers can affect the organization of lipids (see Chapter 5) and thereby influence their function. 1.2.2.1 Lipid Polymorphism To minimize free energy, when phospholipids are in an aqueous medium the hydrophobic fatty acyl chains ("tails") of the molecules orient together due to the hydrophobic effect. The hydrophilic phosphate "headgroup" regions thus become oriented toward the aqueous medium. For a bilayer to form, such as we find in cellular membranes, the average molecular shape of the molecules must be cylindrical. The "shape concept" of lipid polymorphism is illustrated in Figure 1.5 (Cullis, 1986). If the molecular volume occupied by the headgroup and the acyl chains is similar, then a symmetrical bilayer can form (lamellar phase, L a ) . Other organizations of l ipid and water can also exist. For example, i f the headgroup volume is large compared to the acyl chains, and the l ipid concentration is high enough (above the critical micelle concentration, or C M C ) , then micelles may form. Cone-shaped lipids, with either part of the molecule larger than the other (headgroup vs. acyl chain region), may lead to 17 18 the formation of tubes of hexagonal phase lipid (Figure 1.6). H i phase lipid has the headgroups oriented toward the outside of the tubules, while in Hu phase the acyl chains are on the outside. It is important to note that hexagonal phase-preferring lipids can be stabilized into a bilayer configuration by the presence of bilayer-forming lipids or those of opposite molecular shape. DODAC or DOPC, for example, can stabilize DOPE into a bilayer configuration, permitting the formation of DOPE-containing liposomes. The phase adopted by the lipid depends on the intrinsic curvature of the lipid bilayer, temperature, hydration and influences from neighboring lipid molecules i f present. Lipids with a large (or charged) headgroup (such as many of the cationic lipids illustrated in Figure 1.2) exhibit a larger radius of curvature than lipids with a smaller headgroup due to packing constraints. Thus, the transition from lamellar to hexagonal phase occurs at a higher temperature (Tm) relative to lipids with small headgroups due to the larger energy requirement. For example, van Dijck et al. (1978) demonstrated that the enthalpy of the L a to Hu phase transition for DOPC (bulky choline headgroup) is approximately 11.2 kcal/mol while that of DOPE (smaller ethanolamine headgroup) is only about 4.5 kcal.mol. 1.2.2.2 Lipid Phase Behavior The organization of lipids in a bilayer state is also dependent on acyl chain fluidity and headgroup interactions (reviewed in Boggs, 1980). Below a critical temperature, the lipids form a gel state, characterized by a rigid, parallel organization of the lipid chains. Above the phase transition temperature (Tc), the acyl chains become more fluid, more permeable (Bittman and Blau, 1972) and less stiffly organized in the liquid crystalline state, denoted as Lc (Figure 1.7). Liquid Crystalline State Gel State Figure 1.7 Lipids undergo a change in ordering at a characteristic critical temperature. Lateral diffusion of the lipids is slower in the gel state than in the liquid crystalline state. The temperature range at which this transition occurs varies directly with acyl chain length (Papahadjopoulos et al, 1973) and inversely with the degree o f chain saturation (Chapman et al, 1975). Shorter or unsaturated acyl chains are more disordered in the bilayer, making it more fluid. Tc may also be affected by headgroup size and charge. Bilayers formed from lipids containing a charged or bulky headgroup, for example, w i l l be less ordered. L ip id packing constraints also have an influence on transition temperature; the size of the lipid molecules is directly related to the transition temperature. If the lipids are charged, electrostatic repulsion between lipids in the bilayer could also reduce the Tc (Cameron et al, 1981, van Dijck et al, 1978). Generally, the l ipid bilayer is less permeable to small molecules below the Tc. However, permeability of a liposomal bilayer composed of more than one type o f l ipid may be influenced by additional factors such as the packing order, fluidity and lipid-l ipid interactions. Table 2 lists typical lipids used in the preparation of liposomes and their transition temperatures. L ip id Tc (°C) Cationic Lipids D O D A C -8 D D A B 47 Phospholipids Distearoyl PC (18:0, 18:0) 55 Stearoyl, oleoyl PC (18:0, 18:1) 6 Stearoyl, linoleoyl PC (18:0, 18:2) -16 Stearoyl, arachidonoyl PC (18:0, 24:4) -13 Dioleoyl phosphatidylethanolamine (DOPE) (18:1, 18:1) 0 Dioleoyl phosphatidylcholine (DOPC) (18:1, 18:1) -22 Dipalmitoyl PE (16:0, 16:0) 63 Dipalmitoyl PC (16:0, 16:0) 41 Dipalmitoyl PS 55 Dipalmitoyl PG 41 Dipalmitoyl P A 67 Dimyristoyl PC (14:0, 14:0) 24 Dilauroyl PC (12:0, 12:0) -1 Table 2 Lipid transition temperatures (Tc) depend on acyl chain length, degree of chain saturation and headgroup size or charge. 1.2.2.3 Specific Lipids Used The lipid portion of lipoplexes typically is composed of a cationic l ipid and a helper lipid. There are many choices now available for cationic lipids, which may give slightly different results in transfection or in physical studies. The helper l ipid is often D O P E , which gives the lipoplex improved in vitro transfection. D O P E -containing complexes, however, are not stable. For in vivo use and improved stability, a recent development is the replacement of D O P E with cholesterol and sometimes poly(ethylene glycol) (PEG)-lipids. 1.2.2.3.1 Cationic Lipids Since the introduction of cationic lipid-based gene therapy by P. Feigner in 1987, the synthesis of new cationic lipids is an ongoing effort. The first studies used D O T A P (Figure 1.2) and syntheses of new cationic lipids were based initially on its structure. The cationic headgroup consists of one or more amine groups. The hydrophobic portion typically consists of two 18-carbon acyl chains. Variations have been made by modifying the linker group in between the headgroup and the acyl chains, altering the degree of chain saturation, or adding additional hydrocarbon moieties on the amine. A few such examples are illustrated in Figure 1.2. Structure-activity relationships are few at present (Feigner, P. et al., 1989, Bennet et al, 1998, B y k et al, 1998). Bennet et al. found that liposomal l ipid acyl chain length and gel to liquid crystalline phase transition temperature were directly related to the effectiveness of the cationic liposomes in delivering antisense oligonucleotides and to the requirement for helper l ipid (DOPE). B y k and coworkers synthesized a series of cationic lipids that varied in the linker region of the molecule (between the amine headgroup and the acyl chain tail region), which affected the chain shape (linear vs. T-shaped), and found the T-shaped cationic lipids to be more effective transfection reagents. Ansel l et al. (1997) prepared derivatives of D O D A C (dioleyldimethylammonium chloride) with various acyl chain lengths, amino groups and different halide counterions (Br", CI", I"). The parent compound D O D A C , which was used in this thesis, was shown the most effective in the series, with in vitro activity 4-5 fold greater than D O T M A . Reducing the degree of saturation or the length of the acyl chains reduced in vitro transfection activity. Molecules with C f as the counterion were the most 22 effective. Cationic lipid structure affects how the l ipid binds to D N A , resulting in subtle differences in lipoplex morphology (Sternberg et al, 1997). Lipoplex composition, and possibly molecular arrangement, also has a direct effect on how the lipoplex interacts with cellular membranes. Even so, improvements in activity over the older cationic lipids are generally within 1 to 2 orders of magnitude for specific cell lines or tissues. This should not be too surprising; their main purpose, after all, is simply to bind D N A through electrostatic interactions and possibly to enhance cellular uptake. The present study has used mainly D O D A C , as well as D D A B (dimethyldioctadecyl ammonium bromide) and D O T A P (dioleoyltrimethyl ammonium propane) to a lesser extent, as the cationic l ipid component of the lipoplexes. D D A B has saturated acyl chains and no linker group, while D O D A C has monounsaturated acyl chains. The headgroup for D O D A C and D D A B is the same, and both lipids can form liposomes without other l ipid components. The gel to liquid crystalline phase transition temperatures of D O D A C , at -8°C and D D A B at 47°C are quite different (Table 2), due to the difference in acyl chain saturation. These cationic lipids have been used extensively to transfect a variety o f cell types both in vitro and in vivo. 1.2.2.3.2 Phospholipids Phospholipids used in these studies include dioleoylphosphatidylethanolamine (DOPE) and dioleoylphosphatidylcholine (DOPC) (Figure 1.3). Both of these synthetic lipids have been used extensively in conventional liposomes. D O P E is a Hu phase-forming phospholipid and thus by itself, D O P E w i l l not form liposomes. However, it can be stabilized into a bilayer by other phospholipids or cationic lipids of complementary molecular shape (see section 1.2.2.1). It can form stable liposomes, for example, when combined with cationic l ipid at approximately 50 m o l % D O P E or less. The extrusion method of liposome preparation for DOPE-containing cationic vesicles can be performed at or below ambient temperature. This is possible because the gel to liquid crystalline phase transition temperature (Tc) of D O P E is quite low (-16°C). Attempting to hydrate or extrude D O P E -23 containing vesicles at elevated temperatures (>37°C) may produce poor results, in fact, due to the tendency for D O P E to form Hu phase lipid. The presence of salt-containing buffers (e.g. >10 m M NaCl) or strong acids w i l l also induce this change. Dioleoylphosphatidylcholine (DOPC) and egg phosphatidylcholine (EPC) are lamellar phase-preferring phospholipids. D O P C is a semi-synthetic product with a well-defined acyl chain composition. Natural source egg yolk P C contains mixed acyl esters with unsaturated cis-oriented double bonds. Phosphatidylcholine (PC) is the major component of most eukaryotic cellular membranes. Its cylindrical molecular shape allows it to form liposomes readily in an aqueous environment. D O P C has a Tc similar to D O P E due to the long unsaturated acyl chains (18 carbons, one double bond on each chain). The presence of salts or low p H w i l l not affect the ability of P C to form bilayers, and it has a strong stabilizing effect on DOPE-containing bilayers. The main chemical instability of phospholipids is hydrolysis, with formation of lysophospholipids, where one acyl chain of the phospholipid has been cleaved off. Oxidation can also occur i f the phospholipid is unsaturated. Both of these processes are facilitated by high temperatures or oxidative conditions. Stock lipids used in these studies were stored at - 7 0 ° C , and l ipid films were stored under N 2 gas at - 2 0 ° C to minimize the formation of degradation products. Liposome extrusion was performed using pressurized N 2 gas. Stocks were also monitored for changes in color (yellowing) or odor. Occasionally, thin-layer chromatography (TLC) was performed to check lipid purity i f there were any concerns about possible degradation. Liposomes were discarded after 3-4 weeks of storage at 4°C. 1.2.2.3.3 Cholesterol (Figure 1.8) is a steroidal component of animal cell membranes and now commonly used in liposomes. Its main function in liposomes is to modify the gel to liquid crystalline phase (Lc) transition (Ladbroke et al, 1968) by affecting the lipid acyl chain order. The permeability of the membrane is thereby influenced by the presence of cholesterol (Bittman and Blau, 1972, Demel and de Kruijff, 1976). B y decreasing the order of 24 the gel phase and increasing the order of the L c phase, cholesterol reduces the endothermic requirement of the transition. It achieves this effect by its insertion along the long axis in the hydrophobic region (aligned with the acyl chains) of the phospholipid bilayer. This effect increases as the cholesterol content increases (van Dijck et al. 1976). This means that a fluid bilayer can become more rigid and less permeable as cholesterol content is increased, which is an advantage when the liposomes are being used for drug delivery. In the in vivo setting, including approximately 30 mo l% cholesterol in the liposomal formulation grants another advantage, that of reducing rapid clearance from the bloodstream. The liposomal membrane itself may be stabilized by cholesterol, reducing contents leakage (Papahadjopoulos et al., 1973, Finkelstein and Weissman, 1979). Cholesterol prolongs liposome circulation time, due to a decrease in complement opsonization (Weinstein et al, 1981), less uptake by phagocytic cells of the reticuloenthothelial system (RES) (Patel et al, 1983) and less destabilization by lipoproteins (Scherphof, et al, 1978, Ki rby et al, 1980, Damen et al, 1982). These effects may be due to cholesterol's effect on l ipid fluid dynamics (Brulet and McConnel l , 1977), indicating the need for a fluid membrane to allow receptor-mediated uptake of opsonized particles by phagocytic cells such as macrophages. H Figure 1.8 Chemical structure of cholesterol, a commonly used component of liposomes. More recently, cholesterol has been shown to be an effective helper l ipid for cationic liposomes used for D N A delivery via lipoplexes (Templeton et al, 1998, Crook et al, 1998, Sternberg et al, 1998). B y an unknown mechanism, cholesterol either allows the lipoplexes to interact with cells and to be internalized, or plays a role in endocytic escape after internalization. Internalization might occur due to protein binding to the lipoplexes followed by receptor-mediated uptake (such as macrophage scavenger receptors or lipopotein receptors). One could 25 speculate that endocytic escape might occur as a consequence of cholesterol exchange into the inner endosomal membrane, destabilizing the lipoplex, resulting in cationic lipids interacting with endosomal membranes (perhaps in detergent-like fashion), ending in endosomal disruption and DNA escape into the cytoplasm. In addition, using cholesterol as a helper lipid rather than DOPE may impart a greater stability in the cationic liposomes and lipoplexes, which is addressed in Chapter 7. Sternberg and coworkers (1998) recently showed that DDAB/cholesterol (1:1)-lipoplexes are more stable than those made with DDAB/DOPE liposomes. Freeze-fracture electron microscopy showed roughly spherical particles, 200-400 nm, with occasional protrusions ("map-pin" structures), in contrast to DDAB/DOPE lipoplexes which are typically highly aggregated and heterogeneous. The significance of the protrusions is unknown. 1.2.2.3.4 PEG-Lipids Poly(ethylene oxide) commonly known as polyethylene glycol (PEG) is a polymer that can be covalently linked (coupled) to a phospholipid (Figure 1.9). distearoylphosphatidylethanolamine (DSPE)-PEG 2000 O Linker DSPE (C,8) po1y(ethylene oxide) Figure 1.9 Chemical structure of lipid-modified PEG, a common liposome component Lipid-modified PEG can be included in liposomes to provide a hydrophilic and steric barrier for the liposome surface (Woodle and Lasic, 1992, Needham et al, 1992). This allows an enhanced liposome circulation time in vivo due to a reduction in serum protein binding (Du et al, 1997) and reduced uptake by cells of the RES (Allen et al, 1991, Liu and Liu, 1996). This avoidance of detection within the body is what gave rise to the name "stealth liposomes." Vesicle-vesicle 26 aggregation and fusion is also inhibited in PEG-containing liposomes (Mori et al, 1998, Wasan et al, 1998 and see Chapter 7). The PEG concentration and the number of repeat units (the PEG MW) determines the size of the shield (Kenworthy et al, 1995). Lipid-modified PEG with a M W of approximately 2000 u is typically used for liposome applications, which results in a surface barrier that has been estimated to extend approximately 3-4 nm from the liposomal bilayer surface (Zeisig et al, 1996). While increasing PEG concentration increases the dimensions of the surface barrier, Edwards and coworkers (1997) have demonstrated by cryo-transmission electron microscopy (CTEM) that PEG-phospholipid concentrations above 5-10 mol% in 100 nm LUVs may result in separation of the PEG-lipid into micelles or mixed phospholipid-PEG micelles. Cholesterol is often included in PEG-containing liposomes as a stabilizing agent. It has been suggested that cholesterol prevents micelle formation by making the liposomal membrane more rigid and less able to adopt the high degree of lipid layer curvature found in micelles (Edwards et al, 1997). C T E M analysis showed that cholesterol-containing PEG-liposomes were more homogeneous in size distribution. Phase separation of PEG-lipid thus is influenced by the lipid composition of the liposome, and likely the temperature and method of liposome preparation (SUV vs. L U V or M L V ) , or other factors that affect lipid organization or stress the liposomal membrane. The choice of lipid anchor for the PEG also has a direct effect on its stability in the liposomal membrane and the subsequent pharmacokinetic properties of the PEG-liposomes (Webb et al., 1998). The use of PEG-lipids in polynucleotide transfer vehicles has only recently been explored, either for oligonucleotides (Kabanov et al, 1995, Meyer et al, 1998) or for plasmid D N A (Wasan et al, 1996, Hong et al, 1997, Sternberg et al, 1997). Chapter 4 describes an attempt to utilize PEG-lipid in DOPE-containing cationic liposome formulations to prevent 27 excessive lipoplex aggregation. Chapter 7 describes the beneficial use of PEG-lipid to prevent salt-induced aggregation of cholesterol-containing cationic liposomes and lipoplexes. 1.2.3 Entrapment of Drugs in Liposomes The ability of a liposome to retain a compound is governed by liposome properties, drug properties and external conditions. It is beyond the scope of this thesis to give an exhaustive review of this topic, however, trapping of amine drugs into lipoplexes is addressed in Ch. 7. Therefore, a few essential points should be made about how drug trapping inside lipid vesicles is achieved and optimized, with several examples of trapping methods. 1.2.3.1 Passive Drug Loading into Liposomes Drugs may be incorporated into liposomes passively or actively. Passive trapping means that the drug is simply present during liposome formation and incidentally becomes incorporated in either the aqueous phase (highly water-soluble compounds) or in the lipid phase (hydrophobic compounds). For highly water-soluble drugs exhibiting no liposomal membrane partitioning, the maximum concentration achievable inside the liposomes is the same as the drug concentration present at the time of liposome preparation. This process can be optimized for water-soluble drugs by preparing the liposomes in a way that maximizes the internal aqueous contents (trapped volume, Vt) of the liposomes and by minimizing drug leakage. Processes that produce large unilamellar vesicles (LUVs), such as extrusion or ethanol injection, generate liposomes with large Vt (Cullis et al., 1987). Once the external medium of the passively loaded liposomes is changed, such as by dilution or by administration to an animal, the release rate of the loaded compound is determined by the permeability of the drug through the membrane and the stability of the liposomal membrane. For highly water-soluble compounds, the permeability of a given membrane is directly related to the molecular weight of the entrapped compound (Papahadjopoulos et al, 1973), all else being held constant. Factors that influence membrane permeability in general, 28 such as osmotic stress, l ipid acyl chain length, temperature, the inclusion of cholesterol, etc., can affect the rate of efflux of a compound entrapped in liposomes. For hydrophobic compounds, incorporation may be mainly into the lipid bilayer rather than the aqueous core, and so the maximum possible concentration in the liposome may be much greater than that of the initial external concentration. Their incorporation and release is dependent not only on molecular weight and oil/water partition coefficient but also on their propensity to exchange out of the liposome into cellular membranes (Madden et al, 1990) or lipoproteins (K. Wasan et al, 1998). For drug delivery, more molecules per vesicle may be loaded in multilamellar vesicles ( M L V s ) due to the number o f l ipid layers present, however, smaller liposomes may be required to enhance circulation time in vivo. Many compounds, however, are amphipathic, including drug compounds which one might wish to encapsulate into liposomes for therapeutic benefit. Amphipathic compounds inside liposomes would reside not only in the aqueous core but also at the lipid-water interface. Their retention within the liposome depends on their rate of membrane transit. If however, the drug is ionizable (many drugs are weakly basic amines), an ion or p H gradient may be established across the liposomal membrane and thus drive encapsulation of the compound and inhibit egress from the vesicles, as discussed in the following section. 1.2.3.2 Active Drug Loading into Liposomes This method relies on an ion gradient, such as N a + / K + , or a difference in p H between the interior and exterior of the liposomal membrane. Conditions are set such that the drug to be trapped is in the ionized form at the interior p H . This is also referred to as a p H gradient method or "remote loading." In the remote loading method, L U V s are prepared in a high-capacity buffer solution at a p H below the p K a o f the amine drug. A n example of a commonly used buffer for the liposome interior is 300 m M sodium citrate, p H 4.0. Because citrate has three ionizable carboxylates, it has a high buffer capacity. The drug solution is added to the liposomes, then the exterior p H is 29 raised above the p K a by adding an appropriate high p H buffer (such as sodium phosphate). The liposomes may also be passed down a Sephadex column for buffer exchange. The unionized drug (at the higher pH) freely passes into the liposomes, but becomes ionized once inside (at the lower pH). Because of the unfavorable thermodynamics of egress for the charged entity, the ionized drug is trapped inside. This process is illustrated schematically in Figure 1.10. Chapter 7 describes the use of the p H gradient method for loading of transfection-enhancing amine drugs into cationic liposomes and lipoplexes. The energy barrier for the passage of an ion through a l ipid bilayer is highly unfavorable, even for small ions (Parsegian et al, 1969). Some methods of loading drugs into liposomes rely on an ion gradient generated by an ionophore in the liposomal membrane. A n ionophore is a lipophilic macromolecule that allows passage of ions (such as K + , C a 2 + or M n 2 + ) into the liposome interior. Due to the resulting charge imbalance, a membrane potential is set up. Proton flux may occur to balance the charge difference, thereby establishing a transmembrane p H gradient. Examples of K + ionophores used for liposomal drug loading are nigericin (Deamer et al, 1972) and valinomycin (Bally et al, 1986). The p H gradient that subsequently develops is dependent on the relative potassium gradients across the l ipid bilayer. When valinomycin is added to liposomes prepared in KC1 and then exchanged into N a C l for the external buffer, potassium and sodium ions w i l l both cross the membrane. Because of the ionophore, however, the rate of K + egress from the liposomes is much greater than that of N a + , resulting in a membrane potential (interior negative). C a 2 + ionophores used for drug loading include A23187 and X 5 3 7 A (Blau et al, 1984); they operate on the same principle - a potential difference or proton gradient is what drives drug accumulation. 30 OH & .9-53 § " " S I ' S 1 1 p i .2 § B K 5 1 O 2 M 11 I II! CO a s * s 8 1 a P . f ,8 CO I o o ca o B CO § O CO O •3 1 O a o 1 09 P i o & •pal 31 Another method of active drug loading is with the use of ammonium sulfate gradients (Figure 1.11). Liposomes are prepared in ammonium sulfate buffer, and drug is added to the liposome interior lipi^bilayer liposome exterior (ppt) B B Figure 1.11 Bidirectional (coupled) exchange loading can be accomplished via an ammonium sulfate gradient. Adapted from Ceh and Lasic, 1995. external bulk buffer. Uncharged drug crosses the liposomal bilayer, becomes protonated because ammonium sulfate dissociates into ammonium ion, sulfate ion and hydrogen ions. Then the protonated drug either associates with the anionic liposomal membrane, or becomes a sulfate salt and precipitates when the concentration of the salt form becomes elevated. The membrane-associated and precipitated forms are not free in solution, so they do not reduce the drug gradient driving accumulation within the liposome. The ammonia that donated the proton to the drug is no longer charged, so it can cross through the l ipid bilayer to the exterior, accept a proton from exterior drug and thus regenerate the cycle. In addition to ion-gradient driven loading of the drug into the aqueous compartment of the liposomes, certain drugs, such as doxorubicin, go to the lipid-water interface to form membrane-bound forms of the drug. This reduces the drug concentration in the aqueous phase and allows further drug accumulation by increasing the drug gradient. Intravesicular precipitates (sulfate salt o f the drug) can also form once the drug solubility in the interior aqueous 32 compartment is exceeded. For example, doxorubicin has been noted to precipitate inside liposomes. It can be loaded into liposomes at concentrations far exceeding that predicted by the simple Henderson-Hasselbach relationship: [ H + ] i n s i d e / [ H + ] 0 U t S i d e = [drug +]i nside/[drug +] 0utside- Both of these partitioning processes can greatly enhance the concentration achievable inside the liposomes (reviewed in Ceh and Lasic, 1995). A M n 2 + gradient loading method has recently been described which involves a M n 2 + ionophore incorporated into the liposomes, to exchange M n 2 + out of the liposomes and protons into the liposomes, creating a p H gradient to drive drug loading (Fenske et al, 1998). Interestingly, when doxorubicin is the drug being loaded, the formation of Mn-drug complexes can occur inside the liposomes (Cheung et al, 1998). Like the precipitated drug in the example above, drug efflux is limited and the soluble drug gradient remains high, promoting a higher internal drug concentration than would be achievable with the p H gradient alone. exterior buffer liposome interior A H H + A H (Reductant) PMS + 2e P M S H (Lipophilic proton carrier) D R U G 2Fe(CN) 6 2e PMS 2Fe(CN) 6 (Oxidant) 3-->DRUG (trapped) Figure 1.12 A redox system can be used to establish a pH gradient across liposomal membranes for drug loading. To make the pH gradient, liposomes are prepared with the oxidant passively trapped inside the liposomes, then buffer is added which contains the reducing agent. A lipophilic proton acceptor carries the H + across the liposomal membrane. After oxidation by the trapped couple, the protons are released inside the liposome interior, creating an acidic environment. The resulting pH gradient can drive the loading of ionizable drugs. Adapted from Deamer et a l . , 1972. 33 Another coupled method to achieve H + gradient is through a redox system, an example of which was described by Deamer et al. (1972). In their method, liposomes were prepared with the oxidant ferricyanide entrapped, and a lipophilic proton acceptor, phenazine methosulfate, inserted in the membrane. Buffer containing a reductant was added, such as ascorbic acid. Protons are released inside the liposomes after the trapped couple undergoes an oxidation step. This situation creates an acidic interior which can result in drug ionization and entrapment. (Figure 1.12). The most important factors in p H gradient loading are the ability o f the uncharged drug to permeate the lipid bilayer, the ability of the liposomal bilayer to maintain the p H or ion gradient and td retain the drug. These are all influenced by lipid composition. Buffer choice, drug physico-chemical properties and the size of the initial gradient (at least 3 p H units is preferred) w i l l affect the p H gradient (Madden et al, 1990, Harrigan et al, 1992). The partition coefficient and water solubility of the compound both have an influence on the rate of drug uptake and the A. Comparison of ion vs. pH gradient B. Uptake of various drugs into LUVs with 3.5 unit pH gradient Drug T r a p p i n g Efficiency (% ) Ion gradient pH gradient Do xorubic in 95-98 80-98 Vinblastine 90 --Dibucaine 60-98 60-95 Chlorpro mazine 80 -Dopamine 50-90 60 Seratonin 20-40 20-40 Ep inephrine 20-4 0 20-40 U p t a k e a t 2hrs Drug (n m o l / ) i m o l l i p i d) M ito xantrone 198 Ep irubic in 200 Daunorubicin 204 Do xorubic in 203 Vincristine 130 Vinblastine 127 Lidocaine 87 Chlorpro mazine 96 Dibucaine 176 Propranolol 187 Timo lol 97 Quinid ine 74 D o p a m in e 177 Serotonin 78 Imip ramine 188 Quinine 81 Chloroquine 88 Quinacrine 71 Codeine <1 Table 3 Many drugs have been successfully entrapped into liposomes by active loading methods. A : taken from Mayer et al, 1986. B: taken from Madden et al, 1990 and Cullis et al, 1997; L U V : large unilamellar vesicle (-100 nm mean diameter). Uptake studies were done with EPC liposomes at RT.uncharged state. 34 retention within the liposome. If the interior buffer capacity is too low and a large amount of drug is loaded into the liposome, the p H gradient could be dissipated. In that case, drug retention would be determined only by the drug's intrinsic permeability through the membrane. A n y external factors that could influence bilayer permeability, such as temperature, l ipid composition, biological interactions and lipid-mixing would also be important. Nevertheless, a growing number of drug compounds have successfully been encapsulated into liposomes using active loading procedures (Table 3) (Mayer et al, 1986, Madden et al, 1990). 1.3 L i p o s o m e - D N A Complexes 1.3.1 Preparation of Lipoplexes To make lipoplexes, cationic liposomes and D N A are added together by pipetting the two together at the desired ratio. Lipoplex formation occurs rapidly by a self-assembling process, driven by polyelectrolyte interactions and condensation phenomena. Self-assembly refers to the formation of complex macromolecular structures from simpler elements without a requirement for any additional energy input to drive the reaction. This occurs through multivalent charge-charge interactions and hydrophobic interactions. Condensation, or compaction, of the D N A may occur as water molecules are excluded from the structure as it assembles into a complex with the lipids. Lipoplex structure is discussed further in Chapters 5 and 6. If D O P E is present in the formulation, it is important to avoid high salt concentrations or high temperatures, as indicated in section 1.2.2.3.2 above. A n y conditions that promote enucleation can result in precipitate formation, particularly when the charge ratio is approximately unity. Order of addition, charge ratio, mixing rate, l ipid composition, D N A purity, etc., have all been studied as possible influences on lipoplex structure and excessive lipoplex aggregation (Sternberg et al, 1998, Lasic et al, 1998). Many of these issues are addressed in detail in Chapter 2 and throughout this thesis. 35 1.3.2 Physical Properties of Lipoplexes The physico-chemical behavior of lipoplexes and their components has a direct effect on their biological properties, a concept that forms the basis of this thesis. For example, the extent and timing of lipoplex aggregation influences the size of the particles used for transfection. Larger particles may settle onto the cell surface in vitro quite well , but size may limit cellular uptake or totally prevent in vivo delivery to the target population. The optimal structure of lipoplexes is unknown, but surely involves extensive liposome rearrangement. This rearrangement is influenced by l ipid composition and external conditions and due to vesicle-vesicle lipid-mixing events driven by their interaction with D N A , which is addressed in Chapter 6. The lipid-mixing ability of the lipoplexes should have a direct effect on their ability to interact with cellular membranes (introduced in Chapter 2), but may be influenced by the many steps involved in the process of in vitro transfection. 1.4 Hypotheses of the Research and Specific Aims This thesis focuses on the relationship between lipoplex physical properties and biological function. The overall goal of the research project was to use the information gained in the exploration of that relationship to redesign the lipoplexes for optimal activity. HYPOTHESIS 1: L ip id composition, salt concentration and charge ratio directly influence the transfection ability of lipoplexes. Aim 1: To determine the effect of lipid composition, salt concentration and charge ratio on lipoplex particle size. Aim 2 To determine the effect of lipid composition, salt concentration and anion polyvalency on liposome rearrangement and lipoplex structure. Aim 3: To determine the effect of l ipid composition, salt concentration, charge ratio on the lipid-mixing behavior of cationic liposomes and lipoplexes. HYPOTHESIS 2: Aggregation and lipid-mixing of the cationic liposomes following addition of D N A , salt or serum result in the formation of structures that are less capable of further lipid-mixing reactions required for transfection. 36 Aim 1: To relate lipid composition, salt concentration and lipoplex particle size to level of in vitro transfection. Aim 2: To relate lipid-mixing ability o f cationic liposomes and lipoplexes to level of in vitro transfection. H Y P O T H E S I S 3: Retention of basic liposome properties, such as aqueous trapped volume, w i l l facilitate development of liposomes that can deliver D N A and transfection-enhancing drugs. Aim 1: To develop lipoplex formulations that retain a particle size of <300 nm in 150 m M buffered N a C l , protect D N A from DNase I degradation and allow in vitro transfection. Aim 2: To develop lipoplex formulations that achieve p H gradient loading of an ionizable drug. In 1995 when this thesis work was begun, an obvious problem in the field of liposome-based D N A delivery systems was that of salt-induced aggregation of the vesicles. Elimination of this problem would presumably lead to D N A delivery systems that were more stable in a physiological environment and in NaCl-containing solutions typically used for in vivo administration. The early work presented in this thesis focused on using standard techniques for size reduction, in particular, sonication and the incorporation of the liposome aggregation inhibitor P E G - P E . A s demonstrated in Chapter 3, sonication successfully reduced the size of some lipoplex formulations but not others. Interestingly, the lipids protected the plasmid D N A from sonication damage, and transfection ability was unaffected by this process. This implies strong binding between the liposomes and the D N A . To further understand what was happening to the liposomes during the process of complexation to plasmid D N A , the next investigation dealt with the retention of an "aqueous trapped volume" within the liposomes (Chapter 4). Addition of D N A to the liposome was thought to induce major structural changes in the liposomes. It was anticipated that loss of liposomal aqueous contents would occur upon D N A complexation. Surprisingly, a significant retention of contents was noted. 37 It became clear that morphological studies would be helpful to visualize these changes in liposome structure. Cryo-transmission electron microscopy studies were done to determine the effects of addition of plasmid, oligodeoxynucleotides or phosphate anions on the liposomes. The photomicrographs in Chapter 5 clearly show that bilayer destabilization and fusion was occurring. The extent of liposome structural rearrangement was dependent on anion charge density, ionic environment and lipid composition. These parameters were quantified by the use of a R E T lipid-mixing assay, discussed in Chapter 6. In a continuing effort to pair biophysical attributes of the lipoplexes with transfection activity, a multi-step lipid-mixing assay was developed to model the process of in vitro transfection. This was used to determine how continual changes in liposomal membranes may affect their ability to transfect cells. The last chapter describes research identifying formulation approaches using cationic liposomes that could bind and protect plasmid D N A while inducing minimal changes in liposome size and retention of an aqueous trapped volume. It was important that these formulations could be prepared in NaCl-containing solutions. It was demonstrated that incorporation of cholesterol and PEG-modified lipids instead of the highly fusogenic lipid D O P E resulted in stable formulations. These salt-stable liposomes imparted protection of the plasmid from DNase as well as maintenance of transfection activity. The liposomes retained a significant trapped volume following D N A addition. A stable transmembrane p H gradient was established in the salt-stable lipoplexes. Preliminary data are provided which suggest that this gradient can be used to actively trap agents with protonizable amines, such as vincristine. It is believed that liposomes based on these or similar lipid compositions have the potential to be developed as codelivery systems for D N A and transfection-enhancing drugs. The introduction of this concept is a significant contribution to gene therapy research. 38 Chapter 2 Materials and Methods 2.1 Introduction A significant portion of this thesis was devoted to devising appropriate methods to analyze the biophysical behavior of lipoplexes. This chapter describes all methods used in the thesis, however, a more detailed discussion with comparison of additional related methodologies in this field may be found in the Appendix. 2.2 Materials 2.2.7 Materials for Plasmid Preparation Reporter gene plasmids encoding (3-galactosidase ( p C M V P, 7.6 kb), chloramphenicol acetyltransferase (pInexCATv2.0, 4.5 kb) and firefly luciferase (pInexL018, 5.65 kb) were constructed and provided by Inex Pharmaceuticals (Vancouver, B C , Canada). The luciferase plasmid was based on the p C M V p plasmid from Clontech (Palo Al to , C A ) and thus contains the same C M V promoter/enhancer element and SV40 polyadenylation signal. Luria Broth (LB) was obtained from Bio 101 (LaJolla, C A ) . Plasmid purification by anion exchange chromatography was performed using the Qiagen kit (Qiagen, Chatsworth, C A ) . D N A concentration was determined spectrophotometrically with a Model D U 64 (Beckman Instruments, Fullerton, C A ) . Dialysis of plasmid preparations was done using Spectropor cellulose dialysis membranes with a M W cutoff of 12,000 (Spectrum Medical Industries, Houston, T X ) . For preparation of radiolabeled plasmid, tritiated thymidine was purchased from DuPont /NEN (Markham, Ontario, Canada), which had a specific activity of 6.7 Ci /mmol and a purity of 97.6%. 39 2.2.2 Lipids 2.2.2.1 Cationic Lipids D O T A P (dioleoyl-l,2-diacyl-3-trimemylainmonmm propane) was obtained from Avanti Polar Lipids (Alabaster, A L ) . D D A B (dimethyldioctadecyl ammonium bromide) was purchased from Sigma. D O D A C (dioleyldimethyllammonium chloride) was generously provided by Inex Pharmaceuticals (custom synthesis, Dr. Stephen Ansell , Vancouver, B C , Canada). 2.2.2.2 Radiolabeled, Fluorescent and Anionic Lipids Radiolabeled cholesteryl hexadecyl ether (CHE) ( 3 H - or 1 4C-labeled) was obtained from DuPont /NEN. 1 4 C - D O P E was purchased from Amersham (Oakville, Ontario, Canada). 3 H - D O D A C and 1 4 C - D O D A C (used only in Chapter 3 experiments) was provided by custom synthesis (Inex Pharmaceuticals). The fluorescently labeled lipids N-(7-nitrobenz-2-oxa-l,3-diazol-4-yl)-modified phosphatidylethanolamine (NBD-PE) and Lissamine-rhodamine-modified phosphatidylethanolamine (rhodamine-PE) (Figure 2.1) were obtained from Molecular Probes (Eugene, OR) . 8 A r ^ CH3(CH2) ) 4-C-OCH2 | r O Pvhodamine B-l,2-dihexadecanoyl-5«-glycero- 3-phosphoethanolamine, Ixiethylarnmonium salt <CH3CH2)3NH O N-(7-nitrobenz-2-oxa-l,3-diazol- 4-yl)-l,2-dihexadecanoyl-5,n-glycero-3-phospho-ethanolamine, triethylarnmonium salt Figure 2.1 Chemical structures of rhodamine-PE (top) and N B D - P E (bottom). 40 Dioleoylphosphatidylcholine (DOPC) , dioleoylphosphatidylserine (DOPS), dimyristoylphosphatidylethanolamine-poly(eythyl^ glycol)2ooo ( D M P E - P E G ) and distearoylphosphatidylethanolamine-poly(eythylene glycol)2ooo (DSPE-PEG) were purchased from Avanti Polar Lipids or from Northern Lipids (Vancouver, B C , Canada). 2.2.3 Materials for Liposome Preparation The extrusion device used to prepare the liposomes (The Extruder) was provided by Lipex Biomembranes (Vancouver, B C , Canada). Chloroform and methanol were H P L C grade (Fisher or E M Science). The polycarbonate membranes used with the Extruder to limit the size of the liposomes were obtained from Poretics (Mississauga, Ontario, Canada). Assessment of the mean diameter of the liposomes was performed under ambient conditions on a Nicomp submicron particle sizer (Model 270, Pacific Scientific, Santa Barbara, C A ) with the laser operating at 200 k H z and a wavelength of 632.8 nm, and the scattering angle fixed at 90°. Liquid scintillation counting was done by mixing the sample with Pico-Fluor (Packard Instrument Co., Meriden, CT) and analyzing in a Canberra-Packard 1900TR TriCarb counter. Sphingomyelin/cholesterol (70/30 mol%) liposomes (SPH/Chol) were prepared by the Drug Development Unit, Advanced Therapeutics, B C Cancer Agency, under G M P conditions. SPH/Chol liposomes were 100 nm L U V s prepared in 300 m M sodium citrate (pH 4). 2.2.4 Materials for Analysis of DNA Protection and Transfection For D N A extractions, phenol (Gibco-BRL) was first equilibrated with T r i s -HCl (pH 8). Electrophoresis was done in 0.8% agarose gel using a BioRad Model 3000xi programmable power supply (BioRad, Richmond, C A ) , operating at 80-100 V . Gels were visualized after ethidium bromide staining with a U V transilluminator (Ultra Violet Products, San Gabriel, C A ) . Dissolution of gel slices was done with Solvable™ (DuPont/NEN, Boston, M A ) . Analysis o f reporter gene expression using the P-galactosidase assay was done with chlorophenol red galactopyranoside (Boerhinger-41 Mannheim, Germany) as the substrate, and the resultant product was quantified by measuring optical density in a plate reader from Flow Laboratories (TiterTek Multiscan 3IOC, Mississauga, ON). The C A T reporter gene assay required an extraction step in mixed xylenes (Aldrich Chemical, Milwaukee, WI) and the C A T substrate was 1 4 C-labeled chloramphenicol (D-threo-[dichloroacetyl-l,2-14C]-CAT, assay grade, DuPont N E N ) . Analysis of transfection using the luciferase reporter gene was done using a luciferase kit from Promega and a Tropix luminometer (Bio/Can Scientific, Mississauga, Ontario). Optical density for protein assays was measured in the plate reader mentioned above or in a M R X Microplate Reader (Dynex Technologies, Chantilly, V A ) . 2.2.5 Materials and Cell Lines for in Vitro Transfection Chinese hamster ovary cells (CHO) used in Chapter 3 were obtained from American Type Culture Collection ( A T C C ) , Rockville, M D ) and grown in Alpha Min imal Essential Medium supplemented with 10% fetal bovine serum (FBS). B16 cells used in Chapters 6 and 7 were obtained from N C I Tumor Repository (Bethesda, M D ) and maintained in R P M I media with 10% F B S . Tissue culture media were obtained from Stem Cel l Techologies (Vancouver, B C , Canada). Fetal bovine serum (FBS) was from G i b c o - B R L or I C N Biomedical (Aurora, Ohio). Tissue culture plates and plasticware were from Falcon (Becton-Dickinson, Franklin Lakes, NJ) . Lipoplexes were administered to the wells of cultured cells using a positive displacement pipettor fitted with a glass capillary tube (Dade International, M i a m i , F L ) . 2.2.6 Other Materials and Equipment Oligonucleotides (18mer; sequence: T C T C C C A G C A T G T G C C A T , MW=5683) used in Chapter 5 were provided by Genta (San Diego, C A ) . The probe-type sonicator used for experiments in Chapter 3 was a Sonifer Model 350 (Branson Sonic Power). Trapped volume measurements in Chapters 4 and 7 by the filtration method were done using MicroCon30 microconcentrators (Amicon, Beverly, M A ) with a M W 42 cutoff ( M W C O ) of 30,000 daltons. Cellulose dialysis tubing was from Spectrum Medical Industries (Houston, T X ) or from Fisher ( M W C O = 12,000-14,000 daltons). Fluorometric analyses in Chapters 4 and 6 were done with a Perkin Elmer L S 5 0 B luminescence spectrophotometer (Buckinghamshire, England). Triton X-100 was purchased from BioRad (Richmond, C A ) . The C T E M studies of Chapter 5 utilized a Zeiss E M 9 0 2 electron microscope and a custom-made high humidity sample preparation chamber. 1 4C-methylamine used for ion gradient measurements in Chapter 7 was bought from Amersham (Buckinghamshire, England). Vincristine H C I for liposome loading in Chapter 7 was obtained from B C Cancer Agency (Vancouver, B C , Canada). A l l other chemicals or reagents not specified otherwise were from Sigma Chemical Co. (St. Louis, M O ) . 2.3 Preparation of DNA and Liposomes 2.3.1 Plasmids D N A was obtained by transforming competent Escherichia coli (E. coli) bacteria with plasmid D N A . Plasmids contained a gene for ampicillin resistance to ensure that the bacteria growing in the presence of the drug retained the plasmid of interest. Following growth of the bacterial culture and subsequent amplification of the plasmid, the bacteria were harvested by alkaline lysis and the plasmid recovered by anion exchange chromatography by standard techniques (Sambrook et al, 1989) as described below. 2.3.1.1. Plasmid Isolation and Purification Transformed E. coli containing the plasmid of interest were grown from a glycerol stock (0.85 ml bacterial culture + 0.15 ml glycerol, stored at -70°C) and plated on a semisolid matrix containing Luria Broth (LB) , agar and ampicillin (50ug/ml). The bacteria were grown overnight at 37°C. A single bacterial colony was picked from the plate, introduced into 10 ml L B with antibiotic and incubated overnight in a shaker incubator at 37°C. It is important to ensure that the plasmid of interest is present before 43 continuing. This was done by checking for the presence of the desired plasmid using a standard mini-prep method (kit from Qiagen, Chatsworth, C A ) followed by restriction enzyme digestion and analysis by gel electrophoresis. From the 10 ml overnight culture, 5 m l was added to each of two 4 L flasks containing 1 L of L B with selection antibiotic. The bacteria were grown to saturation, usually overnight, in a shaker incubator at 37°C. The optical density at 550 nm (OD55o) o f the culture was about 1.0-1, indicating sufficient bacterial growth. Plasmid was routinely recovered from bacteria by an alkaline lysis procedure, which lyses the bacterial cell while maintaining bacterial D N A attachment to the cell wal l . This procedure enables subsequent precipitation of bacterial D N A and cellular debris, leaving a crude preparation enriched in plasmid. A plasmid D N A purification kit provided by Qiagen was used that utilizes the alkaline lysis method for harvesting, and anion exchange column chromatography for rapid purification. Detailed instructions were provided in the kit by the manufacturer. Using a spectrophotometer, an optical density profile was performed (between 230 and 340 nm) on a diluted sample of the purified D N A . The profile generated a peak at OD26o where D N A absorbs maximally. To determine the concentration and purity of the D N A sample, O D readings at 260 nm and 280 nm were taken and the concentration determined (one OD26o = 50 ug D N A / m l ) . The ratio of absorbance at 260 nm to that at 280 nm was used as an indicator of D N A purity (Sambrook et al, 1989). Typically, the ratio was 1.6 to 1.8, in the acceptable range. Following quantification, the sample was dialyzed to remove residual salts, which can affect the behavior of the D N A in its interaction with the carrier and its transfection efficiency. The entire D N A sample was dialyzed against 4 L of sterile distilled water overnight at 4°C. One hour after the start of dialysis 1 L of the water was replaced with fresh sterile water. After the overnight dialysis, the dialysis bag was transferred to a fresh beaker of sterile water and dialyzed 44 another 1 h at 4°C. The OD260 of the post-dialysis sample at an appropriate dilution in water was checked and concentration and recovery were calculated. The D N A solution was diluted to 1 mg/ml aliquots, which were stored at - 2 0 ° C until use. 2.3.1.2 Preparation of Radiolabeled Plasmid The preparation of tritiated plasmid, described below, is modified from standard protocols due to the need to induce the bacteria to incorporate large quantities of radiolabeled nucleotide (for example, tritiated thymidine) during log phase growth. E. coli containing the plasmid of interest were taken from a glycerol stock and plated onto an M 9 (minimal media) plate (M9: 10 g M 9 salts in 1 L autoclaved, distilled water, or 15 g M 9 salts per L of bacto agar to prepare plates) with appropriate antibiotics for selection. To each 100 m l of media, 5 m l of 20% glucose, 0.1 ml of 1 M M g S 0 4 , 10 ul of 1 M C a C l 2 , and 1.0 m l of 0.1% thiamine H C I were added, plus ampicillin 50 ug/ml). Incubation proceeded at 37°C, for no longer than 16 h to prevent antibiotic depletion and bacteria without plasmid from growing. After plating overnight, a lawn of white opaque colonies resulted. Fresh M 9 media (10-20 ml) was added and the colonies dislodged with a bent glass stirring rod. The liquid containing the bacteria was transferred to 100 m l M 9 and the O D 5 5 o was measured (approximately 0.1 OD) to ensure the bacteria were in log phase growth. The 100 m l M 9 culture was allowed to grow in a 37°C shaker until the O D 5 5 o was about 0.2-0.3, which required about 5 h. Next, 1 m C i tritiated thymidine, or sufficient quantity to reach the desired final specific activity was added to the culture. Incubation proceeded overnight at 37°C and a diluted sample was checked the next day prior to harvesting the bacteria, ensuring that the bacteria had grown to saturation ( O D 5 5 0 «1.5). For plasmid used in experiments in this thesis, the Qiagen kit was used (see section 2.2.1.1), to harvest and purify the plasmid, following the manufacturer's instructions. For transfection-grade radiolabeled plasmid, the final step was dialysis against several changes of water at 4°C overnight. The OD230-340 profile was generated as described above and the final 45 concentration and specific activity were determined following liquid scintillation counting of a known volume. From 50 ul of starting glycerol stock of E. coli carrying pCMVp-galactosidase, for example, typical yield from this procedure was 200 (j.g plasmid, with a specific activity of 80,000 dpm/ug. The 3 H-plasmid was stored at -70°C to minimize radiation-induced breaks. It was used for up to a year, and the quality of the plasmid was checked regularly (e.g. approximately every 2 months as needed) by agarose gel electrophoresis. 2.3.2 Cationic Liposomes High-purity lipids in the desired proportions were dissolved in HPLC-grade chloroform (5-20 mg/ml) and usually radiolabeled with trace amounts (~1 uCi/ml) of 3 H - or 1 4C-cholesteryl hexadecyl ether (CHE) . It is well-established that C H E is a suitable lipid marker for following the fate of conventional liposomes in vitro because it is non-exchangeable and non-metabolizable (Scherphof et al, 1987). The l ipid solution was reduced in volume under a stream of N 2 gas, and dried to a thin film for several hours under vacuum. The lipid films were hydrated by adding an aqueous solution while agitating with a vortex mixer, producing multilamellar vesicles ( M L V s ) . The hydrating solution was 300 m M lactose unless otherwise indicated. In Chapter 7, HEPES-buffered saline (HBS, 150 m M N a C l , 20 m M H E P E S , p H 7), CHES-buffered saline (CBS, 150 m M N a C l , 20 m M C H E S , p H 9) or 300 m M sodium citrate (pH 4) was used for cholesterol-containing liposomes. A l l hydrating solutions were 0.22 um-filtered. If D O P E was present hydration was done on ice. Liposomes were subjected to five cycles of freeze-thaw. The hydrated l ipid suspension was then passed 10 times at room temperature through an extrusion device containing three polycarbonate membranes (0.08 and/or 0.1 pm pore diameter as needed to achieve desired liposome mean diameter). The resulting large unilamellar vesicles (LUVs) had a mean diameter of 100-140 nm as determined by quasi-elastic light scattering (QELS) . Liposomes were stored at 4°C and used within 1 month. 46 Final l ipid concentration was usually determined by use of the radiolabeled C H E . For the liposomes used in the C T E M studies described in Chapter 5, l ipid concentration was determined by an assay of phospholipid phosphorous with the Bartlett assay (Bartlett and Lewis, 1970). Briefly, liposomes were digested with perchloric acid and oxidized, yielding inorganic phosphate (PO4 "). The phosphate was reacted with ammonium molybdate in the presence o f l-amino-2-napthol-4-sulfonic acid and sodium bisulfite, yielding a blue solution. The blue color from the reduced phosphomolybdate complex is proportional to the amount of phosphate present and was read spectrophotometrically at 830 nm. The amount of phosphorus present was determined by comparison to a standard curve. The limit o f detection for this assay was 0.2 pg phosphorus, and it was linear (r > 0.998) in the range of 1.2 to 6.0 ug phosphorus. 2.3.2.1 Sizing of Liposomes and Lipoplexes Q E L S analysis (Pecora, 1972, Holt et al, 1975) was performed using the Nicomp Submicron Particle Sizer. Quasi-elastic light scattering, also known as dynamic light scattering, derives a translational diffusion coefficient for the suspended particles which is translated into a mean diameter. A He/Ne laser shines through the sample and the incident light is scattered by the particles and measured by a detector. The submicron particles exhibit Brownian motion, which is movement due to the bombardment of the suspended particles by solvent molecules. The detector measures the intensity of the scattered light and transmits a photopulse signal to a computer utilizing a mathematical algorithm called an autocorrelator function. Larger particles move more slowly than smaller particles, generating less scattering of the light over time than smaller particles. The photopulse signal vs. time is used to determine the diffusion coefficient o f the particles, which is related to mean diameter. The algorithm takes into account the temperature, solution viscosity and refractive index of the sample, and assumes a spherical particle shape. The dedicated Nicomp software reports the mean particle diameter, standard deviation (SD) and a goodness-of-fit parameter (x2) 47 for a normal (Gaussian) distribution. The SD thus gives an indication of the degree of heterogeneity within a sample. In the results where "average S D " is reported (Chapter 3), this is the mean of the SDs of each sample in that group. This is more representative of the heterogeneity within samples, an important issue in this study, than would be the SD of the means. 2.4 Formation of Lipoplexes Plasmid to be used for making lipoplexes was prepared in water. Liposomes and plasmid D N A were centrifuged separately (7,000-10,000 x g for 2 min) prior to use to pellet any debris or aggregates. Large unilamellar liposomes and plasmid D N A were not pelleted by this procedure (Bally et al, 1997). Lipoplexes were prepared with plasmid D N A and liposome components cooled on ice i f the liposomes contained D O P E , or at room temperature otherwise. Liposomes and D N A were diluted separately in dust-free tubes using the same type of solution that was used to prepare the liposomes. Typically, liposomes were used at a final dilution of 0.5-1 m M total lipid, and D N A concentrations of 5-500 ug/ml in a total volume of 0.2-10 ml . Diluted plasmid D N A was pipetted gently into an equal volume of diluted liposomes. Flocculent aggregation of DOPE-containing liposomes was less likely i f D N A was added slowly to the liposomes and mixed immediately by gentle pipetting rather than by rapid vortex mixing. When D O P E -containing lipoplexes are described in this thesis as being prepared in salt-containing buffers, the liposomes were first prepared in 300 m M lactose, then D N A in the salt-containing buffer was added. Lipoplexes were allowed to form for about 30 minutes unless otherwise indicated. Lipoplexes were used within a few hours of preparation because transfection efficiency is reduced i f stored (Holland et al, 1996) and further aggregation can occur over time. 48 2.5 Characterization of Liposome Carriers of DNA Parameters of the l ipid carrier explored in this thesis which are believed to be important for lipoplex-mediated transfection include particle size and morphology, retention of liposomal trapped volume and lipid-mixing behavior (with self and target membranes). 2.5.1 Measuring Particle Size and Aggregation State Particle size of l ipid carriers can be measured by a number of techniques, such Q E L S already mentioned above and electron microscopy (EM) (Sternberg et al, 1995, Gustafsson et al, 1995, H u i et al, 1996). In this study, the Q E L S method with a Nicomp Submicron Particle Sizer was used, for which accurate measures of vesicle or solid particle diameters can be made in the range of 20-1000 nm. A s aggregation progresses, the analysis shows an increase in polydispersity. Diameters >1 um are not accurately measured with this type of particle sizer, which impedes somewhat the study of highly aggregated lipoplexes. Particle size was estimated by electron microscopy in Chapter 5 only. Aggregation state was also assessed visually, because cloudiness increases slightly as particle size increases, until flocculent white flaky masses, chunks or threadlike particulates appear in the suspension. Turbidity was measured by reading absorbance at 450 nm in a 96-well tissue culture plate in a plate reader (Chapter 5). 2.5.1.1 Sedimentation Assay A simple method to assess the degree of excess aggregation is to subject the sample to centrifugation (10,000 x g) (Chapter 4). Small lipoplexes, liposomes or plasmid D N A alone w i l l not pellet under these conditions, but large aggregates (>1 um) w i l l pellet (Bally et al, 1997). The centrifugation conditions required varied according to the cationic l ipid used. For example, DDAB-containing l iposome/DNA aggregated lipoplexes pelleted within 5 min at room temperature but DODAC-containing aggregates required 30 min of centrifugation at 4°C. The liposomes and/or D N A were radiolabeled and the supernatant was sampled for scintillation counting. The percentage remaining unpelleted was then calculated. 49 Lipoplexes with a D N A concentration of 25 pg/ml and liposome-only controls were prepared in a total volume o f 300 ul . One volume o f D N A was added to one volume o f liposomes to achieve the desired final l i p i d : D N A ratio. Samples were prepared on ice. A sample of 100 ul was taken for scintillation counting. The remaining volume was then diluted with 300 ul of either 300 m M lactose or H B S and incubated for 20 min at room temperature. In the case of H B S addition, the final concentration of N a C l was 90 m M . After centrifugation at 10,000 xg (4°C, 30 min), 100 ul of the supernatant were taken for scintillation counting. The percent lipid remaining in the supernatant was calculated by dividing the post-centrifugation H-l ip id counts by the pre-centrifugation counts. 2.5.1.2 Salt-Induced Aggregation D O D A C / D O P E or D O D A C / D O P C liposomes were prepared in lactose as described above. In the experiments described in Chapter 5 (Table 7), liposomes were diluted into N a C l solution such that the l ipid concentration was 2 m M . The concentration of N a C l was varied between 0 and 150 m M . Liposome mean diameter was assessed by Q E L S within 10 min of dilution. Lipoplexes were prepared with D O D A C / D O P E or D O D A C / D O P C liposomes and pInexCATv2.0 plasmid using 2 m M lipid and 200 ug D N A / m l ( l ip id :DNA = 1 0 nmoles lipid/ \xg D N A ) . Liposome/DNA complex formation was allowed to proceed for 30 min at R T , at which time 20 ul of lipoplexes were diluted into 250 ul of N a C l solution, followed by Q E L S analysis within 10 min of dilution. 2.5.2. Measurement of Trapped Volume The internal aqueous volume of the liposomes and lipoplexes was assessed by measuring the concentration of a radiolabeled water-soluble compound, 1 4C-lactose entrapped in the liposomes. The 1 4C-lactose was first loaded into the liposomes passively by adding it to the liposome hydration buffer prior to extrusion. Release was measured after addition of D N A by filtration and dialysis. The total retentate was determined by subtracting the released amount from the total 1 4C-lactose known to be in the sample. 50 2.5.2.1 Measurement of Trapped Volume by FUtration In triplicate, 400 ul of lipoplexes or liposome-only controls were prepared as described above. One hundred ul were sampled for dual-label ( 3 H-l ip id and I 4C-lactose) liquid scintillation counting. Samples (250 pi) were placed in the top of Microcon30 tubes. The samples were centrifuged at 7800 x g for 5-10 min. A sample of the filtrate (100 pi) was taken for scintillation counting. Neither D N A nor liposomes pass through the filter. For calculation of trapped volume, entrapped 1 4C-lactose dpms were calculated by subtracting the ultrafiltrate 1 4 C dpms from the 1 4 C dpms in the initial unfiltered complex or liposome control sample. L ip id concentration was determined from the specific activity of the 3H-liposomes used and from the known dilution in the sample preparation. Trapped volume was expressed as p i trapped 1 4C-lactose/umole lipid. Nonspecific association of 1 4C-lactose was variable but minimal (0-3% of scintillation counts) and contributed less than 2% of the total reported residual trapped volume. 2.5.2.2 Measurement of Trapped Volume by Dialysis Lipoplexes and liposome-only controls, prepared in triplicate, were sampled (100 pi) for scintillation counting. Samples were dialyzed against 300 m M sterile lactose overnight at 4°C. After dialysis, 100 p i of sample were counted. L ip id and D N A concentrations were unchanged following dialysis. For calculation of trapped volume, entrapped 1 4C-lactose was measured directly by sampling after dialysis. The entrapped 1 4C-lactose dpms were then converted to p i based on the dpm/pl of the 1 4C-lactose solution used to hydrate the liposomes. L ip id concentration was calculated based on the tritium dpms in the sample after dialysis and the known specific activity of the original liposomes. Trapped volume was expressed as pi trapped 1 4C-lactose/pmole lipid. As with the filtration method, nonspecific association of 1 4 C lactose was variable but minimal (0-3% of scintillation counts) and contributed less than 2% of the total reported residual trapped volume. 51 2.5.2.3 Determination of the Effect of Incubation with Tissue Culture Medium on Trapped Volume Lipoplexes and liposome-only controls were diluted 1:1 (v/v) into Eagle Minimal Essential Medium with or without 10% fetal bovine serum. After incubation for 4 h at either 23°C or 37°C, the trapped volume of the liposomes was assessed by filtration as described above. 2.5.3 Cryo-Transmission Electron Microscopy to Study Lipoplex Morphology Changes in liposome morphology were examined following the addition of plasmid D N A , oligodeoxynucleotides or sodium phosphate in the presence or absence of salt. Samples were immediately viewed by C T E M (Chapter 5). Preparation of plasmid/liposome complexes and oligodeoxynucleotide/liposome complexes was done as described above. 2.5.3.1 Preparation of Liposome/N a 2HPO 4 Mixture A 2% volume of 50x Na2HP04 solution was added to the preformed liposomes to achieve the appropriate phosphate concentration (0 to 5 m M ) . L ip id concentration was 1 m M . The mixture was allowed to incubate at R T for 30 min unless otherwise indicated. 2.5.3.2 CTEM Analysis Thin sample films (100-500 nm) were prepared under controlled temperature (25°C) and high humidity in a custom-built chamber (Figure 2.2). A small volume of sample was applied to a copper grid (Figure 2.3) that had been previously coated with the polymer cellulose acetate butyrate. The sample was then blotted to produce a thin sample film across the holes in the polymer film and immediately vitrified by rapid freezing in liquid ethane. This technique is described in detail elsewhere (Bellare et al, 1988, Dubochet et al, 1988). The grid was then transferred to a Zeiss E M 9 0 2 transmission electron microscope (accelerating voltage = 80 k V ) for viewing. Samples were kept cool to prevent formation of ice crystals on the sample surface by maintaining the temperature below 108 K during the transfer and examination procedures. 52 Figure 2.2 Controlled environment vitrification chamber (CEVS). Copper grids coated with cellulose butryl acetate are loaded with 1-2 ul of aqueous sample and blotted. The thin sample film is prepared in high-humidity controlled-temperature environment then plunged by gravity into liquid ethane (108 K) for extremely rapid freezing (vitrification). Samples are viewed immediately, with no further processing, on the cold stage of the microscope. Illustration kindly provided by Dr. Katarina Edwards. 53 Holey Polymer Film Holt sta: 1-6 pm Thickness of sample Rim: ID-5W) nm Figure 2.3 C T E M sample forms a film over the holes in the polymer -coated grid. It should be noted that in viewing the images the two-dimensional projection of a closed liposome appears as a circle with high contrast around the edge, because the projected thickness of the bilayer is greatest at the edges. A flat bilayer disk appears to have even contrast around the edges (Figure 2.4). Larger particles will tend to cluster at the edges of the film where the thickness is greater. electron beam of the microscope Image is transmitted to the detector Figure 2.4 A n illustration of how a three dimensional object is imaged in two dimensions during transmission electron microscopy. 54 2.5.4 Lipid-Mixing Assay This assay is based on resonance energy transfer (Struck et al, 1981). Nitrobenzoxadiazole-modified phosphatidylethanolamine (NBD-PE) and rhodamine-modified phosphatidylethanolamine (Rh-PE) (see Figure 2.1), each at 0.5 mol%, were incorporated into the liposomes. L ip id mixing was detected by an increase in fluorescence from the N B D - P E as it became dequenched upon dilution of the bilayer by incoming lipids. Fluorescence was detected using an excitation wavelength of 465 nm, an emission wavelength of 537 nm with an emission wavelength cutoff filter at 530 nm. The emission filter was employed to eliminate the significant contribution of light scattering from these turbid samples to the fluorescence values. Liposomes were diluted to 2.0 m M with sterile filtered 300 m M lactose, in which the molar ratio of fluorescently labeled liposomes to unlabeled liposomes was 1:1. Initially, fluorescence readings were taken continuously for 150 s. The average value multiplied by a dilution factor was calculated as mean initial fluorescence, F 0 . The dilution factor was determined by measuring the change in fluorescence as liposome-only controls were diluted by a volume of lactose equal to that of the plasmid D N A . This was necessary because dilution of the fluorescently labeled liposomes does not produce a linear reduction in arbitrary fluorescence units under these conditions. Next, an equal volume of plasmid D N A at the appropriate dilution was mixed with the liposomes to form complexes (final l ipid concentration = 1 m M ) and the readings continued for another 150 s (average value = mean fluorescence, F). A t 300 s, a 10% volume of 1% Triton X-100 was mixed with the sample in order to disperse the lipids. Fluorescence in the presence of Triton X-100 was taken to be maximal fluorescence for that sample (average value = F m a x ) . Arbitrary fluorescence units were converted to percent maximal lipid mixing by the formula: ( (F-F 0 ) / (F m a x -F 0 ) ) x 100 55 The initial fluorescence was subtracted from both F and F m a x in order to account for the slight variability in the actual fluorescence readings from sample to sample and between different batches of liposomes. L ip id mixing that occurs between cationic liposomes upon addition of polyvalent anions is extremely rapid, reaching a maximum in <1 min. Subsequently, to simplify the l ipid mixing assay, discrete fluorescence readings were taken after >2 min incubation of the newly formed lipoplexes at room temperature. For lipid mixing assays in the presence of salts, liposomes in sterile 300 m M lactose were mixed with varying amounts of D N A in H B S such that the final concentration of N a C l was 150 m M . A 10% volume of 1% Triton X-100 was mixed with the sample.. The samples were then heated in a 95°C waterbath for 2 min and vortex-mixed again, to solubilize the lipoplexes. After >2 min incubation to re-equilibrate the sample back to room temperature, the fluorescence was measured again. Liposomes at the same dilution as the complexes were included as a control, the fluorescence of which in the absence of Triton X-100 was taken to be background (F 0). With the emission wavelength filter in place, non-fluorescent lipoplexes under otherwise identical conditions did not produce a signal. Percent maximal lipid mixing reported in Chapters 4 and 6 was calculated according to the formula described above. 2.5.4.1 Multi-Step Lipid Mixing Assay Similar to the simple l ipid mixing assay described above, liposomes were prepared in 300 m M lactose and diluted with either 300 m M lactose or H B S to 2 m M lipid. The molar ratio of fluorescent to nonfluorescent liposomes was 1:1. A n aliquot of this dilution was used immediately to prepare lipoplexes in lactose or H B S , with 1 m M lipid and D N A at 100 ug/ml (10:1 l i p i d : D N A ratio, charge ratio (+/-) = 1.6). The fluorescence of the liposomes alone was read for approximately 200 s. A sample of lipoplexes prepared from the same sample of liposomes was then read immediately for another 200 s. One volume of tissue culture medium with or without 10% F B S was then added and the sample was 56 mixed by inversion, followed by fluorescence reading for another 200 s. Anionic liposomes ( D O P S / D O P C , 50/50 mol%) in a small volume were then added such that the molar ratio of D O D A C / D O P E or D O D A C / D O P C to D O P S / D O P C was 1:1, followed by mixing by inversion and a continuation of fluorescence readings for another 400 s. Finally, a 10% volume of 3% Triton X-100 was added, the sample was mixed by inversion, and the fluorescence was read for another 400 s. Controls consisted of samples prepared without D N A to account for changes in fluorescence due to sample dilutions in the course of the assay, and samples prepared without the fluorescent liposomes (same total l ipid concentration) to account for light scattering. The values obtained from the light scattering control were subtracted as background. Dilution factors were calculated from the fold signal reduction of the liposome-only control values at each step of the assay. The data plotted in Chapter 6 were representative experimental samples corrected for both light scattering and dilutions. 2.6 Characterization of DNA Protection For gene transfer to be effective, the D N A must reach its site of action intact. One reason that naked D N A is ineffective for gene transfer to most tissues [except muscle (Acsadi et al, 1991)] is that it is rapidly degraded by nucleases in the plasma compartment and interstitial spaces. Intracellular nuclease-mediated degradation or mechanical damage during sample handling would be detrimental to transfection efforts. It is therefore essential that the D N A be protected until it is released into the cytoplasm of the target cells. This can be achieved by binding the D N A to cationic liposomes. After subjecting the lipoplexes to sonication (Chapter 3) or DNase I treatment (Chapter7), D N A integrity was assessed by agarose gel electrophoresis. 2.6.1 Sonication of Lipoplexes For size reduction (Chapter 3), lipoplexes were sonicated using a microtip (diameter 1/8 inch) in 500 pi of 300 m M lactose. For sonication times < 60 s 57 samples were at room temperature, and on ice for sonication times > 60 s. The Sonifer produces ultrasonic shock waves at a rate of 20 kHz . Output was pulsed to 50 watts with pulse and rest at 1 s intervals. After sonication, the lipoplexes were disrupted by the addition of a 10% volume of lOx H B S , p H 7.4. 3 H - D N A was extracted following the method of B l i g h and Dyer as described above, precipitated by standard techniques, and run on a 0.8% agarose gel in T B E buffer (89 m M Tris-borate, 2 m M E D T A ) . The bands were excised from the gel, dissolved by heating in Solvable, and then analyzed for 3 H - D N A content by liquid scintillation counting. 2.6.2 DNase I Sensitivity DNase I is now routinely used to evaluate the stability of D N A against enzymatic degradation in the context of a carrier system to provide an indication of the potential in vivo usefulness of the carrier. The more protected and/or condensed that the plasmid D N A is by the components of the carrier, the less that DNase I w i l l be able to degrade the plasmid (Crook et al, 1997, Harvie et al, 1998). Lipoplexes containing 1 \ig plasmid were mixed with approximately 0.33 units DNase I and incubated at 37°C for 10 min. This was sufficient to degrade naked D N A into small fragments, which was included as a control. The reaction was terminated by the addition of E D T A to a final concentration of 25 m M . The D N A was then extracted by the method of Bl igh and Dyer (1959). The sample was mixed with 1 part chloroform and 2.1 parts methanol, forming a monophase upon vortex mixing. A n additional 1 part chloroform and 1 part water was then mixed thoroughly with the monophase, forming a two-phase system. The upper, aqueous phase was removed by pipette and the D N A was ethanol-precipitated by standard techniques. 58 2.7 Expression of the Transgene The endpoint of any gene therapy strategy is expression of the transgene in the cells of interest. Following successful cellular uptake of the gene carrier system, the transcription and translation machinery of the cell produces the protein product from the transgene. "Reporter genes," also known as "marker genes," evaluate the ability o f a gene carrier system to deliver exogenous D N A and achieve expression in vitro or in vivo (Mount et al, 1996). These genes encode protein products, usually enzymes, for which there are simple biochemical assays, such as (3-galactosidase ((3-gal), chloramphenicol acetyltransferase ( C A T ) , and luciferase. They share the advantage of providing a simple measure of the effectiveness of a gene therapy vector, in terms of expression of a product. A l l transfections were carried out at approximately 70% confluence. After transfection, cells were inspected by light microscopy for gross evidence of toxicity. Under the transfection conditions used in the experiments in this thesis, minimal toxicity was noted, with 0 to 15%) cell loss. Protein concentration was measured as well as an indicator of cell loss. 2.7.1 Measurement of Cellular Protein Protein concentrations of cells for all transfection experiments were measured spectrophotometrically at 570 nm using the bicinchoninic acid ( B C A ) / C u U protein assay kit (Sigma) standardized with bovine serum albumin. The B C A assay is based on the reduction of Cu(LT) to Cu(I) by protein in a concentration-dependent manner. B C A forms a purple complex specifically with Cu(I) which absorbs at 562 nm in proportion to protein concentration (Smith et al, 1985). In reporting expression of the reporter gene, the quantity of protein product was normalized to total cellular protein. The total cellular protein concentration gives an indication of toxicity, because floating dead cells and debris are removed prior to analysis. Typically, no significant difference was observed in the total cellular protein of untransfected cells (controls) vs. transfected cells. 59 2.7.2 fj-galactosidase (EC 3.2.1.23) p-gal is an enzyme found in many bacterial and eukaryotic cell types that converts o-nitrophenyl P-D-galactoside to o-nitrophenol and D -galactose. The assay for the final product is reasonably rapid and simple to perform, based on the optical density of cell lysates at 570 nm. The plasmid bearing the P-gal gene was used only in Chapter 3 of this thesis. Chinese hamster ovary (CHO) cells were plated at a density of 1500 cells/well in a 96-well plate and allowed to seed for 4 days. A t the time of transfection, the medium was removed from the wells, and 20 ul of prepared lipoplexes, mixed with 180 ul medium in a separate sterile 96-well plate, were transferred to the cell-containing plate. The cells were incubated with the complex for 48 h. The medium was removed from the wells of the microtiter plates. The wells were washed with 100 pi phosphate-buffered saline (PBS, p H 7.2) at room temperature followed by addition of 30 ul/well of lysis buffer (0.1% Triton X-100, 250 m M sodium phosphate, p H 8.0). Fifty ul of bovine serum albumin ( B S A , 0.5% in P B S , p H 8.0) were added to the cell-containing wells. For the standard curve, 50 ul of P-galactosidase standard at various concentrations were added to wells of the same plate containing only lysis buffer. To both cell-containing wells and standards, 150 ul of chlorophenol red galactopyranoside ( C P R G , 1 mg/ml in substrate buffer containing 60 m M N a 2 H P 0 4 , 1 m M M g S 0 4 , 10 m M KC1, 50 m M 0 -mercaptoethanol) were added. A s the red color developed, absorbance at 590 nm was measured then converted to units (U) of enzyme activity/well using the standard curve on each plate. The standard contained 250 to 600 enzyme units/mg protein. One unit w i l l hydrolyze 1.0 umol of o-nitrophenyl-P-D-galactoside to o-nitrophenol and D-galactose per min at p H 7.3 at 37°C. A l l lipoplexes and controls were evaluated in at least three wells on each plate and the values reported are the means of all samples from multiple plates for a given set of conditions. 60 2.7.3 Chloramphenicol Acetyltransferase (CAT) Chloramphenicol acetyltransferase (EC 2.3.1.28) is a bacterial enzyme that catalyzes acetylation of chloramphenicol with the use of acetyl-S-Coenzyme A (CoA). When the bacterial gene is inserted into a plasmid expression vector appropriate for transfecting eukaryotic cells, the C A T enzyme can be quantified by measuring its activity (Gorman et al, 1992). This assay provides better sensitivity than P-gal, however, it involves the use of radioactive materials and the assay procedure is lengthy and labor-intensive. When reporting C A T activity, the transferred cpm of radioactivity was converted to units of C A T activity based on the standard (120,000 units (U)/mg protein) One unit w i l l convert 1 nmol of chloramphenicol and acetyl C o A to chloramphenicol 3-acetate and C o A per min at p H 7.8 at 25°C. Transfection assay results using C A T as a reporter gene are described in Chapter 6. B16 /BL6 murine melanoma cells were plated at 4 x l O 3 cells/well in a 96 well plate. A t the time of transfection, media were replaced with fresh media ± 10% fetal bovine serum (FBS). Transfection proceeded for 4 h with lipoplexes containing 25 pg D N A / m l and 250 nmoles lipid/ml. Twenty p i of lipoplexes were added to each well (0.5 pg D N A / w e l l ) . Plasmid in the absence of liposomes and liposomes alone were included as controls in all experiments. C A T activity was measured based on published methods (Sleigh, 1986, Seed & Sheen, 1988). Briefly, the assay involves addition of D-threo-[dichloroacetyl-1,2-' 4C]-chloramphenicol and N-butyryl C o A to the samples. C A T exchanges the acetyl group on the radiolabeled chloramphenicol with the butyryl group of C o A . The reaction product has a greater hydrophobicity than the acetylated chloramphenicol, permitting a differential extraction in mixed xylenes. The extracted radioactive product is then measured by liquid scintillation counting. Results were quantified by comparison to a standard curve using purified C A T . 61 2.7.4 Luciferase Luciferase (EC 1.13.12.7) is a light-producing enzyme from the American firefly Photinus pyralis. The luciferase gene has been cloned and inserted into vectors for expression in bacterial or eukaryotic cells (de Wet et al., 1985). Luciferase has no mammalian endogenous counterpart, so there is no background interference (Chapman et al, 1992). Luciferase utilizes A T P as a substrate to produce light, which can then be measured in a simple luminometer (de Wet et al, 1987, Brasier et al, 1989). The assay to measure luciferase activity is extremely simple, involving adding the substrate A T P to cell lysates and measuring light production within a specific time frame. Cel l lysis buffer and A T P substrate were provided in a luciferase assay kit and used according to the manufacturer's instructions. Light intensity is proportional to luciferase concentration in the range of 10 M (10 pg/L) to 10 M ( l mg/L). Light emission increases rapidly until about 10 s after substrate addition, continuing steadily for about 5 min. After 5 min, light production is reduced with a half-life of about 5 min, therefore it is important to take the reading during the steady-state phase. In luciferase assays described in Chapters 6 and 7, B16 cells were transfected as described above for C A T assays. During luciferase activity measurements, the luminosity was measured at 1 min after addition of A T P for 30 s. 2.8 Methods of Loading Drugs into Liposomes and Lipoplexes The p H gradient method of loading ionizable drugs into liposomes was used to load amine drugs into cationic liposomes. 2.8.1 Measurement of pH Gradient Liposome ion gradients were established by the addition of an excess of H B S (pH 7.0) or C B S (pH 9.0) to the liposomes initially prepared in p H 4.0 citrate buffer. To measure the gradient, buffer containing 1 4C-methylamine ( M A ) was added (final concentration 1 uCi/ml). Final l ipid concentration was 1 m M . Where lipoplexes of 62 various charge ratios were prepared, D N A concentration varied from 25 to 100 pg/ml. C - M A was used as an ionizable uptake marker by the method of Harrigan et al. (1992). Entrapment of 1 4 C - M A in response to p H gradient (internal liposome p H 4.0, external p H 7.0 or 9.0) was measured by equilibrium filtration using MicroCon30 concentrators. The concentrators retain liposomes and their contents during centrifugation, while free M A passes through the filter. Unentrapped 1 4 C - M A was measured in the filtrates by liquid scintillation counting. This value was subtracted from total 1 4 C - M A added to the samples, to calculate the quantity of 1 4 C - M A trapped. This filtration method is analogous to the manner in which trapped volume was measured with a radiolabeled compound as described previously. The p H gradient was calculated indirectly, using the following formula: p H = l o g ( [ M A ] i n s i d e / [ M A ] 0 U t s i d e ) , where [ M A ] inside = dpm 1 4 C - M A trapped = dpm/ul trapped volume (Vt) x (lipid H dpm/lipid specific activity) and [ M A ] o u ts ide = total 1 4 C - M A added - 1 4 C - M A trapped = dpm/pl. sample volume 2.8.2 Loading of Ionizable Drugs into Cationic Liposomes or Lipoplexes 2.8.2.1 Chloroquine To drive accumulation of chloroquine into cationic liposomes or lipoplexes, the p H gradient method was used. Chloroquine in C B S (pH 9.0) was added to the cationic liposomes prepared in citrate buffer (pH 4.0). Final l ipid concentration was 1 m M and final chloroquine concentration was 10 p M , which is a 30:1 lipid:drug ratio (wt/wt). Lipoplexes were made by the addition of chloroquine in C B S buffer containing sufficient plasmid to give a 5:1 or 10:1 l i p i d : D N A ratio (nmol/pg). Samples were left at R T for 1 h then an aliquot was taken for uptake analysis. Incubation continued at 37°C for an additional 1 h followed by a second sampling for analysis. Uptake of chloroquine was measured by the filtration method. Duplicate or triplicate aliquots were transferred to MicroCon-30 tubes and 63 centrifuged at 10,000 x g for 7 min. The filtrate, containing unentrapped chloroquine, was retained for analysis by fluorescence spectroscopy, (excitation X = 330 nm, emission X = 370 nm). The lower limit of detection was set at a signal to noise ratio >5. A standard curve was done in triplicate with each assay (r > 0.99), with a linear range of 0.5 to 3 u M chloroquine. 2.8.2.2 Vincristine Loading of vincristine into cationic liposomes was done according to the method o f Webb et al. (1995). The liposomes were diluted in triplicate with citrate buffer (pH 4.0). SPH/Chol liposomes served as the positive control for vincristine loading. Vincristine was added to the diluted liposomes followed by a brief vortexing. A n aliquot of 0.5 M N a 2 H P 0 4 (j>H 9.0) was added to bring the total volume up to 1 ml (external p H = 7.4). The l ipid to drug ratio was 13:1 (wt/wt), with 3.2 m M lipid and 160 ug/ml vincristine. To make lipoplexes, plasmid first mixed with the phosphate buffer. L i p i d : D N A ratio was 10:1 (nmoles/ug, equivalent to a charge ratio of 1.3 for liposomes containing 40% D O D A C and 1.6 for liposomes containing 50% D O D A C ) . Incubation proceeded at 37°C for 4 h. Uptake o f vincristine was measured by the filtration method. Triplicate aliquots of 250 pi from each sample were transferred to MicroCon-30 tubes and centrifuged at 10,000 x g for 7 min. The filtrate, containing unentrapped vincristine, was retained for analysis by U V spectroscopy. To analyze filtrates for vincristine concentration, sample triplicates were brought up to 200 ul total volume with d H i O . Eight hundred ul of 95% ethanol were added followed by brief vortexing. Samples were spun at 3000 rpm for 5 min to pellet phosphate precipitates. The supernatant was assayed spectrophotometrically at 297 nm and corrected for background, which consisted of 200 ul sodium phosphate buffer taken through the extraction procedure. The absorbances of the three filtrates from each independent sample were averaged prior to further calculations. Absorbance readings with a signahnoise ratio >8 were converted to vincristine concentrations by comparison to a standard curve, which was linear in the range of 2 to 50 pg/ml (r > 0.99). 64 Chapter 3 Plasmid DNA Is Protected against Ultrasonic Cavitation-Induced Damage When Complexed to Cationic Liposomes* 3.1 Introduction The physical characteristics of lipoplexes are poorly understood, so it is not surprising that transfection using such lipoplexes typically suffers a high degree of variability. This lack of understanding has also hampered the pharmaceutical development of lipoplexes. One common problem, for example, is their aggregation into visible particulates when prepared at high concentrations (Holland and Huang, 1995) or in the presence of tissue culture media. Aggregation of liposomes prepared from cationic lipids and D O P E after addition of D N A is not unexpected; polymorphic, fused structures have been demonstrated by electron microscopy studies (see Chapter 5 and references therein). The multivalent ionic interactions that occur between cationic lipids and D N A would lead to concentration- and l i p i d / D N A ratio-dependent crosslinking of liposomes. Furthermore, phase separation of cationic lipids occurring as a consequence of D N A binding may generate regions rich in D O P E . It is well established that D O P E prefers to adopt a non-bilayer phase that is not compatible with retention of a typical liposome structure (Tilcock, 1986). Attempts to optimize l iposome/DNA formulations for transfection efficiency (Feigner et al, 1994) have not accounted for DNA-induced alteration in vesicle aggregation state nor the structure of the lipoplex. In addition, it is not clear what component(s) of the lipoplex is/are responsible for mediating transfection. These studies used a set of lipoplex formulations that differ in the degree of cationic * Adapted from: Wasan, E.K., Reimer, D.L. and Bally, M.B. (1996). "Plasmid DNA Is Protected Against Ultrasonic Cavitation-Induced Damage When Complexed to Cationic Liposomes" J. Pharm. Sci. 85, 427-433. 65 l ipid acyl chain saturation ( D O D A C vs. D D A B ) and in the helper phospholipid composition (DOPE vs. E P C ) . In this way, it was hoped that insight might be gained into the relationship between lipid composition and lipoplex super-aggregation, as well as its avoidance. D O P E -containing formulations were included in this study because the widely-used D O P E has been shown to have a role in mediating fusion between membranes, such as a liposome and a cell membrane (Siegal, 1986, Litzinger and Huang, 1992). E P C was substituted for D O P E as the neutral l ipid in some formulations because P C tends to be a bilayer-forming lipid, while P E is not. For this reason, it was anticipated that lipoplexes made with P C would have different physical properties than those made with D O P E . Initially it was noted that simple mixing of preformed cationic liposomes and plasmid D N A yielded variably sized lipoplexes, a phenomenon which appeared to be independent of the mixing rate (hand-pipetting or vortex-mixing). Formulations containing P E G - P E were included because previous studies had indicated that the tendency for liposomes to aggregate under conditions where crosslinking reactions occur was reduced when the liposomes contained P E G -P E (Senior et al, 1991). The hypothesis regarding the effect of P E G on lipoplex behavior was that its presence in the lipoplex might reduce particle aggregation (Hong et al, 1997). Determining the factors that influence lipoplex formation is a critical step in lipoplex development as a pharmaceutical product. These parameters must be known for optimization of stability and gene expression. For pharmaceutical purposes, a consistent and uniform size is desirable. Ideally, a mean diameter < 200 nm is preferable. It has been shown, for example, that smaller liposomes have a longer in vivo circulation lifetime (Senior et al, 1985). They also exhibit reduced uptake into the reticuloendothelial system (Gabizon et al, 1988, Gabizon et al, 1990), resulting in an increased probability o f exposure o f the tissue o f interest to the liposomal contents as compared to larger liposomes (Bakker-Woudenberg et al, 1992, Gabizon et al, 66 1992, Mayer etal, 1993). It is expected that similar trends w i l l be observed with lipoplexes injected in vivo. The experiments detailed in this chapter involve the attempt to reduce large lipoplexes into uniformly small particles by mechanical means (microprobe sonication) and also explore the role of charge ratio and l ipid composition on the resulting particle size and D N A integrity. D N A integrity was assessed following the sonication process by two methods: agarose gel electrophoresis to detect D N A breakage, and in vitro expression of the plasmid, for analysis of plasmid function. It was found that the tendency to form larger lipoplexes and the capability of sonication to reduce lipoplex size was dependent on lipid composition and charge ratio. Sonication was used successfully for size reduction of some lipoplexes with retention of D N A integrity, as assessed by gel electrophoresis and in vitro transfection of C H O cells. This implies that more commonly used pharmaceutical processes that employ cavitation or other mechanical forces as a means of particle size reduction may be useful in the preparation of lipoplexes that exhibit uniform size distributions. 3.2 Results To demonstrate the heterogeneity of particle sizes that occur with various liposome formulations, the effect of liposome composition and charge ratio on the size of the lipoplexes was investigated. Lipoplexes were made with D D A B / D O P E , D O D A C / D O P E and D D A B / E P C (all 50/50 mol%) liposomes mixed with pCMV(3 at 50 pg D N A / m l and at increasing l i p i d : D N A ratios of 2:1, 5:1, 10:1, 20:1, 40:1 nmol lipid:pg D N A / m l in a total volume of 500 pi , as described in Materials and Methods. [N.B. These ratios, a convention in this field, can also be expressed as total charge ratios (CR) defined as total moles of positive charge from the lipid 67 component/total moles of negative charge from the plasmid D N A phosphate backbone). Thus 10:1 l i p i d : D N A ratio (nmole/pg) gives a charge ratio (+/-) of 1.62 using liposomes composed of D O D A C / D O P E (1:1 molar ratio, F W = 582 and 748, respectively) and 6.5 kb double-stranded plasmid D N A (average molecular weight of D N A is 660 g/mol per base pair).] Lipoplex mean diameter was measured using quasi-elastic light scattering (QELS) (Figure 3.1). The mean diameter was not significantly different among formulations for uncomplexed liposomes (LC) , or for lipoplexes at ratios of 2:1 (CR(+/-) = 0.3), 20:1 ( C R = 3.2) or 40:1 (CR = 6.4). A t 5:1 (CR = 0.81), D D A B / E P C lipoplexes had a greater mean diameter than D O D A C / D O P E or D D A B / D O P E lipoplexes. This was also evident at the 10:1 ratio (CR = 1.62). A t higher l ipid and D N A concentrations, or in the presence of salt, aggregation was much more evident. A t C R = 1.62, variable mean diameters (200 nm to >1 pm) resulted when D N A concentration was increased from 50 pg/ml to 200 pg/ml. ^ 1000 S a w 800 u QJ 0) g 600 .2 Q 400 ca 200 0 0.4 0.81 1.62 3.2 6.4 LC Charge Ratio (+/-) Figure 3.1 Effect of formulation and charge ratio on lipoplex size was assessed by QELS analysis. Lipoplexes using DDAB/EPC (solid bars), DODAC/DOPE (open bars) and DDAB/DOPE (gray bars) were prepared with 25 ug p C M V p and increasing amounts of lipid (0.5 to 1 mM). These proportions are expressed as total charge ratio (+/-) of the lipoplexes. Data are Gaussian mean diameters +/- average SD (n=3) (See Chapter 2, section 2.3.2.1). *Mean diameters greater than 1 pm are considered aggregated. L C : uncomplexed liposome control (no DNA). 68 I T I P —I i — M , • I---A charge ratio of 1.62 was used for further studies because this ratio had been shown previously to be optimal for in vitro transfection of C H O cells with pCMV(3 complexed with cationic liposomes such as D O D A C / D O P E (Reimer et al, 1998). A t this charge ratio [ l ip id :DNA ratio of 10:1 (nmole/pg)] and a D N A concentration of 50 pg/ml, several liposome formulations differed greatly in their size upon complexation. A s shown in Table 4, mean diameters varied from approximately 300 nm for D O D A C / D O P E and D O D A C / E P C lipoplexes to >1 um, for D D A B / E P C / P E G - P E lipoplexes. LIPID COMPOSITION MEAN DIAMETER+/- SD (nm)a (mol%) LIPOSOMES LIPOPLEXES DDAB/DOPE 153 +/- 81 376 +/- 230 DDAB/DOPE/PEG 414 +/. 344 913 +/- 584 (50/45/5) DDAB/EPC 148 +/- 71 > 1 pm DDAB/EPC/PEG (50/45/5) 161 +/-58 > 1 pm (50/40/10) 135 +/- 66 > 1 pm (50/35/15) 118+/-55 > 1 pm DODAC/DOPE 218+/- 138 306+/- 178 DODAC/EPCb 121 +/-41 304+/- 155 Table 4 The effect of lipid composition on lipoplex size was investigated by preparing L U V s in lactose and performing QELS analysis on the liposomes and lipoplexes prepared from them. Notations: a: Data were representative samples, prepared at a charge ratio of 1.62. D N A concentration was 50 pg/ml; b: D N A concentration was 100 pg/ml. A s mentioned above, smaller, more uniformly sized particles are desirable for therapeutic application. Furthermore, methods that facilitate formation of uniformly-sized particles from lipoplexes prepared using a variety of liposomes, differing in l ipid composition, w i l l allow optimization studies to evaluate lipid composition effects on transfection without the complication of using lipoplexes of with a large degree of polydispersity. To determine i f sonication would be useful as a means of size reduction, the mean diameters of lipoplexes made from several different formulations were compared before and after sonication. Table 5 A shows 69 that while the mean diameters were initially quite large (>1 um) for the formulations shown, sonication for 30 s. reduced the mean diameter to less than 500 nm for D D A B / E P C and D D A B / E P C / P E G - P E lipoplexes (50/40/10 mol% and 50/35/15 mol%). While Table 5 A shows that sonication did reduce particle size, D D A B / D O P E , D O D A C / D O P E and D O D A C / E P C lipoplexes had a small initial size (<400 nm), for which 3 min of sonication provided no additional size reduction (Table 5B). For D D A B / E P C lipoplexes, increasing the sonication time from 1.5 to 6 min did not result in further decreases in mean diameter either for the lipoplexes or for uncomplexed liposomes (Table 5C). LIPOPLEXES3 BEFORE AFTER DDAB/EPC/PEG (nm) (50/45/5)b >1 um >1 um (50/40/10) >1 um 485 +/- 352 (50/35/15) >1 pm 390 +/- 235 DDAB/EPC >1 pm 444 +/- 241 Table 5A The effect of sonication on the size of lipoplexes. After 30 s microprobe sonication, QELS analysis was performed. Data are Gaussian mean diameters (nm) +/- SD of representative samples. Notations: a: lipoplexes prepared with 250 nmol lipid/ml and 25 ug DNA/ml (CR=1.62); b: ratios indicate mol%. LIPOPLEXES BEFORE AFTER DDAB/DOPE 376 +/- 230 441 +/- 256 DODAC/DOPE 306 +/- 178 306 +/- 176 DODAC/EPCa 304+/- 155 398 +/- 263 Table 5B. Experiment was the same as in (A) above, except sonication time = 3 min. Notation: a: D N A concentration was 50 pg/ml. SONICATION TIME (min) DDAB/EPCa BEFORE 1 1.5 3 6 >1 pm nd 277+/- 132 292 +/- 165 455 +/- 371 LC 132+/- 77 110+/- 54 107+/-53 92 +/- 45 nd Table 5C Experiment was the same as in (A) except sonication time was varied as indicated. Data represent Gaussian mean (nm) +/- average SD for 2 separate preparations. Notations: a: lipoplexes were prepared with 125 nmol lipid/ml and 12.5 ug DNA/ml; L C : uncomplexed liposome control; nd: not done. 70 Because sonication might be useful to effect lipoplex size reduction for certain lipid compositions, it was important to know i f D N A were being damaged during the process. Plasmid D N A integrity was evaluated by standard 0.8% agarose gel electrophoresis following extraction of the D N A from the lipoplexes. Figure 3.2 shows photographs of agarose gels of D N A extracted from lipoplexes prepared with D D A B / D O P E (Figure 3.2A) and D D A B / E P C (Figure 3.2B). Lipoplexes were sonicated for 0, 1.5, 3 or 6 min. The resulting gel showed that uncomplexed plasmid D N A was nearly completely sheared by 1.5 min sonication (lanes 11, 12). When complexed with liposomes the D N A was not damaged by the extraction process, although a slight shift in relative proportions of supercoiled vs. relaxed plasmid was noted (lanes 2,3). The reason for the shift is not clear. Increasing sonication time, however, resulted in a decrease in band intensity and an increase in smearing at the bottom of each lane (lanes 4-9). Importantly, a significant portion of the D N A was seen to be intact even after 6 min of sonication when complexed to liposomes (lanes 8-9). It would appear that the D N A from D D A B / D O P E lipoplexes was more sensitive (less protected) to the sonication procedure than the D N A from D D A B / E P C lipoplexes. In comparing D O D A C / D O P E vs. D O D A C / E P C sonicated lipoplexes (Figure 3.2C), D O D A C / E P C appeared to be more protective of the D N A from damage than D O D A C / D O P E . This can be seen by the increased smearing in lane 5 (sonicated D O D A C / D O P E lipoplexes) compared to lane 3 (sonicated D O D A C / E P C lipoplexes). These results suggest that EPC-containing formulations may be more stable. In order to distinguish the relative differences in D N A protection following sonication of different lipoplexes, the amount of intact D N A was quantified. L ip id and D N A concentrations in the lipoplexes ( 3 H - D N A and 1 4 C-l ip id) were determined prior to extraction and just prior to loading the samples on the gel. After electrophoresis, the bands were excised from the gel, along with their corresponding wells. The band and well were separately dissolved and radioactivity 71 determined by scintillation counting. 3 H - D N A counts were then compared with the pre-gel counts to determine the percent of D N A recovered that migrated to the same distance on the gel as untreated plasmid (band %). This was interpreted to be reflective of the percent of plasmid D N A that was intact. It should be noted, however, that approximately 50% of the D N A was consistently lost by the extraction process prior to gel electrophoresis. The amount of D N A lost in the extraction of D N A from control and sonicated samples was comparable. For this reason, this assay is only an approximation of D N A integrity in each sample. A s indicated in Table 6, 83%> of the counts loaded on to the gel were accounted for in the banded fractions (relaxed + supercoiled) when intact, uncomplexed, extracted plasmid D N A was evaluated. When sonicated, however, only 4% of the counts were recovered in the band. Similar results were obtained for unsonicated D D A B / D O P E lipoplexes, where 99%> of the D N A loaded onto the gel migrated as intact D N A . The results indicate that the percent of D N A which remains intact through the sonication process depended on the lipid formulation used in the lipoplex as well as the charge ratio of that specific formulation (Table 6). When D D A B / D O P E and D D A B / E P C lipoplexes at varying charge ratios were sonicated and the D N A assayed by gel electrophoresis, D N A from the sonicated D D A B / E P C lipoplexes at a charge ratio of 1.62 yielded the lowest percent of intact D N A (20%), and a charge ratio of 3.2 yielded the highest percent (43%). For D D A B / D O P E lipoplexes, however, a ratio of 0.81 showed no D N A intact, and only 18%> intact when a charge ratio of 6.4 was used. 72 3 I Figure 3.2 DNA integrity was assessed by agarose gel electrophoresis. Lipoplexes were prepared at a charge ratio of 1.62 [lipid:DNA ratio of 10:1 (nmol/ug)] with DDAB/DOPE liposomes and pCMVp. A, 125 nmol: 12.5 pg) and DDAB/EPC (B, 250 nmol:25 pg). Untreated pCMVp (lane 1), lipoplexes without sonication (lanes 2,3) and uncomplexed, unsonicated, extracted pCMVp (lanes 10,11) serve as controls. Lipoplexes were sonicated 90 s (lanes 4,5), 3 min (lanes 6,7), or 6 min (lanes 8,9). Uncomplexed pCMVp was sonicated for 90 s (lanes 12,13). C: Untreated pCMVp (lane 1) and lipoplexes without sonication (lane 6) serve as controls. Lipoplexes (250 nmol lipid:25 pg DNA) using DODAC/DOPE (lane 2 -not sonicated, lane 3- sonicated) and DODAC/EPC (lane 4 -not sonicated, lane 5 -sonicated) were evaluated. Lane 7-sonicated, uncomplexed plasmid. Sonication time = 3 min. In A, B and C, top DNA band: relaxed plasmid; bottom DNA band: supercoiled plasmid. 73 S A M P L E B A N D W E L L % % untreated D N A 83 0 lipoplexes, no sonication 99 0 sonicated D N A 4 0 sonicated D N A +liposomes 2 0 D D A B / D O P E Charge ratio (+/-) 0.81 0 100 1.62 8 2 3.2 8 15 6.4 18 42 D D A B / E P C Charge ratio (+/-) 0.81 38 28 1.62 20 79 3.2 43 57 6.4 39 25 Table 6 Semiquantitation of D N A integrity following lipoplex sonication was determined. Free plasmid and lipoplexes, prepared with 3 H - D N A , were sonicated for 90 s, followed by D N A extraction and gel electrophoresis. Band% refers to the percent of total 3 H dpm loaded that migrated the same distance as intact, untreated plasmid. Well% refers to the percent of counts that remained in the loading well of the gel. A s indicated in Table 6, a significant proportion of the counts recovered in the gel was present in the well . This was only observed when evaluating sonicated samples. It is unknown how much D N A within the wells was intact, therefore, the data presented may be an underestimate of the fraction of D N A which was intact. The significance of the sonication-induced increase in D N A retention in the wells is not understood. For PEG-PE-containing lipoplexes, results were inconclusive. Both the D N A extraction efficiency and percentage of intact D N A were highly variable between experiments. Within a given experiment the percentage of intact D N A after sonication was similar to that of D D A B / E P C without P E G - P E . On visualization of the gels, in several experiments there appeared to be slightly less D N A protection from sonication damage when P E G - P E was present, but in others there was no apparent difference. Comparisons between experiments could not be done because of the differences in plasmid extraction efficiency. Including P E G - P E in the 74 liposomes did not give a clear advantage in D N A protection from sonication damage compared to liposomes without P E G - P E . While an agarose gel can give information about how much of the total plasmid is intact, demonstration of functional integrity is also important. To determine i f plasmid D N A subjected to sonication while in lipoplexes was still biologically functional, transfection of C H O cells in vitro was performed and (3-galactosidase expression levels were assayed (Figure 3.3). Figure 3.3 In vitro transfection of CHO cells was done with sonicated vs. unsonicated lipoplexes. Liposome formulations evaluated include DDAB/DOPE (n=17), D D A B / E P C (n=17), DODAC/DOPE (n=6) and DODAC/EPC (n=6). CHO cells were transfected with unsonicated lipoplexes (solid bars) and sonicated lipoplexes (hatched bars) (lipid:DNA ratio = 10 nmol/ug; 50 pg DNA/ml) and assayed for P -galactosidase activity. Data are expressed as mean +/- SD (n = 6). Sonicated DODAC/DOPE lipoplexes were significantly different from unsonicated DODAC/DOPE lipoplexes (by one-way Anova, p<0.05). Transfection with unbound D N A either with or without sonication did not result in detectable levels of P-galactosidase expression. DDAB/DOPE DDAB/EPC DODAC/DOPE DODAC/EPC Liposome Formulation 75 Lipoplexes were prepared at a charge ratio of 1.62 and a D N A concentration of 25 pg/ml. Twenty pi of these lipoplexes (0.5 pg D N A ) were mixed with media and added to the cells. The cells were then incubated for 48 hours prior to assaying p-galactosidase activity. Figure 3.3 shows that sonication did not reduce the extent of expression from p C M V p under the conditions used here. Uncomplexed D N A with or without sonication did not result in measurable P-galactosidase expression. The lowest levels o f expression obtained were seen with lipoplexes formed using liposomes composed of D D A B / D O P E / P E G - P E and D D A B / E P C / P E G - P E , with P-galactosidase <0.1 mU/wel l with or without sonication. The highest levels of expression were seen with D D A B / D O P E lipoplexes both with and without sonication (>3.1 mU/well) . It is important to note that the amount of D N A added per well was based on initial D N A concentration and was not adjusted according to the estimated loss of D N A integrity for those samples that were sonicated. The results in Table 6, for example, suggest that as much as 90% of the D N A present in the sonicated D D A B / D O P E lipoplexes was fragmented. It is surprising that the significant loss of D N A integrity had no measurable impact on in vitro transfection efficiency. 3.3 Discussion Lipoplexes made from preformed liposomes and plasmid D N A , while potentially useful for in vivo gene therapy, have a tendency to aggregate. A n attempt was made to reduce the size of the lipoplexes by sonication, and D N A integrity was assessed following this process. Sonication is a commonly employed method for liposome preparation on a laboratory scale (Cullis et al, 1987, Memol i et al, 1995, Arnardottir et al, 1995). It is also used for the disruption of cells and tissues (Crouse et al, 1993, Nielsen et al, 1992). This disruption occurs via the intense mechanical pressures placed on the sonicated medium as gaseous cavitation 76 bubbles produced by oscillations in the medium collapse. This process also produces chemically reactive species such as H2O2 and free radicals, which can also have a significant impact on the sonicated material (Lepoint and Mul l ie , 1994, Chivate and Pandit, 1995). Sonication has been demonstrated to cause D N A strand breaks (Kondo et al, 1985, Mi l l e r et al, 1991a, Mi l l e r et al, 1991b). The hypothesis was that binding of plasmid D N A to cationic liposomes would protect it from damage due to sonication. These experiments determined that not only lipoplex size but also resistance to D N A damage upon sonication was dependent on l ipid composition. The results presented here suggest that D N A was protected from damage due to ultrasonic cavitation when complexed to cationic liposomes. The potential for using other manufacturing processes to effect size reduction of lipoplexes is also briefly discussed. Initially, the effect of liposome composition on the size of lipoplexes was determined. A high degree of variability was found in the mean diameters (see Table 4), although the initial size of all o f the liposomes was small (-150 nm) and reasonably uniform (SD typically 50-60%). Following sonication, mean lipoplex diameter also varied by l ipid composition. In the Q E L S analysis a size reduction was observed for D D A B / E P C and D D A B / E P C / P E G - P E lipoplexes, which were initially large lipoplexes, but not D D A B / D O P E , D O D A C / D O P E or D O D A C / E P C lipoplexes, which were initially small (see Table 5). The results suggest that lipoplexes formed with D N A and cationic liposomes may result in particles that adopt a minimum mean diameter of 300-400 nm. Regardless, lipoplexes were prepared using cationic l ip id /EPC mixtures that exhibited the same size distribution as cationic l ip id /DOPE mixtures using ultrasonic cavitation. It may be suggested that alternative methods of size reduction which employ other types of mechanical force (such as shear force), for example, homogenization (Zheng et al, 1992), high-pressure extrusion (Vidal-Naquet et al, 1989) or microfluidization (Mayhew et al, 1984, Zheng et al, 1994) may also be able to produce lipoplexes of small, uniform diameter. 77 When comparing the particle size of the different lipoplexes used in this study, it should be noted that the charge ratio at a given l i p i d : D N A ratio is the same for the different formulations used. The interaction between the negatively charged D N A and the positively charged lipid is clearly affected by the choice of neutral l ipid (DOPE, E P C , and/or P E G - P E ) and by the specific cationic l ipid used (the saturated D D A B vs. unsaturated D O D A C ) . This is illustrated by the comparison of particle size estimated for the different formulations at equivalent l i p i d : D N A ratios [Figure 3.1, at 5:1 (CR = 0.81) and 10:1(CR = 1.62)]. D D A B / E P C lipoplexes, for example, tend to have large mean diameters, while D O D A C / E P C lipoplexes remain relatively small. This may be anticipated based on the effects of unsaturation on l ipid phase separation in membranes (Cullis et al, 1986). Because unsaturated lipids have a tendency toward increased mobility in a bilayer plane, D N A binding may elicit phase separation and binding of D O D A C to the D N A . Less D O D A C , therefore, would be available for crosslinking reactions between liposomes. In contrast, the saturated, monovalent, cationic l ipid D D A B would exhibit reduced mobility, and hence, reduced binding to D N A in terms o f the number o f cations bound per liposome. This would enhance the propensity for liposome-liposome crosslinking via D N A bridges. Another important component of liposome formulations for D N A transfection is D O P E , which prefers to adopt a hexagonal rather than a bilayer configuration. It is important to recognize that the monovalent cationic lipids used in combination with D O P E function not only to complex D N A , a polyvalent anion, but also stabilize D O P E in a bilayer configuration. Based on the discussion above, the unsaturated monocationic l ipid D O D A C may (due to potential phase separation characteristics) be less able to maintain the stability of the liposome membrane after addition of D N A . It has been shown previously that various PE-containing anionic liposomes are destabilized and fuse in the presence of C a 2 + and other multivalent ions that induce Hu phase 78 formation (this w i l l be discussed in Chapter 6). For such systems, sonication may lead to further destabilization and subsequent D N A damage. In the presence of P E G - P E , a l ipid that was anticipated to interfere with l iposome-DNA binding, D N A binding still occurs, however, more of the D N A is exposed, as indicated by sonication-induced fragmentation of complexed D N A . This situation may make it more likely that a given lipoplex with surface-available D N A is attracted to another particle, eventually leading to increased aggregation. The PEG-PE-containing lipoplexes would be more easily reduced in size by sonication because they likely have a weaker interparticle attraction due to steric hindrance from the P E G . Since the bound D N A was not well-protected, however, the size reduction process employed here may not be appropriate. After sonication of the lipoplexes, it was critical to assess whether the plasmid D N A was remaining intact and functional. Secondarily, it was important to determine whether liposome composition had any influence on the degree of plasmid D N A damage. Results from the gelelectrophoresis analysis showed that protection of the D N A from ultrasonic cavitation-induced D N A fragmentation was dependent on the l ipid composition. D D A B / E P C and D O D A C / E P C provided better protection than D D A B / D O P E and D O D A C / D O P E , respectively (Figure 3.2). PEG-PE-containing liposomes provided less protection against degradation than their non-PEG-PE-containing counterparts. More extensive degradation of the plasmid, as assessed by agarose gel electrophoresis was observed for the lipoplexes prepared with P E G - P E -containing liposomes. A higher charge ratio (+/-) improved protection of the D N A for D D A B / D O P E lipoplexes, but no such ratio dependence was seen for those made with D D A B / E P C (Table 6). The transfection data were consistent with the results discussed above, and indicated that after sonication of the lipoplexes, expression of the reporter gene was achievable (Figure 3.3). 79 D N A can be damaged in a number of ways during sonication. Ultrasonic cavitation can produce many effects on a D N A molecule, including double strand breaks, single strand breaks, rupture of hydrogen bonds, base destruction and possibly crosslinks. These effects may be direct, due to the physical force placed on the molecule, or indirect, secondary to the production of highly reactive sonochemicals such as hydrogen peroxide (Lepoint and Mul l i e , 1994, Mi l l e r et al., 1991b). Transfection assays therefore were performed in order to establish that the D N A in a sonicated lipoplex was still biologically functional. In vitro transfection of C H O cells comparing sonicated and unsonicated lipoplexes showed no detectable compromise of transfection ability due to sonication of D D A B / D O P E , D D A B / E P C , D O D A C / D O P E and D O D A C / E P C formulations. The DOPE-containing formulations yielded higher P-galactosidase expression levels than those containing E P C as the neutral l ipid, consistent with previous studies. From this study three main conclusions may be drawn. First, lipoplexes made from simple mixing of preformed liposomes and plasmid D N A are highly unpredictable with respect to lipoplex size and further control of a uniform particle size distribution as would be required for most pharmaceutical applications would be difficult. Particle size may influence transfection efficiency. Second, P E G - P E , predicted to inhibit interparticle interactions due to its hydrophilic and steric properties, was not effective in preventing aggregation of lipoplexes with the lipid compositions used here. (The use of PEG-l ipids is further addressed in Chapter 7) Third, certain liposome formulations, when bound to the D N A , w i l l protect the plasmid from sonication damage. Microprobe sonication may not be the most practical approach in large-scale settings for size reduction of lipoplexes. However, the results described here indicate that commonly employed pharmaceutical techniques, such as microfluidization or homogenization may have potential in this context for the formation of small, uniform particles. 80 Chapter 4 Cationic Liposome-PLasmid DNA Complexes Used for Gene Transfer Retain a Significant Trapped Volume* 4.1 Introduction A s indicated in Chapters 2 and 3, typical lipoplexes preparations are a heterogeneous mix of particles. The morphology of lipoplexes is highly variable (Sternberg et al, 1998 and references therein). Large fused liposome-like structures, long tube-like strands, stacks of flattened discoids and multilamellar structures with D N A intercalated into the l ipid bilayers have been observed by different investigators. Morphology appears to be highly dependent on multiple factors, including l ipid to D N A ratio, composition, environmental conditions and the technique used to make the observations. These issues are addressed in Chapter 5. It is believed that the observed changes in the vesicles' shapes are due to DNA-induced membrane disruption and/or fusion, which are likely consequences of stresses upon the liposomal membranes arising from the polyelectrolyte interactions. It is unknown, however, to what degree liposomal bilayer integrity is compromised by the interaction of liposomes and plasmid D N A . The goal of the experiments described in this chapter was to determine whether cationic liposomes retained any trapped volume after their complexation to plasmid D N A . This serves two purposes: to further the understanding of the physical nature of lipoplexes used in gene therapy, and to investigate the potential for codelivery of other encapsulated molecules within the lipoplexes. Cationic liposomes composed D O D A C / D O P E (50/50 mol%) encapsulating an aqueous trap marker were used to prepare lipoplexes at various charge ratios. * Adapted from Wasan, E.K. , Fairchild, A . and Bally, M . B . (1997) "Cationic Liposome-Plasmid D N A Complexes Used for Gene Transfer Retain a Significant Trapped Volume" J. Pharm. Sci. 87(1), 9-14. 81 To begin to answer some of the questions about changes in membrane structure that occur when D N A is added to pre-formed cationic liposomes, liposome integrity was assessed by measuring trapped volume before and after complexation to D N A . If there were a complete collapse or lysis of the liposomes when the cationic lipids bind to D N A , then loss of entrapped contents should be observed after formation of the lipoplex. It was anticipated that at charge ratios at which the greatest aggregation occurs (CR « 1) (Wasan et al, 1996, Bal ly et al, 1997, Radler et al, 1997), the loss of entrapped contents would be most evident. The effect of tissue culture medium on trapped volume was also investigated to determine i f media would further disrupt liposome integrity. This was important for the long-term goal of developing codelivery systems for secondary agents within the lipoplexes. A lipid-mixing assay was used to further characterize the aggregation events that influence trapped volume. Aggregation of liposomes could occur without further membrane interactions, however, evidence of morphological changes as observed by electron microscopy (EM) suggested that fusion of liposomal membranes could happen when lipoplexes form (see Chapter 5). L ip id mixing assays in this chapter comprise a preliminary investigation of the role of charge ratio in lipoplex l ipid mixing, to be further addressed in Chapter 6. Results presented here demonstrate loss of an aqueous trapped marker, consistent with the l ipid mixing assay findings, which indicate that liposomal membrane rearrangement occurs during lipoplex formation. Surprisingly, however, a significant trapped volume does remain in the lipoplexes. This result not only gives information about their structure, but also raises the interesting possibility that other bioactive molecules may be delivered at the time of transfection. 4.2 Results The trapped volume (Vt) of cationic and neutral liposomes in the presence and absence of plasmid D N A was investigated. Prior to this study, the trapped volume of these cationic 82 liposomes or any formulation of lipoplexes had not been reported. Cationic liposomes consisting of D O D A C / D O P E (50/50 mol%) or E P C were prepared in 300 m M lactose by extrusion. Trace amounts of 1 4C-D-lactose were added to the hydrating lactose solution. Upon liposome formation, the 1 4C-D-lactose became passively encapsulated within the liposomes and served as an aqueous trapped marker. In preliminary experiments, following dialysis to remove unencapsulated lactose, lipoplexes were prepared with 25 pg D N A / m l and a charge ratio (+/-) (CR) = 1.62. Residual trapped volume (RVt), the trapped volume remaining after addition of D N A , was assessed by filtration as described in Chapter 2 (section 2.5.2.1). The trapped volume of D O D A C / D O P E liposomes in the absence of D N A was 1.45 ± 0.46 ul/umol, while the trapped volume of E P C liposomes was 0.90 ± 0.24 ul/umol (Figure 4.1). Figure 4.1 The effect of binding to D N A on liposomal trapped volume. Trapped volume (Vt) of 2.0 -1.8 -$ 1 - 6 -O £ 1 . 4 -ZL 3 . 1.2 -O 0.8 -~§ 0.6 -Q_ Q. 03 0.4 -I— 0.2 -0.0 - • (") ( + ) cationic liposomes composed of DODAC/DOPE (black bars) or EPC (gray bars) was measured by the filtration method using 14C-lactose as an aqueous trapped marker. Vt was measured in the absence (-) and presence (+) of 25 pg DNA/ml. Data are means ± SD of replicate experiments. For (-), n = 41 (DODAC/DOPE) and n = 44 (EPC). For (+), n = 4 to 6 experiments. There was no statistical difference in Vt between liposomes with and without added D N A . 83 In the presence of 25 pg D N A / m l , R V t was 0.52 ± 0.48 and 0.99 ± 0.03 pl/pmol for D O D A C / D O P E and E P C , respectively. Addition of D N A to E P C liposomes had no effect on trapped volume. The dependence of R V t on lipoplex charge ratio was then investigated. Previous studies in our laboratory and others have demonstrated that the physical properties of the lipoplex, such as particle size, vary with charge ratio (Figure 4.2) (Wasan et al, 1996). E P C (neutral) liposomes, which do not bind D N A , and D O D A C / D O P E cationic liposomes were each mixed with D N A at l i p i d : D N A ratios of 8:1 (CR =0.9 for D O D A C / D O P E lipoplexes), 10:1 (CR = 1.62), 13:1 ( C R = 2.1), 20:1 (CR = 3.2), and 40:1 (CR = 6.4) nmoles lipid/pg D N A . Addition of D N A to E P C liposomes resulted in no change in particle size. However, increases in the mean diameter of the D O D A C / D O P E - D N A lipoplexes indicate that vesicle-vesicle aggregation occurred as D N A concentration was increased from 0 to 125 pg/ml (Figure 4.2A). In addition, close membrane contact between vesicles during the formation of the lipoplexes resulted in l ipid mixing (Figure 4.2B and 4.2C). The observation of l ipid mixing can also occur as a result of either complete membrane fusion, hemi-fusion, or less likely, l ipid exchange. A t a constant lipid concentration of 1 m M and increasing D N A concentrations, the percent of maximal l ipid mixing steadily increased until reaching a plateau, generally at the point at which visible aggregates began to form. Fluorescence readings decrease when gross flocculation occurs. 84 Figure 4.2 Lipid:DNA ratio determines degree of aggregation (A) and lipid-mixing (B, C). A : Mean diameter of DODAC/DOPE lipoplexes at increasing D N A concentration. Lipid concentration was 500 nmol/ml. Data are representative samples, with error bars referring to the intra-sample SD. B: DODAC/DOPE liposomes undergo lipid-mixing upon addition of D N A at the indicated lipid:DNA ratios. Number in parentheses refers to total charge ratio (+/-). C: Example tracing of continuous fluorescence readings during the course of the reaction. At t = 0 (a), the fluorescence of DODAC/DOPE liposomes (1:10 fluorescent to nonfluorescent molar ratio) was read continuously. At (b), D N A was mixed with the liposomes in the cuvette (lipid:DNA ratio = 10:1). At (c), a 10% volume of Triton-X-100 was added. 85 It was hypothesized that loss of trapped marker would be greatest .at charge ratios at which aggregation/lipid mixing reactions occur most frequently, i.e. C R « 1. R V t was assessed for lipoplexes prepared at different l ipid to D N A ratios (charge ratios of 0.32 to 6.48), where the D N A concentration was constant (25 pg/ml) (Figure 4.3). Liposome-only controls were also prepared using D O D A C / D O P E and E P C at the same lipid concentrations as the lipoplexes. R V t was measured by two different methods, filtration (Figure 4.3A) and dialysis (Figure 4.3B), as described in the (sections 2.5.2.1 and 2.5.2.2, respectively). N o significant nonspecific binding of 1 4 C lactose to the liposomes, the Mic roCon filters or the dialysis tubing was observed. Two or three separate preparations of liposomes were used. A l l experiments were performed with triplicate samples from each set of liposomes. Means of the trapped volume vs. charge ratio data were tested for significant differences by one-way analysis of variance, using the Microcal Origin 4.1 computer program (Microcal Software, Northampton, M A ) . There was no significant difference (at p<0.05) between means within each ratio group or between liposome sets, and so the data for all replicates at a given ratio were combined for each method. The results show that R V t was dependent on the charge ratio for D O D A C / D O P E - D N A lipoplexes. The lowest trapped volumes for D O D A C / D O P E - D N A lipoplexes were observed at l i p i d : D N A ratios of 2:1, 5:1 and 10:1 nmoles lipid/pg D N A , which correspond to charge ratios of 0.32, 0.81 and 1.62. The largest trapped volume for D O D A C / D O P E - D N A lipoplexes was observed at a charge ratio of 6.4, the highest ratio used. There was no difference in the R V t values at each ratio between the filtration and dialysis methods. These data indicate that a trapped volume was present in the cationic liposomes after complexation to D N A at all of the charge ratios tested. o E a 2.0-1.8 1.6 1.4 1.2-1.0 0.8-0.6-0.4 0.2-0.0 1.6 1.4 1.2 1.0 0.8 0.6 0.4 0.2 0.0 B i i ML - 1 — 1 — i — • — i — 1 — i — 1 — i — 0 2:1 5:1 1 — i — < — i — ' — i — < — i — • — i — r _ 10:1 20:1 40:1 —[ 1 1— — i — , — | — i — ^ — i — | — i — | — , — | — i — | — i — | — i — | — i 0 2:1 5:1 10:1 20:1 40:1 (0.3) (0.8) (1.6) (3.2) (6.4) Lipid:DNA Ratio (nmohjig) (charge ratio +/-) Figure 4.3 Trapped volume of lipoplexes depends on lipid:DNA ratio. Black squares: DODAC/DOPE lipoplexes. Gray triangles: EPC liposomes. Vt was measured by filtration (A) or by dialysis (B). * Significantly different from liposome-only controls (p <0 .05). Data are means ± SD, n = 6 independent experiments. [DNA] = 25 pg/ml for all ratios. Charge ratio = 0 refers to liposome-only controls. 87 The use of lipoplexes to transfect cells in vitro or in vivo necessitates the presence of salts and serum proteins. These lipoplexes, however, may undergo further aggregation when diluted into saline solutions (Figure 4.4A). Centrifugation of the lipoplexes resulted in pelleting of lipid 5:1 10:1 20:1 DNA Ratio (nmolesr^g) 40:1 50 75 100 125 D N A O g / m l ) Figure 4.4 Lipoplex aggregation is associated with l ip id mixing. The presence o f salt (HBS) causes increased aggregation (A) and l ip id mixing (B) of D N A /liposome complexes. A: Sedimentation o f the complexes upon centrifugation. Percent l ip id in the supernatant is the fraction of the sample which is not aggregated. L C * : Liposome control centrifuged in the absence of D N A or H B S . L C * * Liposome control centrifuged in the absence o f D N A and the presence o f H B S . [ D N A ] = 25 pg/ml. [NaCl] f i n a i = 90 m M . Data represent means ± SD (n=3). Dotted line is drawn to help illustrate the trend. B: Percent maximal l ip id mixing of D O D A C / D O P E liposomes upon complex formation in the presence o f 150 m M H B S . L i p i d concentration = 1 umole/ml. Data represent mean ± SD, n=3. 88 and D N A under conditions that generate large particle diameters according to Q E L S analysis. Figure 4.4A shows the effect of H B S on lipoplex aggregation. Charge ratios greater or less than 1.6 (10:1 l i p i d : D N A ratio on the graph) exhibited less sedimentation, indicating that lipoplex density and aggregation varies by charge ratio. L ip id mixing assays were also performed in the presence of salts. It was found that the overall l ipid mixing was less (lower percent maximal l ipid mixing), but this maximum was reached at lower D N A concentrations when D O D A C / D O P E - D N A lipoplexes were formed in the presence of salt (Figure 4.4B) rather than lactose (Figure 4.2B). If this l ipid-mixing assay was a reflection of the extent of membrane fusion, then after addition of D N A the residual fusogenic ability of the cationic liposomes was reduced. The impact of this factor on transfection is unclear (Stegmann et al, 1997) but it is addressed in more detail in Chapter 6 of this thesis. It was anticipated that the presence of tissue culture medium would lead to increased loss of entrapped contents from the lipoplexes, based on the aggregation that occurs in the presence of salts. To test this hypothesis, lipoplexes were prepared at a charge ratio of 1.62 and a D N A concentration of 25 pg/ml, conditions which in previous studies in our laboratory have been determined to be optimal for in vitro transfection (Reimer et al, 1997). These lipoplexes were diluted 1:1 with tissue culture medium with or without 10% F B S and incubated for 4 h at either room temperature or 37°C. A t the end of the incubation, residual trapped volume was measured by the filtration method (Figure 4.5). No difference was seen in the R V t of lipoplexes between groups incubated with or without serum, or between the groups incubated at room temperature (Figure 4.5A) or 37°C (Figure 4.5B). A s anticipated, D O D A C / D O P E liposomes in the absence of D N A showed loss of contents, as a consequence of salt-mediated aggregation. Importantly, the lipoplexes showed no further loss of entrapped contents beyond that shown by the liposomes alone. 89 DODAC/DOPE EPC + DNA DODAC/DOPE + DNA EPC DODAC/DOPE EPC + DNA DODAC/DOPE + DNA Figure 4.5 Tissue culture medium does not significantly affect trapped volume of liposomes or lipoplexes, at 23°C (A) and 37°C (B). Dark bars: without serum; light bars: with 10% serum. Data are means ± SD, n=3. [DNA]=25 pg/ml. Charge ratio = 1.62. RVt was measured by filtration. 4.3 Discussion These experiments yielded the surprising result that a significant proportion (nearly 30%>) of encapsulated aqueous contents remains associated with D O D A C / D O P E - D N A lipoplexes at charge ratios that promote the greatest aggregation. It was anticipated that nearly complete loss of liposomal contents would occur following complexation based on the major morphological changes that the cationic liposomes undergo following binding to the plasmid D N A . A s shown 90 in Figure 4.2, such changes include aggregation and generation of structures where lipid mixing is facilitated. Considering that the initial trapped volume of D O D A C / D O P E liposomes was 1.45 ± 0.46 pl/pmol, this residual volume is nontrivial. R V t is lowest when the charge ratio of the lipoplex approaches 1 and it is greatest when there is excess positive charge (e.g. excess liposomes). It is likely that these free liposomes can retain their trapped contents because they are not subjected to the stress of the multivalent electrostatic interactions between the cationic lipids of the liposome and the plasmid D N A , which result in membrane rearrangements. Near charge neutrality, it may be suggested that l iposome-DNA binding is optimal and that at this ratio vesicle-vesicle interaction is greatest. This idea is supported by the demonstration of changes in lipoplex mean diameter as a function of charge ratio (Figure 4.2A). This aggregation trend was evident under conditions where salts are present (Figure 4.4A), in agreement with other studies (Sternberg et al, 1998). The D N A , as it is sandwiched in between liposomal bilayers or sheets, promotes the formation of nonbilayer structures associated with membrane fusion. This, in turn, would be accompanied by loss of entrapped contents. Previous studies not involving D N A interactions have demonstrated the loss of cationic liposome contents upon liposome-liposome membrane fusion (Stamatatos et al, 1988). It is not clear whether membrane fusion and content release occur simultaneously (Ellens et al, 1986), however, it is likely that both events are a consequence of vesicle aggregation promoted by addition of D N A to the cationic liposomes. Aggregation can lead to close membrane contact and perhaps dehydration, both requirements for membrane fusion. The fusion process can create membrane defects that allow entrapped contents to be released. Although vesicle aggregation reactions are enhanced in the presence of salt and serum proteins, this did not lead to additional loss of trapped contents beyond that observed for liposomes alone (Figure 4.5). The lower maximal l ipid mixing observed in the presence of salts may be due to an inhibition of 91 the D N A - l i p i d interaction because of charge shielding. The significance of the results presented here is that a measurable portion of the entrapped contents is retained in spite of the potentially disruptive multivalent interactions that occur as lipoplexes form. These results encourage the pursuit of developing codelivery systems consisting of a bioactive molecule entrapped within the lipoplexes. The most obvious choice of molecules to consider for codelivery are those which are known to enhance transfection. Weak bases such as chloroquine have been shown to raise lysosomal p H , such that lipases and nucleases are inactivated (Dean et al, 1984). Thus, treatment with chloroquine is thought to improve transfection of some cell lines by preventing lysosomal degradation of the lipoplexes (Erbacher et al, 1996). Improving delivery of the enhancer and the lipoplexes to the cells simultaneously may result in improved efficacy and even potency of the transfection enhancer. Alternatively, one may consider loading the liposomes with pH-sensitive fusion peptides or polymers that destabilize endosomes or lysosomes (Hughes et al, 1996), thereby promoting release of the lipoplex or its constituents into the cytoplasm. Transfection-enhancing agents that promote lysis of the endosomes/lysosomes by osmotic mechanisms, such as sucrose or polyvinylpyrrolidone (Cifti et al, 1996), represent additional candidate molecules that can be encapsulated into liposomes. Improved delivery of transfection enhancers may overcome some of the cell type-specific differences in the response to these agents i f they are due to differences in uptake of the enhancer or in release of endosomal contents, for example. The choices are numerous, and they are limited only by one's imagination and the ability to load the compounds into the cationic liposomes. It may even be possible to improve loading of ionizable compounds into these cationic liposomes, beyond that achievable by passive trapping as described here, through the use of the ion gradient method based on active loading procedures (Madden et al, 1990). The development of cationic liposome formulations that remain stable in physiological salt 92 concentrations is necessary i f such codelivery systems are to be optimized, which is the topic of Chapter 7 of this thesis. 93 Chapter 5 The Role of Lipid Composition, Salts and Charge in Determining Lipoplex Structure* 5.1 Introduction The purpose of this study was to determine how certain factors (helper l ipid composition, ionic environment, charge density, l i p i d : D N A ratio) affect the structure of lipoplexes. This chapter demonstrates that a variety of structures form as cationic liposomes are mixed with polynucleotides, depending on the conditions. A s cationic liposomes and D N A interact to form a complex, large increases in particle size are observed ("aggregation"). Vesicle-vesicle interactions that lead to increased particle size occur as a consequence of electrostatic and possibly hydrophobic interactions between the cationic liposomes and polynucleotides. These interactions facilitate cross-linking of vesicles between polynucleotide bridges and/or may trigger changes in the liposomal membranes that favor l ipid mixing, fusion, hemi-fusion or other forms of bilayer destabilization. Structure-activity relationships have been difficult to sort out for lipid-based D N A carriers due to the large number of variables involved and the inherent instability of the lipoplexes. Cationic liposomes and lipoplexes clearly undergo significant changes in morphology and structure. These changes are partly a consequence of polyanion induced vesicle aggregation and partly a consequence of membrane-membrane interactions upon close contact.' When assigning roles to the lipid components, therefore, it is important to consider the influences of electrostatic interactions and the presence of salts and/or serum proteins. Furthermore, aggregation-induced perturbation of membrane structure must be examined since it may result in Adapted from: Wasan, E.K. , Harvie, P., Edwards, K , Karlsson, G. and Bally, M . B . (1999) " A Multi-Step Lipid Mixing Assay to Model Structural Changes in Cationic Lipoplexes Used for In Vitro Transfection" submitted to Biochimica et Biophysica Acta-Biomembranes. 94 phase separation of lipid components or formation of non-bilayer structures (e.g. micelles or H n phase lipid) and membrane fusion. In addition, these factors are relevant for the goal of drug-DNA codelivery systems (discussed in Chapters 4 and 7), for which bilayer instability may be undesirable for maximal retention of liposomal contents. Increases in particle size can readily be monitored by light scattering techniques while membrane destabilization, which will be addressed in Chapter 6, can be examined using resonance energy transfer (RET) techniques (Struck et al, 1981). These data can be placed into context by visualization using electron microscopy (Almgren et al, 1996). Several models have been proposed for the molecular structure of complexes formed following addition of DNA to cationic liposomes, using a variety of analytical methods. These structures include smectic phase lipid and DNA sheets (Figure 5.1) (Koltover and Safinya, 1998, Radler and Safinya, Figure 5.1 A proposed model of liposome/DNA complexes - lamellar form. Adapted from Radler et al., 1997. 1997), rods of DNA wrapped in lipid ("spaghetti and meatball") (Sternberg et al, 1994), or a honeycomb-like structure (May and Ben-Shaul, 1997). The consensus at this point seems to be that flat lamellar structures are energetically most favorable, unless specific conditions favor 95 formation of hexagonal phase lipid. The latter is typical for those preparations containing the nonbilayer-forming lipid D O P E and is consistent with part of the C T E M results presented here. Cryo-transmission electron microscopy is an extremely valuable tool for the study of lipoplexes because it is minimally invasive and allows the observation of internal structure. No stains, dyes or metal shadowing are used, eliminating potential interactions with foreign materials. The molecules are frozen from the aqueous state in a form very close to their natural interactions because the rate of thermal energy transfer out of the sample is more rapid than the rate of molecular motion (Dubocet et al, 1988 ). C T E M therefore has significant advantages over other electron microscopy techniques for the visualization of lipoplex structure. 5.2 Results Lipoplexes are often prepared in a nonionic or low ionic strength solution due to the well-known tendency of these complexes to precipitate out of solution as the salt concentration is increased. This is illustrated by the data shown in Table 7 below. Mean Diameter ± SD (nm) DODAC/DOPC DODAC/DOPE NaCl (mM) liposomes lipoplexes liposomes lipoplexes 0 123±43 325±93 114±36 205±71 10 85±22 710±399 91±23 167±53 25 84±23 939±606 93±25 1000° 38 85±22 453±189 95±31 nd 6 50 86±19 555±296 1000 nd 100 85±23 704±346 nd nd 150 84±24 585±292 nd nd medium c 87±27 665±355 1000 1000 medium+serum r f 486±335 344±210 1000 228±101 Table 7 Effect of 1 MaCl on liposome size with and without added plasmid. "1000: aggregated to >1 pm diameter. *nd: not done because next-lowest NaCl concentration resulted in aggregation. cMedium was RPMI 1640. ^Medium contained 10% FBS. 96 First, cationic liposomes prepared using D O P C were stable in salt within the time they were used here (<30 min), but were >400 nm when mixed with serum-containing media. In contrast, the D O D A C / D O P E liposomes aggregated immediately when the N a C l concentration was > 50 m M . Second, addition of D N A to D O D A C / D O P E or D O D A C / D O P C liposomes resulted in aggregation, where the mean diameter as measured by Q E L S was 2- to 3-fold larger than liposomes alone. Further increases in particle size were induced following addition of salt, however, the D O D A C / D O P C lipoplexes aggregated less than the D O D A C / D O P E lipoplexes. Third, although aggregation was observed when lipoplexes were diluted into tissue culture media, the extent was reduced significantly i f the media contained 10% F B S . To further assess the effects of D N A concentration and the presence of N a C l on the mean diameter of lipoplexes, D O D A C / D O P E and D O D A C / D O P C cationic liposomes were prepared in 300 m M lactose and mixed with plasmid or oligonucleotides as described in Chapter 2. The polynucleotides were in either lactose solution or H B S . Wi th the l ipid concentration held constant at 1 m M , complexes were prepared with varying amounts of plasmid (0-125 pg/ml) or oligonucleotides (0-100 pg/ml), yielding net charge ratios (+/-) of 16 to 1.2. A t higher concentrations of polynucleotides under these conditions, visible particulates formed. The mean diameters of the liposomes and complexes were measured by Q E L S within 1 h of complex formation and the results have been summarized in Figure 5.2. The liposomes typically had a mean diameter of 120-130 nm in the absence of N a C l . A s shown in Figure 5.2, particle size increased with increasing polynucleotide concentration as charge neutrality was approached. This increase was more pronounced when the complexes were formed in the presence of H B S (triangle symbols on the graphs). The minimum diameter of liposome-nucleic acid complexes at a charge ratio of 1.6 (most commonly used for in vitro transfection in this thesis) as measured by Q E L S was approximately 300 nm. Interestingly, there was little difference between the 18mer 97 oligonucleotide (Figure 5.2B) and the 4.5 kb plasmid (Figure 5.2A) in their ability to promote vesicle aggregation as measured by light scattering techniques. For comparison, the effects of Na2HP04 (Figure 5.2C), a divalent anion, were tested. Phosphate caused little aggregation, except when added to D O D A C / D O P E liposomes prepared in lactose, where there was an increase in mean diameter as phosphate concentration increased. When D O D A C / D O P E liposomes were mixed with a concentration greater than 5 m M Na2HP04, visible particulates formed. When the D O D A C / D O P E liposomes were diluted into H B S and N a 2 H P 0 4 was immediately added, aggregation was not observed even at concentrations above 5 m M phosphate. This is in contrast to D O D A C / D O P E liposomes in H B S without phosphate present (e.g. Table 1). To facilitate the assessment of the aggregation state of multiple samples, sample turbidity was measured spectrophotometrically by measuring absorbance at 570 nm. A 96-well tissue culture plate was loaded with 50 p i of complexes prepared in triplicate at various concentrations of plasmid, oligonucleotides or phosphate. A s illustrated in Figure 5.3C, changes in turbidity were comparable to the Q E L S results shown in Figure 5.2C, where Na2HP04 was mixed with the liposomes. Turbidity can provide a rough indicator of increasing particle size, until flocculent aggregation occurs and the bulk solution clears. 98 rt 5 rt 1000 800 4 600 400 200 0 10 25 50 100 125 (16) (6.4) (3.2) (1.6) (1.2) D N A Concentration (pg/ml) (charge ratio +/-) 1000* 800 600 400 2004 B r— 0 10 25 50 100 125 (16) (6.4) (3.2) (1.6) (1.2) Oligo Concentration (pg/ml) (charge ratio +/-) 0 0.5 1 3 5 (1) (0.5) (0.17) (0.1) N a 2 H P 0 4 Concentration (mM) (charge ratio +/-) Figure 5.2 The influence of polynucleotide concentration on lipoplex size. Mean particle diameter of liposomes was measured by QELS at increasing concentrations of A : plasmid; B: 18-mer oligonucleotides; C: sodium phosphate. Symbols: • DODAC/DOPE in lactose; • DODAC/DOPC in lactose; • DODAC/DOPE in HBS; T DODAC/DOPC in HBS. Values represent mean ± intra-sample SD for individual samples. *Mean diameter >1000. o S3 © 50 75 100 DNA Concentration (pg/ml) 0.3 0.2 0.1 0.0 B 0 ' 2 5 ^ 50~^ 7 5 ^ 100 155 Oligonucleotide Concentration (pg/ml) 2 ' 3 ' 4 ' 5 N a 2 H P 0 4 Concentration (mM) Figure 5.3 The influence of polynucleotide concentration on lipoplex sample turbidity, at increasing concentrations of A: plasmid; B: 18-mer oligonucleotides and C: disodium phosphate. For charge ratios refer to Figure 5.2. Symbols: • DODAC/DOPE in lactose; • DODAC/DOPC in lactose; • DODAC/DOPE in HBS; T DODAC/DOPC in HBS. Values represent mean ± SD, n=6. 100 While determination of mean particle diameter by Q E L S and gross aggregation by turbidity give valuable information about the bulk sample, these methods tell little about the structure of the lipid/polynucleotide complexes formed. Significant morphological changes take place as the l ipid vesicles interact with polyanions. C T E M was used qualitatively to assess the morphology of the lipoplexes. In the case of D O D A C / D O P E liposomes (Figure 5.4, panel a), dense aggregates formed when mixed with either plasmid (Figure 5.4, panel b) or separately, oligonucleotides (Figure 5.4, panel c). In the case of liposome/plasmid complexes, striated regions of stacked lipid layers were observed, consistent with the distinctive "fingerprint" morphology also described in the C T E M studies of (l,2-dioleoyloxy-3-(trimethylammonium)propane ( D O T A P V D O P E lipoplexes by Gustafsson and Almgren (Gustafsson et al, 1995) and of l,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC)/3P[N-N',N'-dimethylaminoethane)-carbamoyl] cholesterol (DC-Choi) by Battersby et al (1998). The periodicity of the striations was about 6.5 nm, which is in agreement with the aforementioned studies. This indicates that it is l ikely that these are lipid bilayers with D N A sandwiched in between the layers, rather than hexagonal phase lipid. The fingerprint morphology occurred only when using cationic liposomes prepared with D O P E . Addit ion of oligonucleotides to D O D A C / D O P E liposomes in lactose resulted in the formation of dense complexes of indistinct morphology or aggregates o f liposomes (Figure 5.4, panel c). When 3 m M N a 2 H P 0 4 was added to the D O D A C / D O P E liposomes instead of polynucleotides, very large vesicles were observed (Figure 5.4, panel d). This is consistent with the data in Figure 5.2C, where an increase in mean diameter was observed in D O D A C / D O P E liposomes in lactose to which sodium phosphate was added. 101 Figure 5.4 C T E M of D O D A C / D O P E liposomes and lipoplexes in lactose. Bar indicates 100 nm. Panel a: liposomes (100 nm) alone; b: liposomes + 4.5 kb plasmid D N A , 10:1 l i p i d : D N A ratio, 100 pg D N A / m l ; c: liposomes + 18mer oligonucleotide, 10:1 l i p i d : D N A ratio, 100 pg D N A / m l ; d: liposomes + 3 m M N a 2 H P 0 4 . Notations: I: ice crystal on sample surface; P: edge of polymer scaffolding. 102 When this same set of anions was added to D O D A C / D O P E liposomes in the presence of 150 m M buffered N a C l (HBS), the instability of the bilayer structure of these liposomes was even more evident (Figure 5.5). The liposomes alone formed mostly clumps of hexagonal phase lipid (Figure 5.5, panel a). When plasmid D N A was added at the same time as H B S (i.e. prior to equilibration of the liposomes in H B S ) , no vesicles or dense aggregates were found, but rather sheets of l ipid with what appeared to be strands of radiation-sensitive material (Figure 5.5, panel b). The lipoplexes were, in general, quite sensitive to radiation damage from the electron beam of the microscope, which is noted on the electron micrographs as " R . " These sensitive areas may be D N A - r i c h regions, based on the observation that radiation sensitivity was not exhibited by any of our samples containing only lipid. It should be noted that under these conditions samples were visibly aggregated, and large aggregates greater than approximately 1 pm in diameter would be excluded from the observed C T E M sample due to limitations on the dimensions of the thin sample films used in this technique. When oligonucleotides instead of plasmid were added to the D O D A C / D O P E liposomes at the same time as H B S , the vesicles appear to be fused and aggregated, with concentric, thickened or multiple lamellae observed in some regions. (Figure 5.5, panel c). D N A in either oligonucleotide or plasmid form seems to prevent formation of NaCl-induced Hu phase formation (see Figure 5.5 panel a). When Na2HP04 was added to the liposomes instead of polynucleotides, it induced formation of highly irregular, nonvesicular aggregates, but without observable Hu phase formation (Figure 5.5, panel d). 103 a — • b Figure 5.5 C T E M of D O D A C / D O P E liposomes and lipoplexes in H B S . Bar indicates 100 nm. Panel a: liposomes (100 nm) alone; b: liposomes + 4.5 kb plasmid D N A , 10:1 l i p i d : D N A ratio, 100 pg D N A / m l ; c: liposomes + 18mer oligonucleotide, 10:1 l i p i d : D N A ratio, 100 pg D N A / m l ; d: liposomes + 3 m M N a 2 H P 0 4 . Notations: I: ice crystal on sample surface; P: edge of polymer scaffolding; R: radiation damage from the beam of the electron microscope. 104 In the case of D O D A C / D O P C liposomes in 300 m M lactose (Figure 5.6), somewhat different morphology was noted on C T E M analysis as either plasmid, oligonucleotides, or Na2HP04 were added separately. Liposomes alone, prepared by extrusion, were of the expected 100 nm diameter (Figure 5.6, panel a). The appearance of slightly flattened or invaginated vesicles, which also appeared in a few D O D A C / D O P E liposome samples, may be due to slight osmotic stress incurred during sample preparation (i.e. partial evaporation of buffer between the time the sample is blotted on the copper grid and when it is frozen in the liquid ethane). Upon addition of plasmid (Figure 5.6, panel b) or oligonucleotides (Figure 5.6, panel c), large aggregates of fused liposomes were observed and these were particularly sensitive to radiation damage. A s expected, no structures consistent with Hn-phase l ipid were observed in any of the D O D A C / D O P C samples. Non-aggregated liposomes were present in the polynucleotide-containing samples, although this was a minority population. Irregularly shaped particles > 700 nm were common for both types of polynucleotides. Concentric lamellae were observed in aggregates formed from D O D A C / D O P C liposome-oligonucleotide complexes, similar to those of D O D A C / D O P E liposome/oligonucleotide complexes (compare to Figure 5.4, panel c). Interestingly, Na2HP04 caused a reduction in the mean diameter of D O D A C / D O P C liposomes and some invagination of the vesicles (Figure 5.6, panel d). These images are consistent with the Q E L S analysis, which showed a slight reduction in vesicle mean diameter under similar conditions. N o fusion of D O D A C / D O P C into larger structures was observed when Na2HP04 was added. Figure 5.7 illustrates the changes that occur in D O D A C / D O P C liposomes prepared in lactose and diluted in H B S , and after mixing with either plasmid, oligonucleotides or Na 2 HP04. In salt, D O D A C / D O P C liposomes alone had a tendency to form complex plurilamellar structures (Figure 5.7, panel a). 105 Figure 5.6 CTEM of DODAC/DOPC liposomes and lipoplexes in lactose. Bar indicates 100 nm. Panel a: liposomes (100 nm) alone; b: liposomes + 4.5 kb plasmid DNA, 10:1 lipid:DNA ratio, 100 pg DNA/ml; c: liposomes + 18mer oligonucleotide, 10:1 lipid:DNA ratio, 100 pg DNA/ml; d: liposomes + 3 mM Na2HP04. Notations: I: ice crystal on sample surface; P: edge of polymer scaffolding; R: radiation damage from the beam of the electron microscope. 106 Figure 5.7 C T E M of D O D A C / D O P C liposomes and lipoplexes in H B S . Panel a: liposomes (100 nm) alone; b: liposomes + 4.5 kb plasmid D N A , 10:1 l i p i d : D N A ratio, 100 pg D N A / m l ; c: liposomes + 18mer oligonucleotide, 10:1 l i p i d : D N A ratio, 100 pg D N A / m l ; d: liposomes + 3 m M N a 2 H P 0 4 . Notations: I: ice crystal on sample surface; P: edge of polymer scaffolding. 107 Addition of plasmid D N A and H B S to the D O D A C / D O P C liposomes produced loose clumps of large vesicles and structurally amorphous membrane structures, suggesting that membrane fusion is occurring (Figure 5.7, panel b). Addit ion of oligonucleotides in H B S generated very large aggregates that appear to be denser than those formed upon addition of plasmid D N A . Similarly to D O D A C / D O P E liposomes in H B S (Figure 5.5, panel c) and D O D A C / D O P C liposomes in lactose (Figure 5.6, panel c), addition of oligonucleotides results in structurally complex regions containing concentric bilayers (Figure 5.7, panel c). Na2HP04 had no apparent effect on D O D A C / D O P C liposomes in H B S , which appeared as unfused vesicles (Figure 5.7, panel d). 5.3 Discussion This investigation began with the study of the well-known role of salt in promoting aggregation of the lipoplexes composed of either D O D A C / D O P E or D O D A C / D O P C bound to plasmid D N A . Table 7 and Figure 5.2 show the extent of salt-induced aggregation measured by Q E L S , which is marked for DOPE-lipoplexes. Larger structures with an increased degree of lipid-lipid interactions formed as the l i p i d : D N A ratio approached a charge-neutral ratio, suggesting that electrostatic interactions play the predominant role in complex formation. The effect of the presence of salts was to promote particle association rather than to reduce it by surface charge shielding. Several mechanisms may be at work in this situation. Initially C l " binding to the liposomes may reduce liposome-liposome charge repulsion by neutralizing surface charge. Salt bridges may form between vesicles, and structured water at the surface may be disrupted leading to dehydration. D O P E can then revert to Ho phase and lipid mixing or membrane fusion may occur. However, when polynucleotides are added, the ionic interaction between the anionic polymer ( D N A ) and the cationic polymer (liposome membranes) would be of high affinity and cooperative in nature. (These parameters were not measured experimentally 108 in this work.) Water may be excluded as lipids and D N A bind, forming a "self-assembling," condensed structure. The presence of salt ions may prevent full dehydration and condensation, however, leading to the highly irregular, polymorphic structures seen in the C T E M studies in this chapter (Fig. 5.5 and 5.7). When excess liposomes were present (large net positive charge ratio) lipoplex mean diameter was smaller (Fig. 5.2). This may be due to charge repulsion between lipoplexes with a net positive charge. If some lipoplexes were continuing to assemble in the presence of excess free liposomes, the Q E L S analysis would have shown a markedly larger SD and x2 value for the Gaussian distribution, indicating the presence o f several subpopulations. This was not observed. Large aggregates can still assemble when the net charge ratio is slightly positive, however (CR = 1.6 to 3, for example, which is the range typically used for in vitro transfection formulations). This may indicate that not all the positive charge from the liposomes is surface-available during/after lipoplex formation. Based on this observation, and that of the residual trapped volume after lipoplex formation (Chapter 4) and C T E M studies here, there is likely a partial retention of bilayer structure in the lipoplexes at slightly postive charge ratios. Alternatively, the reorganization of the lipids after lipoplex assembly causes the cationic lipid to be sequestered in the lipoplex interior (cationic l ipid domain formation), albeit limited by l ipid mobility and packing constraints. These issues are discussed further in Chapter 6. Charge density of the added component was also important, however. A comparison was made of the micro structures of liposomes interacting with double-stranded plasmid D N A (Figures 5.4-5.7, panel b of each), 18mer phosphorothioate oligonucleotides (Figures 5.4-5.7, panel c of each) or sodium phosphate (Figures 5.4-5.7, panel d of each). The^extent of disruption of liposome structure decreased (as visualized on C T E M ) as the molecular weight or charge density of the added component decreased, reflective of the cooperative nature of the lipoplex assembly process. 109 The C T E M data suggest that D O D A C / D O P E liposomes can assume structures that are dramatically different depending on whether salts are present (compare Figures 5.4 and 5.5, panel b of both). It may also be speculated that polyanion binding may cause disruption of the membrane leading to formation of monomers or micelles that are capable of binding to D N A , eventually leading to formation of lipoplexes (Figure 5.8). Tubular coated DNA Figure 5.8 Schematic of a theoretical mechanism to account for the diverse morphology of lipoplexes, which depends on lipid composition, mixing procedures and environmental conditions proceeding via unknown intermediates. Artwork by Ashley M . Bradford. 110 Janoff et al. have shown evidence that in the transition from lamellar to hexagonal phase lipid for DOPE-containing l ipid mixtures, that monomeric lipids may be released (Perkins et al, 1996). It is possible then that micelles can form in an intermediate step during such transitions, although experimental evidence for this scenario was not generated in this study. However, this could be the basis for the major rearrangement of lipids upon interaction of DOPE-containing cationic liposomes with D N A . Released monomeric l ipid may or may not form micelles, then bind to D N A , leading to the formation of polymorphic nonliposomal structures such as the membrane sheets, tubular lipid-coated D N A or the fingerprint morphology (Figure 5.8). The major limitation of the Q E L S technique and C T E M analysis studies is that as the particle size approaches 1 pm, results are either not reliable or not measurable. If a majority of the lipoplexes is involved in superaggregates (>1 pm), only the fraction of the population still homogeneously dispersed in the suspension is being measured when one uses these techniques. It is important to note, however, that the smaller-sized population is representative, because the particles start smaller then continue to assemble by the same process as discussed above. Particle size growth is rapid initially (within seconds or minutes) followed by a much slower phase of growth over hours or days (Zuidam and Barenholz, 1998). A t high or low charge ratios, less aggregation is noted (see Chapter 4), presumably due to charge repulsion or that the net charge ratio does not reflect the surface-available charge due to l ipid domain formation. After aggregation has occurred the slower phase presumably is due to the reduction in the collision rate as the total number of particles in suspension decreases. The C T E M studies summarized here (see Figures 5.4-5.7) demonstrated that there is a great heterogeneity and subtlety of morphology that cannot be expressed by simply reporting mean diameters o f the lipoplexes. The concepts of local surface charge, l ipid domain formation, lipid packing constraints, D N A accessibility, and the dynamic flexibility of the supramolecular complex are likely to be just as critical as the overall charge ratio and the size of the aggregate. i l l Work in this field must succeed in controlling the substructure of l i p i d - D N A formulations in order for these factors to be studied in greater detail. 112 Chapter 6 Lipid Mixing Behavior of Cationic Liposomes and Lipoplexes and Its Relationship to in Vitro Transfection* 6.1 Introduction M u c h effort has been devoted in the past decade to studying the mechanism of action and intracellular behavior of lipoplexes during transfection in vitro or in vivo (Friend et al., 1996, Thierry et al., 1997). In addition, investigators have attempted to correlate how the physical properties of the lipids, liposomes and lipoplexes influence biological activity (Bennett et al., 1997, Balasubramaniam et al, 1996, Feigner et al, 1994). The relative importance of particular physicochemical properties of lipids and lipid-lipid interactions are still not fully elucidated in this context. Formation of lipoplexes involves electrostatic interactions, which may or may not involve complete intermixing of liposomal bilayers. Membrane fusion is a specific rearrangement event occurring after aggregation (contact) and l ipid mixing (an intermediate step) resulting in the formation of one new bilayer where there once were two. The ability of the lipoplexes to transfer D N A into cells is dependent on the physicochemical attributes of the lipoplexes including l ipid mixing ability; therefore, characterization of binding-induced changes in liposomes is critical for the development of lipid-based D N A delivery systems. To clarify the apparent lack of correlation between lipid mixing of cationic liposomes and in vitro transfection previously observed, a multi-step l ipid mixing assay was developed to model the sequential steps involved in transfection. The roles of anion charge density, charge ratio and presence of salt on l ipid mixing and liposome aggregation were also investigated. The fluorescence resonance-energy transfer method (RET) was used to monitor l ipid mixing as cationic liposomes ( D O D A C / D O P E and D O D A C / D O P C , 50/50 mol%) were combined with * Adapted from : Wasan, E.K., Harvie, P., Edwards, K. , Karlsson, G. and Bally, M . B . (1999) " A Multi-Step Lipid Mixing Assay to Model Structural Changes in Cationic Lipoplexes Used for In Vitro Transfection" Biochimica et Biophysica Acta - Biomembranes, in press. 113 plasmid, 18mer oligonucleotides or NaiHPCv. Transfection studies were performed to determine what effect the presence of salt during complexation had on in vitro transfection with the lipoplexes. These data give new information about the effects o f polynucleotide binding to cationic liposomes, illustrating the complicated nature of anion induced changes in liposome morphology and membrane behavior. The importance of controlling the binding reactions for the preparation of transfection-active, structurally defined lipid-based polynucleotide delivery systems is discussed. This is especially important for the development of lipoplexes that can retain aqueous contents for codelivery applications, as introduced in Chapter 4. To illustrate the problems in assigning mechanisms of activity of l ipid components it is useful to review the roles that have been assigned to the helper l ipid D O P E . D O P E forms nonbilayer structures involved in the fusion of membranes (Siegel, 1986) and it is to this property that transfection-enhancing abilities are usually attributed (Farhood et al, 1995, Hu i et al, 1996). It has been demonstrated, however, that efficient transfection can be achieved using liposomes that do not incorporate non-bilayer forming lipids, such as with DOTAP/cholesterol liposomes (Templeton et al, 1997, Crook et al, 1998). In addition, it may be suggested that the amine group of phosphatidylethanolamine (PE) can interact with the D N A phosphate groups and that such an interaction serves to weaken the binding reaction between cationic lipids and D N A . This, in turn, may promote the eventual disassembly of the lipoplex, a process that has been defined as a critical step required for transgene expression (Wong et al, 1996, X u and Szoka, 1996). It can also be predicted, however, that PE-containing lipoplexes w i l l be more susceptible to factors that promote disassembly prior to accessing a target cell population. This is consistent with results demonstrating that PE-containing lipoplexes afford less protection of plasmid against nucleases then those prepared with phosphatidylcholine (Harvie et al, 1998). 114 The resonance energy transfer technique (RET) has proven to be particularly useful in assessing polynucleotide induced membrane destabilization. It is useful to note, however, that this assay was developed to study lipid mixing mediated by addition of cations to anionic membranes. Various studies of anionic liposome lipid mixing, following addition of monovalent or divalent cations (Bangham and Papahadjopoulos, 1966, Papahadjopoulos and Poste, 1975), polycations (Ash et al., 1978, Oku et al., 1986) and cationic peptides (Pecheur et al., 1998, Rapaport et al., 1994), have formed a basis for studies assessing polynucleotide binding to cationic liposomes (Stamatatos et al., 1988, Bailey and Cull is , 1997). There are similarities in the two systems because electrostatic interactions are the primary driving forces promoting adhesion of the liposomes. Perhaps the most important lesson that can be learned when reviewing these previous studies is that it is difficult to control l ipid mixing reactions between liposomes. This, in turn, makes it difficult to form homogenous membrane structures after induction of l ipid mixing. In this chapter, the effects of plasmid D N A , oligonucleotides and phosphate on cationic liposome aggregation and membrane destabilization are described. Unlike previous studies, however, liposome and lipoplex behavior was examined sequentially as the lipoplexes were formed, diluted into salt-containing buffers and eventually added to anionic vesicles (model cell membranes). While the interaction of cationic liposomes with anionic vesicles or biological membranes has been investigated before (Bailey and Cull is , 1997, Stamatatos et al., 1988), this was the first study examining the l ipid mixing behavior of lipoplexes in a multi-step manner that represents a transfection-relevant scenario. It was observed that the potential for cationic liposomes to undergo lipid mixing reactions generally decreased sequentially following addition of polynucleotides and salts, such that much of the original "l ipid mixing potential" of the 115 cationic liposomes was reduced by the time they come into contact with the model cell membranes. 6.2 Results To gain a further understanding of the vesicle-vesicle interactions occurring during the process of lipoplex formation, l ipid mixing assays were performed using the R E T technique. A s summarized Figure 6.1, the l ipid mixing data suggest that cationic liposomes behaved quite differently depending on the polyanion added. Following addition of plasmid D N A to 1 m M D O D A C / D O P E liposomes in lactose (Figure 6.1 A , squares) there was a progressive increase in lipid mixing as C R (+/-) decreased from 16 to 1.2 with a maximum (%maxLM) of approximately 65% at 75 pg D N A / m l (CR (+/-) = 2.4). D O D A C / D O P C liposomes under the same conditions (Figure 6.1 A , circles) showed a similar pattern, however, % m a x L M only reached 20%, also when C R = 2.4. In the presence of H B S , the % m a x L M observed (20-25%) for D O D A C / D O P E liposomes was comparable to D O D A C / D O P C liposomes in lactose. It should be noted that this value was significantly less then the 60% observed in the absence of H B S . Salt-mediated reductions in l ipid mixing were also evident for the D O D A C / D O P C liposomes in H B S where less than 10% m a x L M was observed. Phosphorothioate oligonucleotides (18mer, 0-100 pg/ml) were also studied for their ability to promote lipid mixing of the cationic liposomes (Figure 6. IB) . When liposomes were in lactose, addition of increasing concentrations of oligonucleotides yielded a pattern of increasing % m a x L M as charge neutrality was approached. In lactose, D O D A C / D O P E vesicles (Figure 6.IB, squares) demonstrated 27% of maximal lipid mixing and this was observed following addition of 100 pg oligonucleotide/ml (CR = 1.6). Oligonucleotide-induced lipid mixing was similar to results obtained for the D O D A C / D O P C (Figure 6 . IB, circles) and D O D A C / D O P E liposomes. In H B S , maximal lipid mixing was observed at high C R (10-25 pg oligos/ml). 116 s S T" ~ 1 1 100 1 1 125 (1.6) (1.2) 45^  B DNA Concentration (u.g/ml) (charge ratio +/-) 0 10 25 50 7 5 100 (16) (6.4) (3.2) (2.4) (1.6) O l i g o C o n c e n t r a t i o n (jxg/mi) (charge ratio +/-) 0.5 1 3 5 (1) (0-5) (0.17) (0.1) Na 2 HP0 4 Concentration (mM) (charge ratio +/-) Figure 6.1 Lipid mixing of liposomes induced by polynucleotides or phosphate. Lipid mixing of liposomes as a percentage of maximum, measured by resonance energy transfer at increasing concentrations of A : plasmid; B: 18-mer oligonucleotides; C: sodium phosphate. Numbers in parentheses refer to charge ratio (+/-). Lipid concentration was 1 mM. Symbols: • DODAC/DOPE in lactose; • DODAC/DOPC in lactose; • DODAC/DOPE in HBS; T DODAC/DOPC in HBS. Values represent mean ± SD, n=3. 117 The extent of l ipid mixing appeared to decrease significantly at concentrations >25 pg/ml for both D O D A C / D O P E (Figure 6. I B , up triangles) and D O D A C / D O P C liposomes (Figure 6. I B , down triangles). The apparent decrease in lipid mixing in H B S at oligonucleotide concentrations > 25 pg/ml may have been due to flocculent aggregation. The l ipid mixing assay, like the turbidity assay in Figure 5.3 of the previous chapter, does not provide quantitative data under these conditions, illustrating an important major limitation of the R E T technique for these kinds of samples. When these studies were conducted with sodium phosphate (0-10 m M ) minimal lipid mixing was observed at up to 5 m M N a 2 H P 0 4 (CR = 0.1, Figure 6.1C). The m a x L M was <15% for D O D A C / D O P E liposomes in lactose following addition of N a 2 H P 0 4 to a final concentration of 3-5 m M (Figure 6.1C, squares). When D O D A C / D O P C liposomes in lactose were mixed with phosphate, there was a consistent decrease N B D - P E fluorescence as phosphate concentration increased, shown in the decrease in % lipid mixing from baseline (Figure 6.1C, circles). This decrease is consistent with the shrinking of the liposomes, or possibly blebbing/budding of new, smaller vesicles as observed by C T E M in Chapter 5 (Figure 5.6, panel d) and the reduction in mean diameter observed by Q E L S (Figure 5.2C). It may be speculated that the binding of the phosphate ions to the cationic liposomes induced lipid packing changes (perhaps to the extent that new vesicles pinched off), resulting in vesicles in which the distance between N B D - P E and rhodamine-PE fluorescent lipids was decreased, causing an increase in fluorescence resonance energy transfer (reduced N B D emission signal). The primary interest in these cationic liposomes is in their use as D N A transfer vehicles for gene therapy applications. Therefore, the relationship between the physical properties and biological behavior of cationic liposomes complexed to plasmid expression vectors is of primary interest. It is clear that addition of oligonucleotides vs. plasmid D N A may result in lipoplexes 118 with structural features that are different from each other. This should be taken into consideration when lipid-based carriers for antisense molecules are being developed based on information about carriers for plasmid D N A . The remainder of this study, however, was focused only on the effects of plasmid D N A on liposome behavior and the effects o f the liposomes on reporter gene expression in vitro. A s mentioned above, an important factor in successful transfection using cationic lipoplexes is the ability to interact with cellular membranes, which is thought to require particles that readily undergo lipid mixing. However, aggregation of cationic liposomes (see Figures 5.2 and 5.3) does not necessarily mean that complete vesicle disruption has occurred, as indicated in the C T E M images of Figures 5.4-5.7 of the previous chapter and the lipid mixing assays summarized in Figure 6.1. Likewise, l ipid mixing during lipoplex formation in simple buffer solution does not necessarily indicate the l ipid mixing potential of the lipoplexes when they reach the target cell membrane. It is believed that the liposomes continue to undergo major structural changes at each phase of the process of in vitro transfection, namely during: 1) interactions with anions in the buffer solution used to prepare or dilute the liposomes; 2) formation of the lipoplex with plasmid; 3) dilution into standard tissue culture medium which may or may not contain serum; and 4) interactions with cellular membranes as the lipoplexes reach the cells to be transfected. Each of these steps w i l l necessarily affect the outcome of subsequent interactions, and examining any one step in isolation gives an incomplete picture. To examine this process as a continuum rather than as isolated steps, a multi-step lipid mixing assay was designed, based on the R E T technique, to model in vitro transfection. D O D A C / D O P E and D O D A C / D O P C lipoplexes were prepared at 10:1 l i p i d : D N A ratio (nmoles:pg, CR=1.6), as previously described. Fluorescence changes were monitored over time throughout the processes of lipoplex formation in lactose or in H B S , dilution into tissue culture 119 medium with or without 10% F B S as well as following addition of D O P S / D O P C anionic liposomes. A t the end of the time course 0.3%> Triton X-100 was added to achieve maximal lipid mixing (Figure 6.2). The addition of anionic liposomes was designed to model contact with target cell plasma membranes. It is also similar to previous studies in which destabilization of lipoplexes was induced by anionic vesicles (Xu and Szoka, 1996, Harvie et al., 1998). Disassembly of the lipoplex at the cellular level to allow passage of free plasmid into the nucleus is presumably a requirement for transfection-competent lipoplexes (Thierry et al., 1997). There are two major points to be made concerning the results of the l ipid mixing assay. First, in the absence of F B S (Figure 6.3 A ) , both helper lipid composition and buffer composition affected l ipid mixing. The structures generated at each step were capable of undergoing reactions that resulted in further dilution of the fluorescent lipids. Second, F B S had a measurable effect on the lipid mixing (Figure 6.2B). For example, in the absence of F B S (Figure 6.2A) increases in percent maximal lipid mixing were observed in step increments for both D O D A C / D O P E and D O D A C / D O P C lipoplexes. A l l samples exhibited approximately 10% m a x L M at the point of D N A addition (200 s). However, there was a great disparity between samples prepared with different lipid compositions or buffers when the tissue culture medium was added (at 400 s). D O D A C / D O P C showed little or no increase in lipid mixing, but D O D A C / D O P E lipoplexes reached approximately 25%> when prepared in lactose and 38%> when prepared in H B S . Addition of anionic liposomes (at 600 s) induced further l ipid mixing except in the case of D O D A C / D O P C lipoplexes prepared in H B S . Interestingly, the largest change in %>maxLM at this step occurred in D O D A C / D O P E lipoplexes prepared in lactose, which reached almost 55%. At all steps, DOPC-containing lipoplexes demonstrated less ability to undergo lipid mixing than D O P E -containing lipoplexes. 120 60 -DODAC/DOPE in lactose DODAC/DOPE in HBS -DODAC/DOPC in lactose DODAC/DOPC in HBS 400 600 Time (sec) Figure 6.2 Representative multi-step lipid mixing assay using RET. Sequentially, at 200 sec, D N A was added to liposomes; at 400 sec, tissue culture medium was added; at 600 sec, anionic liposomes (DOPS/DOPE) were added. A : assay performed using serum-free medium; B: medium containing 10% FBS was used. See text for explanation of calculation of percent maximal lipid mixing. 121 When the tissue culture medium introduced at 400 s contained 10% F B S (Figure 6.2B), little or no additional l ipid mixing occurred, except for D O D A C / D O P C lipoplexes prepared in lactose, which went from approximately 5% to 10% m a x L M at this step. The initially higher level of lipid mixing observed upon addition of D N A to the liposomes (at 200 s) in Figure 6.2B compared to Figure 6.2A reflects the variability inherent in lipoplex formation. It should be noted that the trend of DOPE-containing lipoplexes demonstrating greater l ipid mixing ability than DOPC-containing lipoplexes was highly consistent. The subsequent addition of anionic membranes (at 600 s) to the lipoplexes diluted into serum-containing media showed that the membrane structures formed were not capable of undergoing reactions that would cause additional l ipid mixing. These data support the idea that serum is inhibitory for anionic membrane-induced destabilization of the D O D A C / D O P E lipoplexes (Harvie et al, 1998). D O D A C / D O P C lipoplexes prepared in lactose were the exception, rising from 10% to 2 0 % m a x L M at this step, although it is not clear why this should be the case. For D O D A C / D O P C lipoplexes prepared in H B S , l ipid mixing was eliminated after addition of medium with or without F B S (Figure 6.2A and 6.2B, at 400 s). The observed reductions in % m a x L M ( D O D A C / D O P E in H B S , D O D A C / D O P C in H B S ) could be due to changes in membrane organization induced by anionic protein binding after addition of serum-containing media, although protein binding was not evaluated in this study. To summarize the results of the multi-step l ipid mixing assay: although lipid mixing was quite similar between liposomes of differing lipid compositions or buffers at the point of D N A addition, considerable differences arose at the point of addition of tissue culture medium. DOPE-containing lipoplexes had a greater l ipid mixing ability than DOPC-containing lipoplexes, especially at the point of adding anionic liposomes (model cell membranes). F B S in the media reduced both overall l ipid mixing and the differences due to l ipid composition. To determine i f 122 there was a correlation between lipid mixing and successful transfection, transfection of B16 /BL6 mouse melanoma cells was done in vitro using D O D A C / D O P E and D O D A C / D O P C lipoplexes, prepared in either lactose or in H B S (Figure 6.3). DNA DODAC/ DOPE DODAC/ DOPC B b,c a,b' DNA B,— DODAC/ DOPE b,c DODAC/ DOPC Figure 6.3 In vitro transfection of B16 cells for comparison to lipoplex lipid mixing properties. A: Expression of chloramphenicol acetyltransferase gene transfected into B16/BL6 murine melanoma cells using lipoplexes (CR (+/-) = 1.62) prepared in either 300 mM lactose or 150 mM buffered NaCl (HBS). Black bars: samples prepared in lactose; hatched bars: samples prepared in HBS. Data represent the mean ± SD, n=6. "Significantly different from free DNA. ^Significantly different from DODAC/DOPC in lactose. "Significantly different from DODAC/DOPE in lactose. ^Significantly different from DODAC/DOPE in HBS. (p<0.05) B: Expression of luciferase gene transfected into B16/BL6 murine melanoma cells in medium with or without 10% FBS using cationic liposome/DNA complexes prepared in either 300 mM lactose or HBS. Bars (diluent groups): black- in lactose, with serum; grey- in lactose, no serum; hatched-in HBS, with serum; open- in HBS, no serum. Data represent mean ± SD, n=6. "Significantly different from DODAC/DOPE for this diluent group; ^significantly different from lactose with serum for this lipid composition; "significantly different from HBS with serum for this lipid composition. (p<0.05) 'Significance refers to all 4 diluent groups for DNA alone. The first experiment used a plasmid containing a reporter gene encoding chloramphenicol acetyltransferase ( C A T ) . Transfection with DOPE-lipoplexes prepared in lactose resulted in 16 ± 1.7 m U C A T activity/pg protein and DOPC-lipoplexes yielded 8.5 ± 2 . 1 mU/ug. Lipoplexes prepared in H B S resulted in 13.8 ± 2.6 and 9.1 ± 1.1 mU/pg for D O P E - and DOPC-lipoplexes, respectively (Figure 6.3A). No significant toxicity was noted by visual inspection or in the measurement o f total cellular protein concentrations after transfection compared to untransfected controls. In the second experiment, samples were prepared in the same way, as described in the Methods, but using a plasmid encoding the luciferase reporter gene using the same vector backbone and promoter as the CAT-encoding plasmid. Lipoplexes were made in either 300 m M 123 lactose or H B S . Luciferase assay results, as shown in Figure 6.3B, indicate there was a significant difference between cells transfected with D O D A C / D O P E vs. D O D A C / D O P C lipoplexes except when they were prepared in lactose and diluted into medium containing serum. A l l groups transfected with D O D A C / D O P C lipoplexes showed a significantly lower luciferase activity than groups transfected with D O D A C / D O P E lipoplexes. The luciferase activity of groups receiving D O D A C / D O P C lipoplexes was not significantly different from those receiving plasmid D N A alone, except for the significantly greater activity when they were prepared in lactose and diluted into serum-free medium. F B S reduced transfection ability of all lipoplexes. A s in the first experiment, surprisingly, the presence of H B S during lipoplex formation did not have a major effect on transfection. These data are consistent with other reports suggesting that there is often only 5 to 10-fold difference in transfection observed when comparing lipoplexes prepared with D O P C versus D O P E (Farhood et al., 1995, Hu i et al., 1996). These differences, however, were dependent on the plasmid expression vector used. The increased transfection that was observed for the D O P E -lipoplexes transfected with the C A T plasmid compared to the luciferase plasmid was striking. In comparing the results of the multi-step l ipid mixing assay (Figure 6.2) and the transfection studies (Figure 6.3) it is clear that for successful transfection, the lipoplexes do not necessarily have to be capable of supporting the extensive l ipid mixing that can be measured through a R E T assay. There is a tendency to see improved transfection when using lipoplexes that do undergo lipid mixing following addition of anionic membranes, however, the improvements are typically no more then 2- to 5-fold and may depend on the plasmid vector being used. In comparing Figures 6.3A and 6.3B, for example, DOPC-lipoplexes promoted transfection nearly as well as DOPE-lipoplexes when the C A T plasmid was used, but not when the luciferase plasmid was used. While the two plasmid preparations were isolated and purified 124 by the same methods, it is unclear i f this difference in expression was due to some physical factor involving the plasmid preparation. Alternatively, it may have been due to a difference either in the intracellular processing of lipoplexes or the expression of the respective reporter genes in this cell type. 6.3 Discussion The purpose of this study was to determine how certain factors (helper lipid composition, ionic environment, charge density, l i p i d : D N A ratio) affect the l ipid mixing ability of lipoplexes and subsequently their in vitro transfection ability. Structure-activity relationships have been difficult to sort out for lipid-based D N A carriers due to the large number of variables involved and the inherent instability of the lipoplexes. It was demonstrated that a variety of structures form as cationic liposomes are mixed with polynucleotides (see Chapter 5), depending on the conditions, and that they differ in their l ipid mixing capacity. The differences in transfection ability, however, are surprisingly small (2-3 fold) despite this heterogeneity. This implies that there may be either enough similarity in the microstructure to render the larger-scale morphological differences less important, or that there are multiple routes of uptake possible, some of which are not fusion-dependent. A third possibility, not addressed here, is that plasmid-dependent factors were the most important. Currently there is much discussion regarding how the "l ipid mixing potential" of lipoplexes is important for transfection (Stegmann, 1987, Bailey and Cull is , 1997, M o k and Cullis , 1997, Eastman et al., 1997, L i and Hui , 1997). It is believed by many that the optimal interaction of lipoplexes with the cellular plasma and endosomal membranes depends on the ability of the lipids to form nonbilayer structures that allow membrane fusion. The fundamental assumption is that a certain minimal degree of lipid mixing character is mandatory for destabilization/interaction with cellular membranes. This is necessary so that the lipoplex can 125 enter the cell, be released from endosomes or lysosomes, and for the release of the associated D N A . Release of the D N A has been shown necessary for nuclear uptake and subsequent transcription of the exogenous D N A (Zabner et al, 1995). It may be overly simplistic to expect a direct correlation between lipid mixing of liposomes alone in a low ionic strength solution and transfection of cells due to the many changes that the liposomes/lipoplexes undergo during the process. In an actual transfection procedure, generally liposomes are pre-formed, then the liposomes are incubated with D N A to form lipoplexes and finally the lipoplexes are administered to cells in tissue culture or to an animal. Each step w i l l have an impact on subsequent events. It is therefore important to think about each step separately before considering subsequent changes. First, the liposomes must interact with the plasmid, changing the liposome conformation and its net surface charge density. This step w i l l be influenced by the ionic strength during lipoplex formation (Chapter 5, Table 7). Changes also occur in the plasmid itself, often described as D N A "condensation," which in this context is often assessed indirectly by measuring the relative degree of exclusion of DNA-intercalating dyes, such as ethidium bromide (Xu and Szoka, 1996, Eastman et al, 1997) or TO-PRO-1 (Reimer et al, 1995). A decrease in sensitivity of the plasmid to degradation by DNase I is another feature of this change. (DNase I sensitivity experiments are discussed in Chapters 2, 3 and 7 of this thesis.) Second, the lipoplex may incur further structural and electrostatic surface changes when it encounters tissue culture medium (or in vivo, the blood stream or tissue fluids). These media are highly complex, containing various electrolytes, proteins and carbohydrates. The R P M I 1640 medium that was used in this study, for example, contains calcium nitrate, magnesium sulfate, potassium chloride, sodium chloride, sodium phosphate, the essential amino acids, vitamins, glucose, sodium bicarbonate, etc. It is reasonable to expect an interaction of these components 126 with the lipoplexes, which may affect their "lipid mixing abilities," or perhaps affect l ipid organization within the lipoplex. Third, the lipoplexes must interact with cellular membranes, both on the exterior and the interior of the target cell, which also contain membrane proteins and carbohydrates in addition to charged and uncharged phospholipids. This interaction is likely somewhat different from the interaction of cationic liposomes and anionic liposomes in a low ionic strength environment in the absence of serum proteins. Also at this step, the D N A may regain its sensitivity to DNase (Harvie et al, 1998) and its ability to bind D N A intercalating dyes ( X u and Szoka, 1996). These characteristics indicate an increase in D N A accessibility, although not necessarily complete release of the D N A (Wong et al., 1999). The major advantage of the multi-step lipid mixing assay used here is the ability to monitor the sequential changes in l ipid mixing that occur in the lipoplexes during the steps analogous to those involved in cellular transfection. This multi-step assay may be more appropriate for trying to correlate l ipid mixing with transfection ability than previous studies on this topic. Analysis of the electrostatic interactions between cationic liposomes and polynucleic acid and how they determine the structure of lipoplexes typically used for gene transfer has recently been addressed (Radler and Safinya, 1997, Zuidam and Barenholz, 1998, Bruinsma and Mashl, 1998, Dan, 1998, Koltover and Safinya, 1998). The complexation of lipids and D N A is well described by existing polyelectrolyte theory, specifically Manning condensation (Manning, 1978, Manning and Mohanty, 1997). In polyelectrolyte interactions, condensation of the polymer is dependent on the linear charge density of the oppositely charged ions. Here, the lipid bilayer and/or D N A molecules are the charged polymer being condensed. Cationic lipids or salts condense the D N A , while polynucleotides condense/rearrange the cationic l ipid bilayer. Crosslinking of liposomes by ions of opposite charge (salt bridging) can condense the lipids 127 through dehydration of the l ipid bilayer as the counterions bind to the charged l ipid headroups. When certain lipids are present in the lipid bilayers, such as phosphatidylethanolamine, this may result in the formation of nonbilayer structures followed by membrane fusion. This interaction has been addressed extensively using monovalent and divalent cations to induce fusion of anionic liposomes (Diizgiines et al., 1981a,b, Wilschut et al., 1985). Divalent cations cause very rapid fusion of anionic vesicles (on the order of 1 min), and the percent maximal l ipid mixing achieved depends on the cation/anion ratio, salt concentration, temperature, etc. The effects of multivalent synthetic cations on anionic liposome behavior have also been addressed. Oku and coworkers demonstrated that polymeric cations induce maximal anionic liposome fusion (by contents release, l ipid mixing assays and turbidity changes in PS /PC liposomes) when the C R = 1 (Oku et al., 1986). Divalent cations such as C a 2 + and M g 2 + require much higher concentrations relative to the anionic vesicles to achieve the same degree of liposome fusion as charged polymers. This is analogous to the l ipid mixing data presented here, in which sodium phosphate did not produce the same degree of percent maximal lipid mixing in the cationic liposomes compared to the polynucleotides, even when the apparent net C R ~ 1. L ip id mixing was also reduced in the presence of salt. Thus, charge density is important for induction of l ipid mixing of cationic liposomes by anions, similar to the results of previous studies investigating the effect of cations on anionic liposomes. Polynucleotides also bind to cationic liposomes very rapidly and efficiently (Zuidam and Barenholz, 1998). It has been shown by several methods in this thesis that the degrees of structural change induced by double-stranded plasmid, single-stranded oligonucleotides and sodium phosphate are different, as expected. When Na2HP04 was used as the anion instead of polynucleotides in this investigation, formation of dense complexes was much less pronounced, in agreement with studies by M i t a et al. (1977), which showed that a certain critical length o f charge is necessary to induce the formation o f condensed DNA-cat ion 128 complexes. Here, the critical length is somewhere in between a single phosphate divalent anion and an 18mer polynucleotide. These results also are supported in studies by L i u et al. (1998), which showed that the binding efficiency of polyphosphates to a cationic surfactant depends on the degree of polyphosphate polymerization. Thus it is becoming clear that the reactions between cationic liposomes and D N A can be understood based on previous work with anionic liposomes and cations, bearing in mind that both the spacing of the charges and the length of the opposing charged molecules are critical. This may explain in part why the cationic l ipid headgroup charge is important in lipoplex formation (Wong et al., 1996) and in transfection with lipoplexes (Byk et al., 1998). Controlling lipid interactions to generate defined structures from plasmid and pre-formed liposomes, however, may prove to be considerably more problematic. For example, much effort has been spent studying anionic model membrane fusion in an attempt to generate vesicles that w i l l fuse in a controlled manner, for example, without loss o f contents. There has been some degree of success in regulating the rate of fusion of anionic liposomes by modifying the l ipid composition, altering the temperature of the system, and/or through the use of pH-sensitive fusion-active lipids or peptides (Eidelman et al, 1984, Conner et al., 1984, Diizgiines et al, 1984). Having the ability to control the kinetics of lipid mixing, however, does not mean one w i l l be able to control the final structure of lipoplexes. Intermediate events in the fusion process are fleeting and unstable, such as lipidic particles imaged during freeze-fracture electron microscopy (Verkleij et al., 1980). Other possible intermediate structures, such as inverted micellar intermediates (Evils) (Siegal, 1996), are considered theoretical, or their lack of capture by present microscopy techniques is attributed to their ephemeral nature. If intermediates are difficult to study experimentally and the only aspect of l ipid mixing that can be readily controlled is the rate, then it is unlikely that lipoplexes made from pre-formed DOPE-containing liposomes and plasmid w i l l ever be 129 developed beyond the present stage of heterogeneous, unstable lipoplexes with variable transfection efficiency. Chapter 7 addresses the need to redesign the liposome formulation to increase its stability upon D N A binding, and proposes several effective new formulations. Alternative methods of forming lipid-based D N A carriers in a more controlled manner are currently under development (Zhang et al., 1996). A major focus of this chapter is the effect of l ipid composition and ionic environment on liposomal l ipid mixing and its relationship to in vitro transfection. It was observed in the multi-step lipid mixing assay (Figure 6.2) that there was not a large difference between D O D A C / D O P E and D O D A C / D O P C lipoplexes in l ipid mixing ability in the presence of serum. This was also borne out in the transfection experiments done in the presence of 10% F B S , where the difference between treatment groups was small (Figure 6.3). In the absence of serum, there was a clear difference in the ability of P E - vs. PC-containing lipoplexes to undergo l ipid mixing at the point o f addition of anionic liposomes. In the transfection experiment using the luciferase reporter gene, the PE-containing lipoplexes maintained their ability to transfect cells, even when prepared in salt and when the transfection was done in serum-containing media. D O D A C / D O P C lipoplexes did not maintain lipid mixing ability as well as DOPE-containing lipoplexes. It may be suggested that lipoplexes with l ipid compositions that favor l ipid mixing at each step have a tendency to exhibit improved transfection, but the correlation is not strong. What was also remarkable, however, was that the inter-group differences were more pronounced when the luciferase plasmid was used rather than the C A T plasmid for transfecting the same cell line. The two plasmid preparations were purified by the same method and the lipoplexes were prepared in the same way by the same individual. One cannot rule out a variance in physical factors related to the plasmid preparation. However, it is also possible that: 1) sequence-level differences in the two plasmids affected gene expression in this cell line; 2) 130 secondary or tertiary structure of the two plasmids was different and this had an impact on lipoplex structure. (It was not determined, for example, the relative proportions of supercoiled vs. relaxed plasmid in the two plasmid preparations); and 3) intracellular processing of the lipoplexes (degradation, D N A release, D N A nuclear transport) differs due to plasmid factors that are influenced by lipoplex lipid composition or environment. These issues are critically important but are just beginning to be addressed in the field of D N A delivery and gene therapy. Conclusions about the roles of lipids used in the preparation of cationic liposomes or the structures adopted during lipoplex formation are tenuous when these processes are studied in isolation. For this reason, this thesis describes changes in aggregation (light scattering techniques and C T E M , Chapter 5) and membrane structure (lipid mixing R E T assay and C T E M ) under conditions that mimic those used when transfecting cells in vitro. It was demonstrated that changes in aggregation and membrane structure can occur at each step and it was concluded that the ability of the resulting membrane structures to undergo lipid mixing reactions influences vesicle aggregation and membrane structure. Although more has been learned about the structure of cationic liposomes as they interact with polynucleotides, it is still unknown exactly what the "active species" is for transfection purposes. In vitro transfection tends to be highly variable, and usually this is attributed mainly to cellular variability factors such as cell cycle, growth fraction, cytokine production, extracellular environment, etc. or to the "size" of the lipoplexes based on light scattering data. It is unknown i f there are certain structural features that facilitate cellular interactions other than the ability to allow membrane fusion. The similarity between D O P E - and DOPC-lipoplexes in the C A T transfection experiment brings up the possibility that there may be multiple mechanisms of cytoplasmic delivery (fusion dependent, fusion independent). Perhaps some substructure is present in varying degrees in all the preparations that is mediating the gene transfer. 131 Chapter 7 Development of Salt-Stable Cationic Liposomes and Lipoplexes and Characterization of Transmembrane Ion Gradients for Drug Loading 7.1 Introduction Previous chapters of this thesis have illustrated the tendency of cationic lipoplexes to aggregate into large, heterogeneous, ill-defined particles. The aim, however is to develop cationic liposomes that maintain size stability at physiologic salt concentrations, protect plasmid D N A from physical and enzymatic degradation and promote transfection. Large aggregates may be suitable for transfecting cultured cells but are clearly unacceptable for systemic administration to humans, an overall long-term goal of gene therapy research. For lipoplexes to become an effective mode of in vivo gene therapy, two aspects are of primary concern: (1) distribution of the lipoplexes in the tissue(s) of interest after administration; and (2) cellular uptake/processing of the D N A to generate transgene expression. It is hoped that lipoplex formulations can be developed which meet both of these demanding criteria. Effective cellular processing, which was addressed in Chapter 2, requires interaction between lipoplexes and the cell, lipoplex uptake, release of biologically active D N A , then finally expression. Concerns related to lipoplex biodistribution are avoidance of rapid clearance from the bloodstream, targeting to the appropriate site, and diffusion within the tissue of interest. Cationic lipid/DOPE-type liposomes, which are moderately effective in vitro, seem to meet one of the two general criteria by promoting transfection. However, it is clear that the aggregation and lipid mixing reactions of DOPE-containing cationic lipoplexes in the presence of salts generate large particles (400 nm to » 1 pm in diameter). Large aggregates are limited in their distribution and exhibit rapid clearance after i.v. administration (Osaka et al, 1997, Parker et al, 1997). They diffuse poorly in tissues even upon direct injection (Nomura et al, 1997). DOPE-containing cationic lipoplexes may be effective once they reach the cell. If distribution is 132 limited, however, and only a few cells in the tissue are transfected, the biological result of transgene expression in those few cells may be inadequate to elicit a change in the condition one is attempting to treat. It is believed therefore, that smaller, more homogeneous lipoplexes are desirable for systemic administration. This study explored alternatives to D O P E as a helper lipid. Specifically, this investigation examined whether using cholesterol as a helper l ipid (Templeton et al, 1997, Crook et al, 1998) and including poly(ethylene oxide) in the l ipid composition would eliminate the tendency to aggregate while maintaining in vitro transfection ability. Cholesterol has a stabilizing effect on liposomal membranes, while D O P E has a tendency to revert to nonbilayer configurations and to promote membrane fusion reactions. Poly(ethylene oxide), also known as poly(ethylene glycol) or P E G , has been used with conventional liposomes to enhance in vivo circulation time (Klibanov et al, 1990, Papahadjopoulos et al, 1991). When P E G is coupled to a phospholipid, such as D S P E or D M P E as used here, it can be incorporated into liposomal membranes. A s discussed in Chapter 1, P E G provides a steric barrier to liposome-liposome fusion (Mori et al, 1991) and to uptake by cells of the reticuloendothelial system (RES) such as phagocytic macrophages (Allen et al, 1991). The use of P E G in nonviral D N A delivery systems is beginning to be explored for both liposomal (Hong et al, 1997) and polymeric D N A delivery systems (Wolfert et al, 1996), which have a tendency to aggregate. This includes liposome or polymer-based D N A delivery systems containing peptides for fusion or targeting, which are currently under development. Cholesterol and PEG- l ip id containing lipoplexes are described in this chapter which do not undergo super-aggregation in salt-containing buffers. The helper l ipid substitution (cholesterol for D O P E ) could, however, render the lipoplexes less able to escape the endocytic compartment of the cells. It is generally believed that the mechanism of D O P E in transfection is to promote fusion between lipoplexes and cellular membranes, including endosomal membranes. L ip id 133 compositions that generate lipoplexes less prone to self-fusion are likely less able to fuse with cellular membranes and may need additional pharmacological assistance. Loaded agents that promote escape of the carriers from the endocytic system of the cell could be an advantage (Truong et al, 1998). For example, chloroquine is a frequently used lysosomotropic agent. Alternatively, other compounds that affect the various steps involved in transfection could be useful when coencapsulated in the lipoplexes. For the concept of codelivery to be successful, the secondary agent must be loaded into the lipoplexes in high enough concentrations to be effective. Conventional liposomes can be loaded with certain ionizable compounds at high concentrations by active loading techniques (see Chapter 1), such as by means of a p H gradient (Mayer et al, 1990). The cationic liposomes used in this study maintained a residual trapped volume after lipoplex formation (see Chapter 4), an important parameter for the encapsulation of water-soluble drugs. It was proposed that the salt-stable cationic liposomes developed here might also have the ability to establish and maintain a p H gradient in the presence of D N A . The use of ion gradient-based liposome loading techniques, however, required first the development of l ipid compositions that would not aggregate in salt-containing buffers. 7.1.1 Theoretical Considerations in the Use of Methylamine to Measure pH Gradients The validity of using a weak base like methylamine ( M A ) to determine the p H gradient across a liposome bilayer (interior acidic) can be described mathematically in terms of simplified kinetics. The following derivation is adapted from a review by Cull is et al. (1997). The starting assumptions are: the protonated form of M A is practically membrane impermeant; the compound does not partition into the membrane; the temperature is above the gel to liquid crystalline phase transition temperature (Tm) for the lipids of the bilayer; and the gradient is not significantly dissipated upon M A uptake into the liposome interior (high buffer capacity). Firstly, the total concentration of M A initially added to the system is given by: 134 [ M A ] 0 t o t a l = [ M A ] 0 + [ M A H + ] 0 , (Eq. 1) where the subscript o denotes the outside of the liposomes. M A and M A H + are the neutral and protonated forms of methylamine, respectively. The dissociation constant of M A , a weak base, is given by: Kd = [ M A ] 0 [ H + ] 0 / [ M A H + ] 0 (Eq. 2) Thus, [ M A ] 0 = Kd ( [MAH+] 0 ) / [H + ] (Eq. 3) Substituting Eq . 3 into Eq. 1 and rearranging gives: [ M A ] 0 = [ M A ] t o t a l / (1 + [ H + ] 0 / Kd) (Eq. 4) Because the p K a of M A is 10.6 and the p H of the buffer external to the liposomes was approximately 7.0, [ H + ] 0 » K d , which allows simplification of Eq . 4 to: [ M A ] 0 = (Kd [ M A ] t o t a l ) / [ H + ] 0 (Eq. 5) Initially, there is no methylamine inside the liposomes ([MA]j = 0, where i denotes liposomal interior). The total number (N) of molecules of M A available for uptake is determined by the external volume times the drug concentration, N = ( [ M A ] 0 t o t a l ) ( V 0 ) (Eq.6) The rate of uptake into the liposomes is given by: d [ M A ] 0 t o t a l / dt = - d [ M A ] 0 / dt (Eq. 7) The available surface area of the liposomes (S) and the permeability (P) of the drug through the lipid bilayer must also be take into account in describing the rate of uptake, however, which gives: dN/dt = P S ( [ M A ] 0 - [MA]j). Because [MA]j = 0 initially, and using Eq . 6, dN/dt = -(PS / V 0 ) / ( [ M A ] 0 - [MA],) = -(PS / V 0 ) / [ M A ] 0 (Eq. 8) Upon substituting Eq. 7 into Eq. 8, ' d[MA]0tota7dt = - ( P S / V 0 ) [ M A ] 0 (Eq. 9) which can be expressed in terms of the external p H (from Eq. 3) to yield: d[MA]0tota7dt = - {(PSKd)/V 0 [H + ] 0 } [ M A ] 0 t o t a l (Eq. 10) 135 Thus the rate constant for M A uptake (k) can be defined as k = (PSKd)/V0[H ] 0 and d [MA] 0 t o t a l / d t = -k [ M A ] 0 t o t a l (Eq. 11) which integrates to: [ M A ( t ) ] 0 t o t a l = [ M A ( 0 ) ] o t o t a l e- k t, where t refers to time (Eq. 12) A t a given time t, the methylamine concentration inside the liposomes would be: [MA(t) ] , t o t a l = ( [ M A ( 0 ) o t o t a l - [MA(t)]0 t o t a ,)Vo/Vi (Eq. 13) If all the methylamine initially added to the system goes inside the liposomes, which have a total internal volume Vj, then V o [ M A ( 0 ) ] o t o t a l = V , [MA(max) ] i t o t a l (Eq. 14) where [MA(max)] j t o t a l is the maximum possible concentration of methylamine inside the liposomes. Based on Eq. 12, this relationship can be expressed as: [MA(t) ] , t o t a l = [MA(max)]i t o t a l (1 - e k t ) (Eq. 15) Recall that k = (PSKd)/V 0[H +] 0, so that to maximize uptake of a weak base, the external p H should be set to maximize the proportion of the unionized (membrane permeant) form of the compound, while maintaining [H+]»Kd. In addition, V 0 can be minimized by keeping the liposome concentration high. Finally, assuming that Kd inside the liposomes is the same as Kd outside the liposomes, the Henderson-Hasselbach equation can be used: [ H + ] 0 / [ M A H + ] 0 = [H+]i / [ M A H + ] j (Eq. 16) To reach equilibium, M A w i l l continue entering the liposomes until [MA]j = [ M A ] 0 , but when M A enters the acidic liposome interior, it becomes M A H + as long as protons are available. Because it was assumed in the beginning that the internal buffer capacity of the liposomes is high, Eq . 5, can be used to describe the equilibium situation: [ M A H + ] i / [ M A H + ] 0 = [FT], / [ H + ] 0 s [MA] i t o t a l / [ M A ] 0 t o t a l (Eq. 17) Therefore it should now be clear that the uptake of a weak base such as M A is dependent not only on the membrane permeability of the drug but also on the magnitude of the p H gradient. For example, a p H gradient of 2 units should result in 100 times greater concentration of M A 136 inside the liposomes compared to outside. The p H gradient can be determined by measuring the M A gradient, or other easily detected weak base that can rapidly penetrate the liposomal bilayer but not leak out rapidly once trapped in the ionized form (for example, 3 H -tetraphenylphosphonium). For M A , the largest gradient that can be reliably measured experimentally is approximately 3.6 p H units, because with larger gradients, the external [H + ] approaches ICj, and the above relationship does not hold. (Harrigan et al, 1992). 7.2 Results Based on the observation that the inclusion of D O P E in the cationic liposomes causes lipoplex aggregation, an alternative helper l ipid was sought. Templeton et al. (1997) showed that DOTAP/cholesterol lipoplexes were effective for transfection of many tissues following systemic administration to mice. In the present study, the inclusion of P E G - l i p i d was anticipated to inhibit liposome-liposome aggregation in salt. To determine i f cationic lipid/cholesterol/PEG liposomes could form stable lipoplexes in 150 m M N a C l , a series of cationic liposomes was prepared in H B S . Q E L S analysis before and after D N A complex formation was used to assess aggregation. Some of the compositions included D S P E - P E G or D M P E - P E G (0.5 to 2 mol%) for additional stabilization. Table 8 lists the mean diameters of these cationic liposomes and the lipoplexes prepared from them at a l i p i d : D N A ratio of 10:1 nmol/pg. Prior to D N A addition, liposomes were approximately 100 nm in mean diameter. DOTAP/cholesterol liposomes, containing no PEG- l ip id , aggregated when D N A in H B S was added to them to form lipoplexes. DOTAP/choles terol /DSPE-PEG (70/29.5/0.5 mol%) lipoplexes were approximately 440 nm. DOTAP/choles terol /DSPE-PEG (70/29/1 mol%), DOTAP/choles te ro l /DSPE-PEG (70/28/2 mol%) and DOTAP/choles te ro l /DMPE-PEG (70/28/2 mol%) lipoplexes all had mean diameters in the 200 nm range by Q E L S analysis, which were stable beyond 48 h. Thus, the inclusion of approximately 30 mo l% cholesterol and 1 to 2 mo l% D S P E - P E G in 100 nm D O T A P liposomes 137 conferred size stabil i ty (mean diameter <300 nm) o f l i p o s o m e / D N A complexes for an extended time. Composition (molar ratio) Liposomes (nm) Lipoplexes (nm) DOTAP/cholesterol (70/30) 104 ± 3 0 971 ± 5 3 9 DOTAP/choles te ro l /DSPE-PEG (70/29.5/0.5) 128 ± 4 3 4 4 2 ± 175 DOTAP/choles te ro l /DSPE/PEG (70/29/1) 107 ± 3 6 223 ± 102 DOTAP/choles te ro l /DSPE/PEG (70/28/2) 111 ± 3 6 2 1 6 ± 3 0 DOTAP/cho l e s t e ro l /DMPE/PEG (70/28/2) 158 ± 5 5 162 ± 4 9 Table 8 Cationic liposomes and lipoplexes containing cholesterol and P E G are stable in 150 m M N a C l . Liposomes were prepared in H B S and extruded at R T . Lipoplexes were prepared with 25 pg D N A / m l and 250 nmoles total l ipid/ml (10:1 l i p i d : D N A ratio). Particle sizing by Q E L S was done after 30 min incubation at R T . Inhibi t ion o f l iposome aggregation by P E G cou ld also be an indica t ion that P E G was inh ib i t ing l i p o s o m e - D N A bind ing . It was necessary to assess the abi l i ty o f P E G - l i p i d containing l iposomes to protect the p l a smid D N A from nuclease degradation. L ipop lexes and naked p la smid D N A were incubated w i t h D N a s e I fo l lowed gel electrophoresis o f the extracted p lasmid (Figure 7.1). 1 2 3 4 5 6 7 8 Figure 7.1 Cholesterol-containing cationic liposomes impart DNase I protection. Lane 1: untreated plasmid; 2: treated free plasmid; 3: D O T A P / D O P E lipoplexes; 4: D O T A P / D O P C ; 5: D O T A P / C h o l / D M P E - P E G (70/28/2); 6: D O T A P / C h o l / D S P E - P E G (70/28/0.5); 7: D O T A P / Cho l /DSPE-P E G (70/28/1); 8: D O T A P / C h o l / D S P E - P E G (70/28/2). Lipoplexes were prepared with 25 pg D N A / m l and 250 nmol total l ipid/ml in H B S , except for samples 3 and 4, which were prepared in 300 m M lactose. 138 DNase protection using these formulations prepared in H B S was superior to that of D O T A P / D O P E or D O T A P / D O P C lipoplexes prepared in lactose. DNase I degraded the free plasmid (lane 2). D O T A P / D O P E and D O T A P / D O P C lipoplexes provided some protection, as indicated by the faintly visible bands migrating the same distance as untreated free plasmid. There was appreciable D N A digestion in the D O T A P / D O P E and D O T A P / D O P C lipoplexes after DNase treatment, however, indicated by the appearance of a low M W smear toward the bottom of the gel (lanes 3 and 4). M u c h less degradation was seen in lanes 5-8, corresponding to DOTAP/cholesterol /PEG lipoplexes prepared in H B S . Slightly more low M W fragments were seen in lane 5, which contained plasmid extracted from DNase-treated DOTAP/choles te ro l /DMPE-PEG (70/28/2 mol%) lipoplexes than those in which D S P E - P E G (0.5 to 2 mol%) was used. This may be because D M P E - P E G , with its shorter acyl chain, is less stable than D S P E - P E G in the lipoplexes. The salt-stable lipoplexes were at least as effective as those prepared with D O T A P / D O P E liposomes for transfection of B16 /BL6 melanoma cells in vitro as previously described. Under serum-free conditions, DOTAP/cholesterol /PEG (70/29.5/0.5 mol%) lipoplexes showed superior transfection compared to D O T A P / D O P E lipoplexes (Figure 7.2A). Where results refer to significant differences between groups, a one-way A N O V A test was performed. Differences were considered significant when p <0.05. (Analyses were done using the Mic roCa l Origin 5.0 software program.) Further increasing the PEG- l ip id concentration to 1 or 2 mol%, which was necessary to maintain particle size under 300 nm (Table 8), decreased luciferase expression significantly, although no additional toxicity was noted by visual observation or total cellular protein content. Based on these results and the particle size data, further studies used an intermediate concentration of 1.5 mo l% D S P E - P E G . With this concentration of PEG- l ip id , these lipoplexes 139 consistently had stable mean diameters <300 nm that still could promote measurable transfection in vitro. Consideration of the cationic l ipid component was also important. U p to this point of the study, D O T A P was used in combination with cholesterol based on the work of Templeton et al. (1997). Previous studies in this laboratory, however, had shown excellent transfection results with D O D A C as the cationic lipid reagent. A direct comparison was made of the transfection ability o f lipoplexes made from D O D A C vs. D O T A P , with 50 m o l % D O P E or D O P C (Figure 7.2B). D O P E was chosen as the helper lipid because it consistently generates transfection in vitro when used in combination with a variety of cationic lipids. D O P C was included as an alternative helper l ipid, albeit a less effective one. The level o f expression in cells receiving D O T A P / D O P E - or DOTAP/DOPC-l ipoplexes was not significantly different from free D N A . Cells receiving D O D A C / D O P E lipoplexes expressed 25-fold higher levels of luciferase activity than those receiving D O T A P / D O P E . D O D A C was therefore substituted for D O T A P in subsequent studies. The transfection ability o f cationic lipid/cholesterol/PEG lipoplexes is an important criterion in their development as D N A carriers, as is their stability in electrolyte solutions. For developing the salt-stable lipoplexes further as carriers o f secondary drug compounds it was necessary then to characterize the liposomes and lipoplexes under the conditions required for maximal drug encapsulation, i.e. in salt-containing buffers with an ion or p H transmembrane gradient. 140 Figure 7.2 Transfection of B16 cells with lipoplexes prepared in HBS. Liposomes containing DOPE were first prepared in lactose prior to lipoplex formation in HBS. Data represent mean ± SD (n = 6). Percentages indicated on abscissa of A refer to m o l % of DSPE-PEG in DODAC/DOPC/Cholesterol/PEG lipoplexes. In (A): *significantly different (p<0.05) from free D N A ; **significantly different from 1%. In (B): ^significantly different from free DNA; ** significantly different from DOTAP/DOPE. 141 Cationic liposomes and lipoplexes containing 28.5 mo l% cholesterol and 1.5 mo l% D S P E -PEG2000 did not aggregate under p H gradient conditions. Cationic lipid/cholesterol/PEG liposomes and lipoplexes were prepared in citrate buffer (pH 4.0) then diluted with citrate buffer or an excess of H B S (pH 7.0). Mean diameter was determined by Q E L S analysis and the results are shown in Table 9A. A s expected, the neutral liposomes exhibited no change in their mean diameter when plasmid D N A was added. DODAC/DOPC/cho les te ro l /PEG liposomes were approximately 100 nm before plasmid addition and approximately 240 nm afterwards. Liposomes alone were stable under these conditions for at least 7 days. For effective encapsulation of water-soluble compounds within liposomes, an adequate internal aqueous volume (trapped volume, Vt) is essential. It was shown in Chapter 4 that D O D A C / D O P E liposomes retain aqueous contents after lipoplex formation. The same method was used in this study to assess the trapped volume of the salt-stable lipoplexes. The trapped volume of cationic lipid/cholesterol/PEG liposomes and lipoplexes prepared in citrate buffer (pH 4.0) then diluted with citrate buffer or an excess of H B S (pH 7.0) was measured. In the absence of D N A , DODAC/choles terol /PEG (70/28.5/1.5 mol%) liposomal trapped volume (Vt = 1.73 ± 0.5) was less than that of the neutral liposomes composed of DOPC/cholesterol/PEG (70/28.5/1.5 mol%) (Vt = 4.62 ± 1.02) under p H gradient conditions (Table 9B). D N A was added to DODAC/choles terol /PEG (70/28.5/1.5 mol%) liposomes to achieve a charge ratio (+/-) of approximately 2.2. The trapped volume of the resulting lipoplexes was only 0.55 ± 0.12 pl/pmol. To use the observation that the V t of DOPC/cholesterol/PEG liposomes (Vt = 4.62 ± 1.02 in the absence o f a p H gradient) was nearly 3-fold greater than DODAC/choles terol /PEG liposomes, D O P C was then included in the cationic liposomes. It was anticipated that the trapped volume would be larger than that of DODAC/choles tero l /PEG (70/28.5/1.5 mol%) liposomes and lipoplexes. Thereafter, 30 mol% D O P C was included in the cationic liposomes. 142 For DODAC/DOPC/choles te ro l /PEG (40/30/28.5/1.5 mol%) liposomes under p H gradient conditions, V t = 3.18 ± 1.2 pl/pmol. This trapped volume is slightly larger than that of DOPC/cholesterol/PEG liposomes when a p H gradient is in effect (Vt = 1.2 ± 0.15). DODAC/DOPC/choles te ro l /PEG liposomes retained their liposomal trapped volume after D N A complexation. N o significant loss of V t was observed in most samples (Vt = 3.17 ± 1.8). A. Composition (molar ratio) p H Gradient Established D N A Added Mean Diameter (nm) DOPC/cholesterol/PEG (70/28.5/1.5) (-) (-) 122 ± 3 7 (+) (-) 113 ± 32 (+) (+) 114 ±32 DODAC/DOPC/cholesterol/PEG (40/30/28.5/1.5) (-) (-) 121 ± 3 2 (+) (-) 104 ± 2 3 (+) (+) 242 ± 85 B. Composition (molar ratio) p H Gradient Established D N A Added Trapped Volume (p,l/pmol) DODAC/cholesterol/PEG (70/28.5/1.5) (+) (-) 1.73 ±0 .5 (+) (+) 0.55 ±0.12 DOPC/cholesterol/PEG (70/28.5/1.5) (-) (-) 4.62 ± 1.02 (-) (+) 2.75 ±0.84 (+) (-) 1.20 ±0.15 (+) (+) 1.47 ±0.36 DODAC/DOPC/cholesterol/PEG (40/30/28.5/1.5) (+) (-) 3.18 ± 1.2 (+) (+) 3.17 ± 1.8 Table 9 Mean diameters and trapped volumes of liposomes and lipoplexes. A: Liposomes were prepared in citrate buffer, pH 4.0. Liposomes were diluted into HBS, pH 7.0, to create a pH gradient (+) or into citrate buffer [(-) gradient], which either contained D N A (+) or not (-), followed by QELS analysis. Data represent mean ± intrasample SD. For cationic liposomes + D N A , the charge ratio was approximately 2.2. B: Liposomes were prepared as in (A). Where indicated, D N A in HBS was added to achieve a charge ratio (positive:negative) of approximately 2.2 and 50 pg DNA/ml . Trapped volume was measured by the filtration method using 14C-lactose as an aqueous trapped volume marker. Data represent mean ± SD of 3 to 7 samples. 143 Because it was clear that the salt-stable lipoplexes could maintain their trapped aqueous contents, it was proposed that they might also be able to maintain an ion gradient. If a stable p H gradient could be established then the loading of ionizable drugs, such as putative transfection enhancers, might be possible by active loading procedures (see section 1.2.2.2). The ion gradient was established by the difference in buffer p H between the liposome interior and the bulk phase and measured by the redistribution of 1 4C-mefhylamine ( M A ) in response to the transmembrane p H gradient (Harrigan et al, 1992). A t p H 7, M A is in the nonionized (neutral) form and can readily cross the liposomal bilayer. Neutral M A w i l l continue to enter the liposome interior to establish an equilibrium of neutral M A inside and outside the liposomes. Once it reaches the liposome interior, however, it becomes ionized at the lower p H . M A in the ionized form cannot cross the bilayer and so it becomes trapped, as long as the p H gradient is in effect. Thus, M A accumulates inside the liposomes and its concentration is directly proportional to the magnitude of the p H gradient. The p H gradient therefore can be determined by measuring the methylamine gradient, as described in the methods (see section 7.2.1). It was hypothesized that the p H gradient might vary according to the lipoplex charge ratio because it was observed previously that C R has a direct effect on trapped volume and lipid mixing behavior of D O D A C / D O P E liposomes (Wasan et al, 1998). Destabilization of liposomal bilayers when D N A complexation occurs could impinge upon the ability of the liposomes to maintain the ion gradient due to leakage. Lipoplexes were prepared at C R (+/-) = 0.5 - 4.5 and the 1 4 C - M A gradient [(dpm 0 U t Side - dpmj n Sjd e)/pmol total lipid] was measured at 1 h, with the results shown in Figure 7.3. Plasmid D N A alone did not affect the distribution of 1 4 C -M A across the Mic roCon filter. The methylamine gradient in neutral liposomes (DOPC/cholesterol/PEG) was not significantly different (p < 0.05, by one-way Anova) when D N A was added (samples 1 and 2). Cationic liposomes (DODAC/DOPC/choles terol /PEG) (sample 3) and lipoplexes at C R = 0.5 (sample 4) had similar M A gradients. Samples 5 to 8, 144 which represent single samples of cationic lipoplexes at increasing charge ratios maintained the gradient at all ratios of l ipid to D N A tested. The lowest value, for C R = 4.5, was close to the mean value for neutral liposomes in the presence of D N A . 3.5-, [3. 1 2 3 4 5 6 7 8 Figure 7.3 Ion gradients of DODAC/DOPC/cholesterol/PEG lipoplexes. The gradient was measured by the methylamine uptake method after 1 h at RT. Where error bars are indicated, data represent mean ± SD (n>4). Samples- 1: DOPC/Cholesterol/DSPE-PEG2 0 0o (70/28.5/1.5 mol%) (neutral liposomes) prepared in 300 m M sodium citrate (pH 4) to which HBS (pH 7) was added; 2: Neutral liposomes to which plasmid D N A in HBS (pH 7) was added; 3: DODAC/DOPC/cholesterol/DSPE-PEG20oo (40/30/28.5/1.5) prepared in 300 mM citrate, pH 4 (cationic liposomes) to which HBS (pH 7) was added; 4 to 8: Cationic liposomes to which plasmid D N A in HBS (pH 7) was added, to yield charge ratios of 0.5, 0.8, 1.0, 3.2 and 4.5, respectively. Bars 5 to 8 correspond to single representative samples. These methylamine gradients translate into a p H gradient of approximately 2 units (see section 2.8.1), which was the same at 8 h, the longest timepoint measured. It was anticipated that a p H gradient could be used to drive the loading of amine drugs into the liposomes and lipoplexes, similar to the way in which M A was accumulated. This concept was introduced in the thesis introduction (see section 1.2.3.2 and Figures 1.10-1.12). The first compound to which this technique was applied in this study was the putative transfection enhancer chloroquine. DODAC/DOPC/choles te ro l /PEG (70/30/28.5/1.5 mol%) liposomes were prepared in citrate 145 buffer, p H 4.0. CHES-buffered saline (CBS, p H 9.0) was used as the external buffer to maximize the applied p H gradient (see section 7.1.2). C B S containing the chloroquine was added to the liposomes all at once to establish the gradient. To form the lipoplexes, plasmid was first added to the chloroquine/CBS solution. N o loading of chloroquine was detected in either liposomes or lipoplexes at R T for 1 h or after an additional 1 h at 37°C. A s an alternative model ionizable compound, vincristine was loaded into the cationic liposomes and lipoplexes by the p H gradient method. Liposomes were prepared as described above, then vincristine was added at a lipid:drug ratio of 13:1 (wt/wt) using the p H gradient method. Negative controls for loading consisted of liposomes and lipoplexes prepared in the same way but with citrate buffer as the external medium instead of phosphate buffer so that no p H gradient was present. The positive control was sphingomyelin/cholesterol (SPH/Chol) liposomes for which vincristine encapsulation by p H gradient loading is well-established (Mayer et al, 1993). Preliminary experiments indicated that 4 h of incubation at 37°C were required to achieve 90-100% encapsulation of vincristine in control SPH/Chol liposomes (drug:lipid ratio 1:20) (Figure 7.4 inset). D N A had no effect on the loading of SPH/chol. D O D A C / D O P C / C h o l / P E G liposomes encapsulated 1.63 ± 0.24 pg vincristine/pmol l ipid (34 ± 3.7% of the initial vincristine) when the p H gradient was present compared to 3.76 ± 0.47 pg/pmol loaded into the SPH/Chol liposomes. N o vincristine was retained in D O D A C / D O P C / C h o l / P E G liposomes without the p H gradient. The corresponding lipoplexes loaded 0.98 ± 0.29 pg/pmol (21.9 ± 6%) with the gradient, which was significantly less than the liposomes alone (p = 0.042). A s an additional control, D O D A C / D O P E liposomes and lipoplexes prepared in lactose were also tested for their ability to accumulate vincristine. N o p H gradient was established because these liposomes are not stable in salt. A drug concentration of 0.45 ± 0.13 pg/pmol (14 ± 2.9%) was associated with the D O D A C / D O P E liposomes and lipoplexes, 146 Figure 7.4 Loading of vincristine into liposomes via a pH gradient. Liposomes and lipoplexes were incubated with vincristine at 37°C in the presence (+) or absence (-) of a pH gradient. In lipoplexes, lipid:DNA ratio was 10:1 (nmol/pg). * Significantly different from sample 2. Inset: Percent uptake of vincristine into control sphingomyelin/cholesterol liposomes over time. Samples: 1: Sphingomyelin/cholesterol liposomes (+) 2: DODAC/DOPC/cholesterol/DSPE-PEG liposomes (+) 3: DODAC/DOPC/cho les te ro l /DSPE-PEG lipoplexes (+) 4: DODAC/DOPC/cho les t e ro l /DSPE-PEG liposomes (-) 5: D O D A C / D O P E liposomes (-) 147 which is probably reflective of passive trapping occurring during liposome rearrangement upon addition of the phosphate buffer during the loading procedure. (Recall the effects of sodium phosphate on l ipid mixing discussed in Chapters 5 and 6.) Based on an average trapped volume for D O D A C / D O P C / C h o l / P E G lipoplexes of 1.26 pl/pmol (see Table 8B), and assuming no partitioning of the drug into the liposomal membrane, the internal vincristine concentration was approximately 0.77 mg/ml. This concentration is 6.4-fold higher than the initial concentration of free vincristine in the samples. These results suggest that a small p H gradient was in effect (i.e. less than 1 unit) after lipoplex formation at the four hour timepoint. 7.3 Discussion In summary, DODAC/DOPC/choles terol / PEG2000 cationic liposomes which avoid the problem of salt-induced super-aggregation yet maintain transfection properties are described. This formulation has an excellent trap volume and can maintain a stable p H gradient of several units. The ability to maintain an ion gradient suggests that these liposomes may be amenable to development of codelivery systems consisting of D N A and ionizable drugs for transfection enhancement or other applications. Although chloroquine could not be loaded by the p H gradient method in these liposomes, vincristine was successfully loaded. Aggregation prevention is a significant advance for the pharmaceutical development of cationic liposome/plasmid D N A complexes for gene therapy. One set of confounding factors in assessing the pharmacological properties of lipoplexes is their physical polymorphism, instability and lack of robustness in preparation. Reasons for variability in activity can be ambiguous; they can be due to physico-chemical parameters of the liposomes and D N A , or due to biological factors. Elimination of the aggregation phenomenon, which is achieved here, should help to limit this problem by stabilizing the lipoplexes. 148 Substituting cholesterol and D O P C for D O P E as the helper lipids in lipoplex formulations resulted in more stable complexes (Tables 8 and 9). This is l ikely due to a more rigid, less fusogenic bilayer in the cholesterol-containing liposomes. The mechanism o f its role as a helper lipid, however, has not yet been elucidated. It is anticipated that cholesterol w i l l have value in the systemic administration of the lipoplexes (Templeton et al, 1997), by reducing opsonization and increasing circulation time. These presumptions are based on the effects of cholesterol in conventional liposomes used for drug delivery. D O P C was added to the DODAC/choles terol /PEG formulation in an effort to achieve larger trapped volume. Addition of D O P C did not reduce stability in salt, as indicated by comparing mean diameters of the formulations listed in Tables 8 and 9. Lipoplexes containing D O P C also had a larger V t than DODAC/choles terol /PEG lipoplexes at an equivalent charge ratio. The inclusion of <2 mo l% PEG- l ip id to the lipoplexes further reduced their tendency to aggregate in salt-containing buffers (Table 8). A s with conventional liposomes, the P E G may be forming a hydrophilic steric barrier over the lipoplex surface (or part of the surface) that inhibits liposome-liposome contact, preventing the aggregation reaction from starting. It was considered possible that the steric barrier over the liposome surface due to the P E G would prevent lipoplex formation altogether. Clearly, the data in this chapter support the idea that complexation between plasmid D N A and the cationic liposomes does occur when D S P E - P E G is included at molar ratio of up to 2%. The mean diameters of DOPC/choles terol /DSPE-PEG and D O D A C / DOPC/cholesterol /DSPE-PEG liposomes are quite similar in size (Table 9). The mean diameter of the DODAC-containing liposomes, however, more than doubles when D N A is added, indicating lipoplex formation. The DNase incubation experiments indicated that addition of the PEG- l ip id containing liposomes to the plasmid D N A protected the D N A from enzymatic degradation, which also supports the idea that PEG-containing cationic liposomes can bind D N A . 149 Several possibilities exist concerning the structure of the salt-stable lipoplexes that may account for the trapped volume results. Liposomes loaded with the aqueous trap marker were mixed with plasmid DNA, which would cause liposome-crosslinking as the first step of the process. The first possibility is that due to the lipid composition modifications made here (sterically hindered due to PEG, less fusogenic due to cholesterol), no further structural changes occur. This is illustrated schematically in Figure 7.5A below. This idea is akin to the beads-on-a-string model of lipoplex morphology (Feigner et al, 1987). If some of the liposomes do begin to fuse, perhaps not all the liposomes go through the process. In that case, the trapped volume measured may represent relatively few, intact liposomes fully loaded, rather than liposomes or lipoplexes that have fused and leaked part of their contents (scenario B). A third possibility, somewhat more remote, is that all the liposomes have fused and that the final new structure of complexed lipid and D N A has an internal aqueous volume (scenario C). Such a structure would have less aqueous trapped volume than the liposomes prior to D N A addition because of partial loss of contents upon fusion during the D N A complexation reaction. A B C Figure 7.5 Schematic illustration of three possible fates of liposomal entrapped aqueous contents (shaded regions) after interaction with plasmid D N A (depicted as strands). In A , liposomes become aggregated together but do not fuse. In B , some liposomes fuse during the D N A complexation reaction, but not all . In C, al l the liposomes fuse into a new structure which possesses internal aqueous contents. Nevertheless, the results of the trap volume (Table 9) and ion gradient measurements (Figure 7.3) are quite encouraging for the development of lipoplexes that encapsulate other small 150 molecules for additional biological functionality. These results also open the possibility of encapsulating strongly ionic compounds for basic investigations into cationic liposome or lipoplex behavior. For example, it is very difficult to encapsulate carboxyfluorescein, A N T S / D P X , or terbium chloride into D O D A C / D O P E liposomes because the lipid reverts to hexagonal phase l ipid clumps rather than liposomes. These compounds can now be used to study leakage and contents mixing phenomenon of the new salt-stable cationic liposomes developed here, to further our understanding of the lipoplex self-assembly process. In spite o f the improved stability against nuclease attack, the addition of PEG- l ip id at molar ratios >1% decreased lipoplex transfection ability in vitro (Figure 7.2). When 0.5 mol% P E G -lipid was included in the liposomes, the resulting lipoplexes transfected B16 melanoma cells at least as well as conventional D O D A C / D O P E lipoplexes. The dramatic decrease in luciferase expression as a function of PEG- l ip id concentration probably reflects an inability o f the lipoplexes to come into direct contact with the cells because of the steric barrier imposed by the P E G polymer layer. It is also possible that PEG- l ip id containing complexes do not settle onto the cell monolayer as well as D O D A C / D O P E large aggregates but their relative densities or sedimentation rates were not determined. However, this should not raise great concerns because these salt-stable lipoplexes are being developed for systemic use in vivo, and this activity may not be reflective of their in vivo efficacy in transfection. It was hypothesized that the stabilization of the lipoplexes with PEG- l ip id , which likely hindered interaction with cells on the tissue culture plate, might actually be beneficial in vivo. P E G might prevent rapid R E S uptake or destabilization in vivo. It is hoped that the transfection activity of the salt-stable lipoplexes may be improved by the coencapsulation of transfection-enhancing compounds. Some examples of lysosomotropic compounds that have been used experimentally in attempts to enhance in vitro transfection include chloroquine (Erbacher et al., 1996), monensin (Legendre and Szoka, 1992, Erbacher et 151 al, 1996), brefeldin A (Budker et al, 1996), sucrose (Ciftci et al, 1996), glycerol or bafilomycin (Zauner et al, 1998). These compounds disrupt the function or integrity of endosomes or lysosomes by different mechanisms. Compounds which have activity in endosomes/lysosomes and inhibit their function are known as lysosomotropic agents. Typically, cells are pre-treated before transfection. Chloroquine, the most frequently used putative transfection enhancer for cationic lipoplexes, buffers endosomal p H , reducing the acidity of the compartment. Thus lysosomal enzymes, which are pH-dependent, would be less effective in degrading the plasmid D N A . The effects of the lysosomotropic agents seem to be cell-line dependent (Strydom et al, 1993) and formulation-dependent, varying in effectiveness when used in combination with lipoplexes of differing l ipid composition. When D N A carrier uptake is receptor-mediated, such as by means of a targeting ligand, endosomolytic agents seem to be especially helpful (Zenke et al, 1990, Mis l i ck et al, 1995, Forminaya et al, 1998). However, some untargeted liposomal or polymeric D N A delivery methods are not improved by pretreating the cells with chloroquine (Legendre and Szoka, 1992, Haensler and Szoka, 1993, Mack et al, 1998). In the latter case, either the lipoplexes/polyplexes still cannot release all the D N A into the cytoplasm, regardless o f endosomal/lysosomal p H , or nonendocytic processes predominate in carrier uptake and D N A release. Presently the effectiveness of chloroquine as a transfection enhancer must be determined empirically. Alternatively, drugs that stimulate D N A repair or transcription could enhance transgene expression when co-loaded into the lipoplexes. A number of chemotherapeutic agents generate various D N A repair responses (reviewed in Pratt et al, 1994, Chaney and Sancar, 1996). Changes in transgene expression have been noted when chemotherapeutic agents were administered with lipoplexes, for example cisplatin (Son and Huang, 1996) or vincristine (Mortimer et al, 1999). A s an anticancer agent, cisplatin causes D N A crosslinking, leading to double strand breaks. The presence of D N A strand breaks induces D N A repair processes. 152 Although it is presently only theoretical and highly speculative, it is possible that stimulation of D N A repair could be harnessed to enhance transgene expression because of the induction or recruitment of nuclear factors important in D N A synthesis and transcription. It is also possible that the main effect is synchronization of cells. The highest level o f transfection is observed at M phase of cell cycle (Mortimer et al, 1999), presumably because at mitosis, the nuclear membrane dissolves, allowing access of the transgene to cellular transcription machinery. Caution must be taken to ensure that such an agent does not damage the transgene itself, but it is possible that the packaging of the D N A with the lipids may afford protection, as it does against DNase degradation or D N A intercalating dye binding. A s a related approach, directly including transcription factors specific for the transgene promoter have been considered for use in D N A delivery vehicles (Poxon et al, 1997). Other parts of the pathway between D N A uptake and degradation or transgene expression have been identified as points of intervention (Reston et al, 1995). Microtubule inhibitors, such as vincristine, colchicine, nocodazole or cytochalasin D are candidates as transfection enhancers because intracellular vesicular transport requires microtubule activity. The processing of early endosomes to late endosomes/lysosomes involves microtubule function. In fact, microtubule inhibitors are often used to uncover the intracellular processing routes of endocytosed materials. Vincristine's mechanism of action involves inhibition of microtubule formation, leading to metaphase arrest of mitotic cells and in vitro synchronization. Vincristine might also retard lipoplex transfer from early endosomes to lysosomes, with the delay thereby enhancing the chance of D N A escape into the cytoplasm. The encapsulation of vincristine within the lipoplex could increase its potency at the site of action (the cells to be transfected) and reduce toxic systemic exposure. Preliminary studies ongoing in this laboratory have shown that pretreatment of cells in tissue culture with vincristine results in a moderate increase in transgene expression after lipoplex administration (L. Nicholson and M . Bal ly , unpublished results). It should be 153 emphasized, however, that vincristine is being used in this thesis only as a model compound for the loading of ionizable drugs into cationic liposomes and lipoplexes. The effects of lipoplex -encapsulated vincristine on transgene expression were not investigated. High concentrations of ionizable drugs can be loaded into conventional liposomes. This process is amenable to optimization with the lipoplexes described here by varying the ratio of lipid, D N A and drug. This may be possible because the lipoplex charge ratio did not significantly influence the transmembrane p H gradient (Figure 7.3). Encapsulation of drugs into liposomes is affected by l ipid composition, the technique employed and certainly the physico-chemical properties of the drug itself. This chapter demonstrated the inability to encapsulate chloroquine while up to 25% of vincristine was retained in DODAC/DOPC/choles te ro l /DSPE-P E G lipoplexes, possibly reflecting a difference in membrane permeability of the two compounds. The chemical structures of chloroquine and vincristine are shown in Figure 7.6. 4 Figure 7.6 Chemical structures of chloroquine diphosphate (top) and vincristine sulfate (bottom). Source: Sigma Aldrich Internet website (http//www.sigma-aldrich.com). 154 The pKa's of chloroquine are 8.2 and 10.8 while those of vincristine are 5.0 and 7.4. Chloroquine may buffer the acidic p H of the liposome interior better than vincristine. Thus, chloroquine might be reducing the p H gradient immediately when it enters the liposomes, resulting in no retention of the drug by the time the first sample was measured. The external p H was approximately 9.0 for both loading experiments. Most of the vincristine would have been unionized at that p H and able to cross into the liposome, but part of the chloroquine would be ionized and therefore impermeant. The higher M W of vincristine may make its membrane permeability less than that of chloroquine. It is possible that extending the incubation time could improve vincristine loading, because in preliminary experiments, a lower concentration of vincristine was measured in both the cationic liposomes and control sphingomyelin/cholesterol liposomes at 1 h. Hydrophobicity alone, however, does not always directly correlate to retention in liposomes. Table 10 shows the uptake and retention of several drugs related to and including chloroquine and vincristine. Neither water solubility nor octanol/water partition coefficient alone is sufficient to explain differences in drug retention. Quinine Chloroquine Vincristine Vinblastine Doxorubicin Uptake at 15 min a (nmol/pmol lipid) 148 104 178 175 202 Uptake at 2 h a (nmol/pmol lipid) 81 88 130 127 203 Log octanol/water partition coefficient 1.7 4.3b 2.8 1.68c 1.1 Apparent maximum solubility in 300 m M citrate (mM) 1.05 585 >35 19.1 0.24 Table 10 Comparison of liposome uptake of drugs with differing physico-chemical properties. Adapted from Madden et al, 1990. "Into EPC/cholesterol liposomes with a 3 unit pH gradient (interior acidic) at 23°C. bIn 0.1M NaOH (Ferrari et al, 1991). cIn 0.9% NaCl (Grieg et al, 1990). 155 A second area o f consideration is that in the liposome loading experiments described in this chapter, the drug and D N A were added at the same time to the liposomes. During the time when the p H gradient was being established, communication between internal and external liposome environments might have reduced the p H gradient by loss/exchange of buffers. Different results might have been obtained i f the D N A had been added to form the lipoplexes and allowed to come to an equilibrium prior to establishing the p H gradient for drug loading. Thirdly, investigators who have successfully encapsulated chloroquine in anionic or neutral liposomes have chosen lipid compositions that are in the gel state at physiological temperature (Agrawal et al, 1987, Peeters et al, 1989). The liposomes used in this study, however, were probably in the liquid crystalline state, because D O D A C and D O P C have phase transition temperatures (Tc) well below 0°C. Fortunately, there are a number of potential transfection-enhancing agents, some of which have physical properties that would allow formulation of the compound into the lipoplex for codelivery applications. Modifications to the l ipid composition could also be made to optimize the retention of a particular agent. Overall, these are encouraging results. The data suggest that future studies with these or similar lipoplexes, with or without loaded transfection-enhancing compounds, may show that in vivo transfection is achievable by a systemic route. The goal, which is now realistic, is to reach transfection levels equal to or greater than that of the conventional lipoplexes presently in use, without the pharmaceutical problems associated with variable preparations and salt-induced aggregation. 156 8. Summarizing Discussion 8.1 Summary of the Work In the beginning of this project, it was understood that lipoplexes were heterogeneous and inefficient D N A delivery systems, but advantageous because of their low toxicity and ability to promote uptake of nonviral D N A into cells. It was believed that with modifications to the lipid composition and method of preparation, lipid-based carriers could be an effective modality of gene therapy. A s the work progressed, it became clear that understanding the fundamental biophysical interactions between the liposomes during the complexation reaction and subsequent to it would provide insight into how to optimize the formulations. It was hoped that a relationship between lipoplex structure and function could be discerned. It was unclear, for example, whether the liposomal bilayers were fusing together to form a new structure with the bound D N A or i f the liposomes were aggregating together intact after D N A binding. This was investigated by studying the fate of intraliposomal contents, examining lipoplex structure through electron microscopy studies and by performing lipid-mixing assays. Loss of contents, major morphological changes and significant lipid bilayer mixing during complexation to D N A suggested that bilayer fusion was occurring. A multi-step lipid-mixing assay was developed to mimic the processes of in vitro transfection. Each step causes structural changes in the liposomes and lipoplexes and it was hypothesized that these alterations in l ipid organization may influence lipoplex behavior. D O D A C / D O P E lipoplexes were more fusogenic than D O D A C / D O P C lipoplexes even in the presence of salts, which corresponded to the greater transfection ability of DOPE-containing lipoplexes compared to those containing D O P C . To avoid the extensive aggregation that occurs when D O P E is included, however, the lipid composition was then modified to produce salt stable lipoplexes. B y including cholesterol and DSPE-PEG2000 in the cationic liposomes instead of D O P E , the lipoplexes did not aggregate in salt but were still transfection-competent. Liposomal entrapped contents were retained in the 157 salt-stable lipoplexes. This observation lead to preliminary studies on the encapsulation of ionizable drugs into the lipoplexes using a p H gradient to drive drug entrapment. Vincristine was successfully encapsulated into the salt-stable cationic lipoplexes, indicating the potential for codelivery systems for D N A and drugs. 8.2 Physical Characterization of Lipoplexes Contributes to Advances in Gene Therapy This thesis began with the idea that to increase the effectiveness of gene therapy using liposome-based D N A carriers the physical structure and biophysical properties of the carriers must be understood to provide a rational basis for further design improvements. The ideal D N A lipid-based D N A carrier would have the following properties: • Protects D N A from enzymatic degradation • Delivers D N A to the desired target organ and desired cell type after systemic administration • Is taken up in sufficient quantity by the desired cell type • Releases the D N A intracellularly in the appropriate cellular compartment prior to D N A degradation • Can be made in a reproducible manner • Can be scaled up for pharmaceutical manufacturing • Is nontoxic and nonimmunogenic A s mentioned above, one of the primary areas of concern was the tendency for cationic liposome/plasmid D N A complexes to aggregate in salt-containing solutions. For the eventual development of D N A carriers for systemic delivery, small and stable particles are desirable. This thesis focused on characterizing the aggregation phenomenon and preventing its occurrence. B y determining how lipid composition influenced particle size, the cationic liposomes were reformulated to prevent lipoplex precipitation in the presence of salts. L ip id composition also 158 affected liposome and lipoplex fusion behavior during the processes of in vitro transfection. Less fusogenic lipoplexes tended to be less effective transfection reagents. Herein lies the dilemma: highly fusogenic lipoplexes are large, heterogeneous and give poorly reproducible transfection results, while less fusogenic, stable, small, monodisperse lipoplexes may have more desirable pharmaceutical properties but transfect poorly. This project suggested a means to facilitate transfection with the salt-stable lipoplexes and move beyond this impasse. B y including an adjuvant, namely a lysosomotropic agent, intracellular D N A release might be improved and thereby enhance transfection. This class of transfection enhancing agents is only one possibility however, because the process of in vitro or in vivo transfection is multipartite, providing many points of potential pharmacologic intervention. The preliminary proof of principle is provided in this work, in that secondary agents can be coencapsulated within the lipid carrier. 8.3 The Next Steps A continuation in this line of work would likely include an optimization of the lipid and PEG- l ip id composition of the salt-stable lipoplexes to maximize transfection and to improve drug uptake and retention. Maintenance of an ion gradient was observed here for up to eight hours, suggesting that cholesterol-containing cationic liposomes should be amenable to p H gradient drug loading. The choice of loaded agent is also an important consideration. Several lysosomotropic agents as well as other classes of potential transfection-enhancing drugs are candidates, as discussed in Chapter 7. The ideal agent must not only be able to be loaded into the cationic liposomes, but must also be effective in promoting D N A release or activity within the cell. Thus encapsulation of the lysosomotropic agent should result in higher transgene expression levels. The timing of D N A release and secondary agent release from the l ipid carrier w i l l therefore be critical for transfection optimization. 159 The use of P E G to prevent lipoplex aggregation was explored in this thesis and is an area of recent interest in the literature. The use of PEG-l ipids of varying chain length to modify P E G retention and thereby the timing of surface exposure (Mori et al., 1998) w i l l surely be useful in the application of cationic liposomes to D N A delivery. Ideally, the P E G - l i p i d would be retained long enough to permit lipoplex preparation and in vitro handling without aggregation prior to systemic administration. After administration, the P E G - l i p i d would then leave the lipoplex at a rate inversely proportional to the l ipid anchor chain length, thus directly influencing the pharmacokinetic profile of the carrier (Adlakha-Hutcheon et al, 1999). The chain length should be chosen to maximize delivery of the lipoplexes to the target organ. The stabilized lipoplexes should exhibit longer circulation times, which would improve delivery to tissues other than organs of the reticuloendothelial system, similar to what has been observed for stealth liposomes discussed in Chapter 1. This approach could provide a better platform for targeted delivery as well . After release or exchange of the PEG- l ip id out of the lipoplex, the lack of steric hindrance and subsequent exposure of the cationic surface could then promote cellular uptake of the lipid carriers. Coupling the sophisticated use of PEG-lipids with coencapsulation of transfection enhancers could result in further improvements in lipid-based gene therapy beyond that of only modifying the l ipid composition. Whether or not the plasmid D N A must be in a condensed form for optimal transfection is another area that must be resolved. Nevertheless, there is enthusiasm for lipid-based technology, and interest in its clinical application is growing, as indicated by clinical gene therapy trials ongoing worldwide. Up to 1996, only about 10% of human gene therapy clinical trials involved the use of liposome-based vectors (Anderson et al., 1996). B y 1998, closer to 20% used nonviral methods (Wivel and Wilson, 1998). 160 8.4 The Future of Gene Therapy A s we continue to learn more about controlled release delivery systems for macromolecules, whether the carrier is lipid-based, polymeric, peptide or some combination thereof, there w i l l l ikely be small increments in the efficiency of nonviral D N A delivery. Likewise, as we develop better methods to target the cell type of interest, specificity w i l l improve. Regardless of what the carriers are composed, however, they w i l l continue to face the limitations inherent in any particulate delivery system: limited biodistribution after systemic administration, typically to liver, spleen and to a lesser extent, the lung and to tumors i f present. This limited range may be due to interactions with cells of the reticuloendothelial system or simply a sieving effect of the organs with extensive capillary beds. The former may be reduced through the use of surface shielding, such as with the PEG-l ipids discussed in this thesis. In the latter case, particle size can only be reduced to a certain point and this type of clearance may be unavoidable for nonviral D N A carriers in the present state. Direct or local administration to the tissue of interest may be more appropriate for particulate D N A delivery methods. If this technology is to yield viable pharmaceutical products then the problem of rapid elimination must be overcome. One should bear in mind, however, that a similar conclusion was reached regarding the potential of conventional liposomes for drug delivery 20 years ago. Further work since then has resulted in the development of several commercial drug products formulated with lipids or liposomes. Some studies suggest that delivery of the D N A to the target cells is necessary but clearly insufficient (Reimer et al., 1998). A major hurdle to overcome is the short duration of expression achieved by plasmid D N A , whatever the delivery method. The explosion in molecular biology and cellular biology research has brought the previously inconceivable idea of gene therapy through to the present stage of early application to treat disease in human beings. These preliminary tests have shown that maintaining expression of a transgene is difficult. 161 (I l l I I I I II 1 DNA Chromosome gene i • mm—i—H—»-°NA TranJtipfion 5-UT • yur O H M M H M H K Z - J 5AAAAAM mRNA Traqfcfon m 1 i — Z Z ^ B O ^ M J C O O H Protein Glycosylation, M ^ p , and membrane insertion i i £k fh Ah rOj JUUW> ,5,4 3,6 , ,7 8 9,W,11,18, 12—If 15 16,1718 ITh r™**on*yP« I I II I I I I I I I I III III . = in+ame deMon • = rrcxsense • j j * j5 SI * — spacing iMr Protein product Figure 8.1 Depiction of the multiple steps involved in producing a protein. DNA consists of coding and noncoding sequences. The coding sequences serve as a template for mRNA production (transcription), which in turn is used to produce nascent peptide (translation). Some proteins are modified after translation, such as by glycosylation. After assembly and folding, the proteins can then go to the appropriate cellular compartment and begin functioning. D N A also contains noncoding regions, some of which are regulatory elements. Mutations in DNA, either in the coding regions or in regulatory regions, can have an adverse effect on the production of functional proteins. By sequencing and functional analysis, genes responsible for certain diseases have been identified. For replacement of faulty genes with functional copies, known as gene therapy, the normal regulation of the native gene should be well understood. Tissue-specific expression of specific genes, for example, results from the regulation of transcription or translation of the gene through the interaction of D N A binding factors, coregulatory proteins and/or gene promoter modification (such as methylation of CpG islands in promoter regions of DNA). In addition, until the interaction of the target host cell factors with the gene ofinterest and its regulatory regions are fully described, the extent and duration of expression from the newly introduced transgene may be suboptimal. 162 It is unreasonable to expect complete success of gene therapy protocols without further improvements in the design of plasmids for expression in mammalian cells. Because it w i l l be very difficult to deliver the D N A only to the specific cells affected by the disease, ideally, one would like to be able to induce high-level expression only in the cells of interest and be able to turn it off again at w i l l . This requires a high level of understanding and control over the promoters and enhancers used, and knowledge of how the entire transgene interacts with the transcription and translation machinery of the host cells (Figure 8.1). Ongoing work in this area w i l l be essential for optimizing human gene therapy. Another area of great concern is the induction of immune response, not only to the D N A delivery system, but also to D N A sequences (such as unmethylated C p G repeats of bacterial origin) and the newly expressed protein product. Contributions from the field of immunology in how to avoid such a response w i l l also be necessary. 8.5 Conclusions Three hypotheses were introduced in Chapter 1. The first stated that physico-chemical properties of the cationic liposomes and lipoplexes w i l l be identified that are important for achieving and/or optimizing transfection. This research has shown that cationic liposomes with D O P E had greater increases in particle size after the addition of D N A or salt-contining solutions than those with D O P C . Electron microscopy indicated major structural rearrangements of the liposomes occurred after D N A addition that depended on the l ipid composition and the presence of salts. The fusogenic behavior of the liposomes, believed to be a critical factor for sucessful transfection, was influenced by l ipid composition, l i p i d : D N A ratio and the presence of salts. The second hypothesis was that aggregation and fusion of the cationic liposomes following addition of D N A , salt or serum results in the formation of structures that are less capable of further fusion reactions required for transfection. It was found through the multistep l ipid-163 mixing assay presented in Chapter 6 that the ability to maintain fusogenic properties after dilution into salt-containing media and serum was observed in lipoplexes that transfect relatively well . D O D A C / D O P E lipoplexes were still fusion competent after dilution into serum-containing tissue culture media while D O D A C / D O P C lipoplexes were not, even though initially the liposomes had similar levels of lipid mixing during complexation to D N A . This hypothesis was supported by data throughout the thesis showing that the less fusogenic cationic l ip id /DOPC-containing lipoplexes do not transfect as well as cationic l ip id /DOPE lipoplexes. The third hypothesis was that retention of basic liposome properties, such as aqueous trapped volume, w i l l facilitate development of liposomes that can deliver D N A and secondary drugs. Replacement of D O P E with cholesterol and PEG- l ip id resulted in salt-stable lipoplexes that can transfect cells in vitro. The large trapped volume and ability to maintain an ion gradient was used to load a model ionizable amine drug, vincristine, into the liposomes via a p H gradient. This demonstrates that cationic liposome D N A carriers may be useful for codelivery of secondary agents. In conclusion, analysis of the biophysical properties of cationic liposome/plasmid D N A complexes did lead to the development of new formulation strategies to modify the properties of nonviral D N A delivery systems. Specific roles of the l ipid components were identified that contributed to lipoplex aggregation problems and in vitro transfection. A new avenue for further optimization of lipoplexes has been opened. 164 9. References Abra, R. M . and Hunt, C . A . (1981) Liposome disposition in vivo. III. Dose and vesicle-size effects. Biochim. Biophys. Acta 666(3), 493-503. Acsadi, G . , Dickson, G. , Love, D . R., Jani, A . , Walsh, F. S., Gurusinghe, A . , Wolff, J. A . and Davies, K . E . (1991) Human dystrophin expression in mdx mice after intramuscular injection of D N A constructs. Nature 352(6338), 815-818. Adlakha-Hutcheon, G . , Bal ly , M . B . , Shew, C. R. and Madden, T. D . (1999) Programmable fusogenic vesicles: systemic application as delivery vehicles for the anticancer agent, mitoxantrone. Nature Biotech., in press. Agrawal, A . K . , Singhal, A . and Gupta, C. M . (1987) Functional drug targeting to erythrocytes in vivo using antibody-bearing liposomes as drug vehicles. Biochem. Biophys. Res. Comm. 148(1), 357-361. Al len , T. M . , Austin, G . A . , Chonn, A . , L i n , L . and Lee, K . C. (1991a) Uptake of liposomes by cultured mouse bone marrow macrophages: influence of liposome composition and size. Biochim. Biophys. Acta 1061(1), 56-64. Al len , T. M . , Hansen, C. , Martin, F., Redemann, C. and Yau-Young, A . (1991b) Liposomes containing synthetic l ipid derivatives of poly(ethylene glycol) show prolonged circulation half-lives in vivo. Biochim. Biophys. Acta 1066(1), 29-36. Al ley , M . C. , Scudiero, D . A . , Monks, A . , Hursey, M . L . , Czerwinski, M . J., Fine, D . L . , Abbott, B . J., Mayo, J. G . , Shoemaker, R. H . and Boyd, M . R. (1988) Feasibility of drug screening with panels of human tumor cell lines using a microculture tetrazolium assay. Cancer Res. 48(3), 589-601. Almgren, M . , Edwards, K . and Gustafsson, J. (1996) Cryotransmission electron microscopy of thin vitrified samples. Curr. Op. Colloid Interfacial Sci. 1, 270-278. Anderson, W . F. and Bordignon, C. , eds. (1996) Human gene therapy clinical trials in Europe. Hum. Gene Ther. 7(10), 1258-1259. Ansell , S. M . , Feng, F. , Lau, A . and Ahkong, L . (1997) The design and synthesis of simple quaternary ammonium salts for gene therapy formulations. Meeting Abstract, Artificial Self-Assembling Systems for Gene Delivery, Cambridge Healthtech Institute, Coronado, C A , Oct., 1997. Arnardottir, H . B . , Sveinsson, S. J. and Kristmundsdottir, T. (1995) The use of a high intensity ultrasonic processor equipped with a flow cell in the production of reverse-phase liposomes. Int. J. Pharmaceut. 117, 237-241. Arndt-Jovin, D . J. and Jovin, T. M . (1989) Fluorescence labeling and microscopy of D N A . Meth. Cell Biol. 50,417-448. 165 Arscott, P. G . , L i , A . - Z . and Bloomfield, V . A . (1990) Condensation of D N A by trivalent cations. 1. Effects of D N A length and topology on the size and shape of condensed particles. Biopolymers 30, 619-630. Ash , P. S., Bunce, A . S., Dawson, C. R. and Hider, R. C. (1978) The effect o f synthetic polymers on the electrical and permeability properties of l ipid membranes. Biochim. Biophys. Acta 510, 216-229. Ausubel, F. M . , Kingston, R. E . , Moore, D . D . , Seidman, J. G . , Smith, J. A . and Struhl, K . , eds. (1997) Protocols in Molecular Biology. Wiley and Sons, Boston, M A . Bailey, A . L . and Cull is , P. R. (1997) Membrane fusion with cationic liposomes: effects of target membrane lipid composition. Biochem. 36(7), 1628-1634. Bainton, D . (1991) The discovery of lysosomes. Cell Biol. 91, 66S-76S. Bakker-Woudenberg, I. A . , Lokerse, A . F., ten Kate, M . T. and Storm, G . (1992) Enhanced localization of liposomes with prolonged blood circulation time in infected lung tissue. Biochim. Biophys. Acta 1138(4), 318-326. Balasubramaniam, R. P., Bennett, M . J., Aberle, A . M . , Malone, J. G . , Nantz, M . H . and Malone, R. W . (1996) Structural and functional analysis of cationic transfection lipids: the hydrophobic domain. Gene Ther. 3(2), 163-172. Bal ly , M . B . , Mayer, L . D . , Hope, M . J. and Nayar, R. (1993) Pharmacodynamics of liposomal drug carriers: methodological considerations, in Liposome Technology (Gregoriadis, G. , ed.) C R C Press, Boca Raton, F L , pp.27-41. Bally, M . B . , Mayer, L . D . , Loughery, H . , Redelmeier, T., Madden, T. M . , Wong, K . , Harrigan, P. R., Hope, M . J. and Cullis , P. R. (1988) Dopamine accumulation in large unilamellar vesicles induced by transmembrane ion gradients. Chem. Phys. Lipids 47(2), 97-107'. Bally, M . B . , Zhang, Y . - P . , Wong, F. M . P., Kong, S., Wasan, E . and Reimer, D . L . (1997) L i p i d / D N A complexes as an intermediate in the preparation of particles for gene transfer: an alternative to cationic l iposome/DNA aggregates. Adv. Drug Deliv. Rev. 24, 275-290. Bangham, A . D . and Papahadjopoulos, D . (1966) Biophysical properties of phospholipids. I. Interaction of phosphatidylserine monolayers with metal ions. Biochim. Biophys. Acta 126(1), 181-184. Bangham, A . D . , Standish, M . M . and Watkins, J. C. (1965) Diffusion of univalent cations across the lamellae of swollen phospholipids. J. Moi. Biol. 13(1), 238-252. Barenholtz, Y . , Amselem, S. and Lichtenberg, D . (1979) A new method for preparation of phospholipid vesicles (liposomes) - French press. FEBS Lett. 99(\), 210-214. Barthel, F. , Remy, J.-S., Loeffler, J.-P. and Behr, J.-P. (1993) Gene transfer optimization with lipospermine-coated D N A . DNA and Cell Biol. 12(6), 553-560. 166 Bartlett, E . M . and Lewis, D . H . (1970) Spectrophotometric determination of phosphate esters in the presence and absence of orthophosphate. Analyt. Biochem. 36(1), 159-167. Battersby, B . J., Grimm, R., Huebner, S. and Cevc, G . (1998) Evidence for three-dimensional interlayer correlations in cationic l i p i d - D N A complexes as observed by cryo-electron microscopy. Biochim. Biophys. Acta 1372, 379-383. Behr, J. P., Demeneix, B . , Loeffler, J.-P. and Perez-Mutul, J. (1989) Efficient gene transfer into mammalian primary endocrine cells with lipopolyamine-coated D N A . Proc. Natl. Acad. Sci. USA 86, 6982-6986. Bellare, J. R., Davis, H . T., Scriven, L . E . and Talmon, Y . (1988) Controlled environment vitrification system: an improved sample preparation technique. J. Electron Microsc. Tech. 10(1), 87-111. Bennett, C. F. , Mirejovsky, D . , Crooke, R. M . , Tsai, Y . J., Feigner, J., Sridhar, C. N . , Wheeler C. J. and Feigner, P . L . (1998) Structural requirements for cationic l ipid mediated phosphorothioate oligonucleotide delivery to cells in culture. J. Drug Targeting 5, 149-162. Bertling, W . A . , Gareis, M . , Paspaleeva, V . , Zimmer, A . , Kreuter, J., Numberg, E . and Harrer, P. (1991) Use of liposomes, viral capsids and nanoparticles as D N A carriers. Biotech. Appl. Biochem. 13, 390-405. Bielinska, A . , Kukowska-Latallo, J. F. , Johnson, J., Tomalia, D . A . and Baker, J. R. (1996) Regulation of in vitro gene expression using antisense oligonucleotides or antisense expression plasmids transfected using starburst P A M A M dendrimers. Nucl. Acids Res. 24(\ 1), 2176-2182. Bittman, R. and Blau, L . (1972) The phospholipid-cholesterol interaction. Kinetics of water permeability in liposomes. Biochem. 11, 4831-4839. Bl igh, E . G . and Dyer, W . J. (1959) A rapid method of total l ipid extraction and purification. Can. J. Biochem. Physiol. 37(8), 911-917. Blumfield, V . A . (1991) Condensation of D N A by multivalent cations: considerations on mechanism. Biopolymers 37, 1471-1481. Bogdanov, A . A . Jr., Weissleder, R., Frank, H . W. , Bogdanova, A . V . , Nossif, N . , Schaffer, B . K . , Tsai, E . , Papisov, M . I. and Brady, T. J. (1993) A new macromolecule as a contrast agent for M R angiography: preparation, properties, and animal studies. Radiol. 187(3), 701-706. Brasier, A . R., Tate, J. E . and Habener, J. F. (1989) Optimized use of firefly luciferase assay as a reporter gene in mammalian cell lines. Biotechn. 7(10), 1116-1122. Bruinsma, R. and Mashl , J. (1998) Long-range electrostatic interaction in D N A cationic lipid complexes. Europhys. Lett. 41, 165-170. Brulet, P. and McConnel l , H . M . (1977) Structural and dynamical aspects of membrane immunochemistry using model membranes. Biochem. 16(6), 1209-217. 167 Budker, V . , Gurevich, V . , Hagstrom, J. E . , Bortzov, F. and Wolff, J. A . (1996) pH-sensitive, cationic liposomes: a new synthetic virus-like vector. Nature Biotech. 14(6), 760-764. Buschle, M . , Cotton, M . , Kirlappos, H . , Mechtler, K . , Shaffher, G . , Zauner, W. , Bimstiel , M . C. and Wagner, E . (1995) Receptor-mediated gene transfer into human T lymphocytes via binding of D N A / C D 3 antibody particles to the C D 3 T cell receptor. Hum. Gene Ther. 6(6), 753-761. Byk, G . , Dubertret, C , Escriou, V . , Frederic, M . , Jaslin, G . , Rangara, R., Pitard, B . , Crouzet, J., Wi ls , P., Schwartz, B . and Sherman, D . (1998) Synthesis, activity, and structure-activity relationship studies of novel cationic lipids for D N A transfer. J. Med. Chem. 41, 224-235. Byk, G . , Soto, J., Mattler, C, Frederic, M . and Scherman, D . (1998) Novel non-viral vectors for gene delivery - synthesis o f a second-generation library o f mono-functionalized poly-(guanidinium)amines and their introduction into cationic lipids. Biotech. Bioeng. 61, 81-87. Bywater, M . , Bywater, R. and Hellman, C. (1983) A novel chromatographic procedure for purification of bacterial plasmids. Analyt. Biochem. 132, 219-224. Cameron, D . G . , Gudgin, E . F. and Mantsch, H . H . (1981) Dependence of acyl chain packing of phospholipids on the head group and acyl chain length. Biochem. 20, 4496-4500. Canonico, A . E . , Plitman, J. D . , Conary, J. T., Meyrick, B . O. and Brigham, K . L . (1994) No lung toxicity after repeated aerosol or intravenous delivery of plasmid-cationic liposome complexes. J. Appl. Phys. 77(1), 415-9. Caplen, N . J., Kinrade, E . , Sorgi, F., Gao, X . , Gruenert, D . , Geddes, D . , Coutelle, C , Huang, L . , Alton, E . W . and Williamson, R. (1995) In vitro liposome-mediated D N A transfection of epithelial cell lines using the cationic liposome D C - C h o l / D O P E . Gene Ther. 2, 603-613. Chancerelle, Y . , Mathieu, J., Viret-Soropogui, R., Tosetti, F. , Alban, C. and Kergonou, J. F. (1993) Immunization of rabbits with proteins reacted with malonic dialdehyde ( M D A ) : kinetics and specificity of the immune response. Biochem. Molec. Biol. Intl. 29(1), 141-148. Chaney, S. G . and Sancar, A . (1996) D N A repair - enzymatic mechanisms and relevance to drug response./. Natl. Cancer Inst. 55(19), 1346-1360. Chang, N . -S . and Mattison, J. (1996) Rapid separation of D N A from ethidium bromide and cesium chloride in ultracentrifugation gradients by a desalting column. BioTechniques 14(3), 342-343. Chapman G . D. , L i m , C. S., Gammon, R. S., Culp, S. C , Desper, J. S., Bauman, R. P., Swain, J. L . and Stack, R. S. (1992) Gene transfer into coronary arteries o f intact animals with a percutaneous balloon catheter. Circ. Res. 71(1), 27-33. Chapman, D . (1975,) Phase transitions and fluidity characteristics of lipids and cell membranes. Quart. Rev. Biophys. 8, 185-235. 168 Chen, X . , L i , Y . , X i e , Y . , A iz icov ic i , S., Snodgrass, R., Wagner, T. E . and Platika, D . (1995) A novel nonviral cytoplasmic gene expression system and its implications in cancer gene therapy. Ca. Gene Ther. 2(4), 281-289. Cheung, B . C. L . , Sun, T. H . T., Leenhouts, J. M . and Cullis , P. R. (1998) Loading of doxorubicin into liposomes by forming Mn 2 + -d rug complexes. Biochim. Biophys. Acta 1414(1-2), 205-216. Chivate, M . M . and Pandit, A . B . (1995) Quantification of cavitation intensity in fluid bulk. Ultrason. Sonochem. 2(1), S19-S25. Chonn, A . and Cull is , P. R. (1995) Recent advances in liposomal drug-delivery systems. Curr. Opin. Biotech. 6(6), 698-708. Ciftci, K . , Smiley, E . , Labhasetwar, V . , Bonadio, J. and Levy, R. J. (1996) Effect of lysosomotropic agents on gene expression in vitro. Pharmaceut. Res. Suppl. 13(9), S390. Conner, J., Yatvin, M . B . and Huang, L . (1984) pH-sensitive liposomes: acid-induced liposome fusion. Proc. Natl. Acad. Sci. USA 81, 1715-1718. Cooper, M . J. (1996) Noninfectious gene transfer and expression systems for cancer gene therapy. Sent. Oncol. 23(1), 172-187. Cotton, M . , Baker, A . , Saltik, M . , Wagner, E . and Buschle, M . (1994) Lipopolysaccharide is a frequent contaminant of plasmid D N A preparations and can be toxic to primary human cells in the presence of adenovirus. Gene Ther. 1, 239-246. Cotton, M . , Langle-Rouault, F., Kirlappos, H . , Wagner, E . , Mechtler, K . , Zenke, M . , Beug, H . and Birnstiel M . L . (1990) Transferrin-polycation-mediated introduction of D N A into human leukemic cells: stimulation by agents that affect the survival of transfected D N A or modulate transferrin receptor levels. Proc. Natl. Acad. Sci. USA 57(11), 4033-4037. Cox, R. A . (1968) The use of guanidinium chloride in the isolation of nucleic acids, in: Methods in. Enzymol. (Grossman, L . and Moldave, E . , eds.) Vol .12(B), p.120. Cowan, P. J., Shinkel, T. A . , Witort, E . J., Barlow, H . , Pearse, M . J. and d 'Apice, A . J. (1996) Targeting gene expression to endothelial cells in transgenic mice using the human intercellular adhesion molecule 2 promoter. Transplantation 62(2), 155-160. Cristiano, R.J . , Smith, L . C , Kay , M . A . , Brinkely, B . R. and Woo, S. L . (1993) Hepatic gene therapy: efficient gene delivery and expression in primary hepatocytes utilizing a conjugated adenovirus-DNA complex. Proc. Natl, Acad. Sci. USA 90(24) 11548-11552. Crook, K . , McLachlan, G. , Stevenson, B . J. and Porteous, D . J. (1996) Plasmid D N A molecules complexed with cationic liposomes are protected from degradation by nucleases and shearing by aerosolisation. Gene Ther. 3(9), 834-839. 169 Crook, K . , Stevenson, B . J., Dubouchet, M . and Porteous, D . J. (1998) Inclusion of cholesterol in D O T A P transfection complexes increases the delivery of D N A to cells in vitro in the presence of serum. Gene Ther. 5, 137-143. Crouse, C. A . , Ban, J. D . and D'Aless io , J. K . (1993) Extraction of D N A from forensic-type sexual assault specimens using simple, rapid sonication procedures. BioTechniques 15(4), 643-647. Cui , C , Wani, M . A . , Wight, D . , Kopchick, J. and Stambrook, P. J. (1994) Reporter genes in transgenic mice. Transgen. Res. 3(3), 182-194. Cullis , P. R., Hope, M . J. and Tilcock, C. P. S. (1986) L ip id polymorphism and the role of lipids in membranes. Chem. Phys. Lipids 40, 127-144. Cullis , P. R., Hope, M . J., Bally, M . B . , Madden, T. D . , Mayer, L . D . and Janoff, A . S. (1987) Liposomes as pharmaceuticals, in: Liposomes: from Biophysics to Therapeutics, (Ostro, M . J . , ed.) Marcel Dekker, New York, pp.39-72. Damen, J., Regts, J. and Scherphof, G . (1982) Transfer of [14C]phosphatidylcholine between liposomes and human plasma high density lipoprotein. Partial purification of a transfer-stimulating plasma factor using a rapid transfer assay. Biochim. Biophys. Acta 712(3), 444-452. Dan, N . (1998) The structure of D N A complexes with cationic liposomes-cylindrical or flat bilayers? Biochim. Biophys. Acta 1369, 34-38. Davis, H . L . , Schleef, M . , Moritz, P., Mancini , M . , Schorr, J. and Whalen, R. G . (1996) Comparison of plasmid D N A preparation methods for direct gene transfer and genetic immunization. Biotechn. 21(1) 92-99. de Wet, J. R., Wood, K . V . , DeLuca, M . , Helinski, D . R. and Subramani, S. (1987) Firefly luciferase gene: structure and expression in mammalian cells. Molec. Cell Biol. 7(2), 725-737. de Wet, J. R., Wood, K . V . , Helinski, D . R. and DeLuca, M . (1985) Cloning of firefly luciferase c D N A and the expression of active luciferase in Escherichia coli. Proc. Natl. Acad. Sci. USA 82(23), 7870-7873. Deamer, D . W. , Prince, R. C. and Crofts, A . R. (1972) The response o f fluorescent amines to p H gradients across liposome membranes. Biochim. Biophys. Acta 274, 323-335. Dean, R. T., Jessup, W . and Roberts, C . R. (1984) Effects of exogenous amines on mammalian cells, with particular reference to membrane flow. Biochem. J. 217, 27-40. Delort, J. P., and Capecchi, M . R. (1996) T A X I / U A S - a molecular switch to control expression of genes in vivo. Hum. Gene Ther. 7(7), 809-820. Demel, R. A . and de Kruijff, B . (1976) The function of sterols in membranes. Biochim. Biophys. Acta 457(2), 109-132. 170 Du, H . , Chandaroy, P. and Hui , S. W . (1997) Grafted poly-(ethylene glycol) on l ipid surfaces inhibits protein adsorption and cell adhesion. Biochim. Biophys. Acta 1326, 236-48. Dubochet, J., Adrian, M . , Chang, J. J., Homo, J. C , Lapault, J., M c D o w a l l , A . W . and Schultz, P. (1988) Cryo-electron microscopy of vitrified specimens. Q. Rev. Biophys. 21, 129-228. Diizgiines, N . , Ni r , S., Wilshut, J., Bentz, J., Newton, C , Portis, A . and Papahadjopoulos, D . (1981) Calcium- and magnesium-induced fusion of mixed phosphatidylserine /phosphatidylcholine vesicles: effect of ion binding. J. Membr. Biol. 59, 115-125. Diizgiines, N . , Paiement, J., Freeman, K . B . , Lopez, N . B . , Wilschut, J. and Papahadjopoulos, D . (1984) Modulation of membrane fusion by ionotropic and thermotropic phase transitions. Biochem. 23(15), 3486-3494. Diizgiines, N . , Wilschut, J., Fraley, R. and Papahadjopoulos, D . (1981) Studies on the mechanism of membrane fusion: Role of headgroup composition in calcium- and magnesium-induced fusion of mixed phospholipid vesicles. Biochim. Biophys. Acta 642, 182-195. Dzau, V . J., Mann, M . J., Morishita, R. and Kaneda, Y . (1996) Fusogenic viral liposome for gene therapy in cardiovascular diseases. Proc. Natl. Acad. Sci. USA 93 (21), 11421-11425. Eastman, S. J., Siegal, C , Tousignant, J., Smith, A . E . , Cheng, S. H . and Scheule, R. K . (1997) Biophysical characterization of cationic lipid: D N A complexes. Biochim. Biophys. Acta 1325(1), 41-62. Edwards, K . , Johnsson, M . , Karlsson, G . and Silvander, M . (1997) Effect of polyethylene glycol-phospholipids on aggregate structure in preparations of small unilamellar liposomes. Biophys. J. 73, 258-266. Eidelman, O., Schlegel, R., Tralka, T. S. and Blumenthal, R. (1984) pH-dependent fusion induced by vesicular stomatitis virus glycoprotein reconstituted into phospholipid vesicles. J. Biol. Chem. 259(1), 4622-4628. Ellens, H . , Bentz, J. and Szoka, F. C. (1984) pH-induced destabilization of phosphatidylethanolamine-containing liposomes: role of bilayer contact. Biochem. 23, 1532-1538. Ellens, H . , Bentz, J. and Szoka, F. C. (1986) Fusion of phosphatidylethanolamine-containing liposomes and the mechanism of the L a -Hn phase transition. Biochem. 25, 4141-4147. Erbacher, P., Bousser, M . T., Raimond, J., Monsigny, M . , Midoux, P. and Roche, A . C. (1996) Gene transfer by DNA/glycosylated polylysine complexes into human blood monocyte-derived macrophages. Hum. Gene Ther. 7(6), 721-729. Erbacher, P., Roche, A . C , Monsigny, M . and Midoux, P. (1996) Putative role of chloroquine in gene transfer into a human hepatoma cell line by DNA/lactosylated polylysine complexes. Exp. Cell Res. 225(1), 186-194. 171 Farhood, H . , Gao, X . , Son, K . , Yang, Y . - Y . , Lazo, J. S., Huang, L . , Barsoum, J., Bottega, R. and Epand, R. (1994) Cationic liposomes for direct gene transfer in therapy of cancer and other diseases. Ann. New York Acad. Sci. 716, 23-35. Farhood, H . , Serbina, N . and Huang, L . (1995) The role of dioleoyl phosphatidylethanolamine in cationic liposome mediated gene transfer. Biochim. Biophys. Acta 1235, 289-295. Fasbender, A . , Marshall, J., Moninger, T. O., Grunst, T., Cheng, S. and Welsh, M . J. (1997) Effect o f co-lipids in enhancing cationic lipid-mediated gene transfer in vitro and in vivo. Gene Ther. 4, 716-725. Feigner, P. L . and Ringold, G . M . (1989) Cationic liposome-mediated transfection. Nature 537(6205), 387-388. Feigner, P. L . , Gadek, T. R., Holm, M . , Roman, R., Chan, H . W. , Wenz, M . , Northrop, J. P., Ringold, G . M . and Danielsen, M . (1987) Lipofection: a highly efficient, lipid-mediated D N A -transfection procedure. Proc. Natl. Acad. Sci. USA 84,14-12-14X1. Feigner, J. H . , Kumar, R., Sridhar, C. N . , Wheeler, C. J., Tsai, Y . J., Border, R., Ramsey, P., Martin, M . and Feigner, P. L . (1994) Enhanced gene delivery and mechanism studies with a novel series of cationic l ipid formulations. J. Biol. Chem. 269(4), 2550-2561. Fenske, D . B . , Wong, K . F. , Mauer, E . , Mauer, N . , Leenhouts, J. M . , Boman, N . , Amankwa, L . and Cullis , P. R. (1998) Ionophore-mediated uptake of ciprofloxacin and vincristine into large unilamellar vesicles exhibiting transmembrane ion gradients. Biochim. Biophys. Acta 1414(1-2), 188-204. Ferkol, T., Pellicena-Palle, A . , Eckman, E . , Perales, J. C , Trzaska, T., Tosi, M . , Redline, R. and Davis, P. B . (1996) Immunologic responses to gene transfer into mice via the polymeric immunoglobulin receptor. Gene Ther. 5(8), 669-678. Ferrari, M . E . , Nguyen, C. M . , Zelphati, O., Tsai ,Y. L . and Feigner, P . L . (1998) Analytical methods for the characterization of cationic l ipid nucleic acid complexes. Hum. Gene Ther. 9(3), 341-351. Finkelstein, M . C. and Weissmann, G . (1979) Enzyme replacement via liposomes. Variations in lipid compositions determine liposomal integrity in biological fluids. Biochim. Biophysc. Acta 587(2), 202-216. Fishman, G . I., Kaplan, M . L . and Buttrick, P. M . (1994) Tetracycline-regulated cardiac gene expression in vivo. J. Clin. Invest. 93(4), 1864-1868. Flasher, D . , Konopka, K . , Chamow, S. M . , Dazin, P., Ashkenazi, A . , Pretzer, E . and Duzgunes, N . (1994) Liposome targeting to immune deficiency virus type 1-infected cells via recombinant soluble C D 4 and C D 4 immunoadhesin (CD4-IgG). Biochim. Biophys. Acta 1194(1), 185-196. Fominaya, J., Uherek, C. and Wels, W . (1998) A chimeric fusion protein containing transforming growth factor-alpha mediates gene transfer via binding to the E G F receptor. Gene Ther. 5(4), 521-530. 172 Fraley, R., Subramani, S., Berg, P. and Papahadjopoulos, D . (1980) Introduction of liposome-encapsulated SV40 D N A into cells. J. Biol. Chem. 255, 10431-10435. Friend, D . S., Papahadjopoulos, D . and Debs, R. J. (1996) Endocytosis and intracellular processing accompanying transfection mediated by cationic liposomes. Biochim. Biophys. Acta 1278(1), 41-50. Fung, B . K . K . and Stryer, L . (1978) Surface density determination by fluorescence energy transfer. Biochem. 17(24), 5241-5248. Gabizon, A . A . (1992) Selective tumor localization and improved therapeutic index of anthracyclines encapsulated in long-circulating liposomes. Cancer Res. 52(4), 891-896. Gabizon, A . and Papahadjopoulos, D . (1988) Liposome formulations with prolonged circulation time in blood and enhanced uptake by tumors. Proc. Natl. Acad. Sci. USA. SJ(18), 6949-6953. Gabizon, A . , Price, D . C. , Huberty, J., Bresalier, R. S. and Papahadjopoulos, D . (1990) Effect of liposome composition and other factors on the targeting of liposomes to experimental tumors: biodistribution and imaging studies. Cancer Res. 50(19), 6371-6378. Gao, X . and Huang, L . (1996) Potentiation of cationic liposome-mediated gene delivery by polycations. Biochem. 35(3), 1027-1036. Gerloni, M . , Billetta, R., Xiong, S. D . and Zanetti, M . (1997) Somatic transgene immunization with D N A encoding an immunoglobulin heavy chain. DNA and Cell Biol. 16(5), 611-625. Gorman, C. M . , Moffat, L . F. and Howard, B . H . (1982) Recombinant genomes which express chloramphenicol acetyltransferase in mammalian cells. Molec. Cell Biol. 2(9): 1044-1051. Gustafsson, J., Arvidson, G. , Karlsson, G . and Almgren M . (1995) Complexes between cationic liposomes and D N A visualized by c ryo -TEM. Biochim. Biophys. Acta 1235(2), 305-312. Haensler, J. and Szoka, F. C. (1993) Polyamidoamine cascade polymers mediate efficient transfection o f cells in culture. Bioconj. Chem. 4(5), 372-379. Hallahan, D . E . , Mauceri, H . J., Seung, L . P., Dunphy, E . J., Wayne, J. D . , Hanna, N . N . , Toledano, A . , Hellman, S., Kufe, D . W . and Weichselbaum, R. R. (1995) Spatial and temporal control of gene therapy using ionizing radiation. Nature Med. 7(8), 786-791. Halle, J . P., Bourassa, S., Leblond, F . A . , Chevalier, S., Beudry, M . , Chapdelaine, A . , Cousineau, S., Saintonge, J. and Yale, J. F. (1993) Protection of islets of Langerhans from antibodies by microencapsulation with alginate-poly-L-lysine membranes. Transpl. 55(2), 350-354. Hanna, Z . , Fregeau, C , Prefontaine, G . and Brousseau, R. (1984) Construction of a family of universal expression plasmid vectors. Gene 30, 247-250. Hara, T., L i u , F., L i u , D . and Huang, L . (1997) Emulsion formulations as a vector for gene delivery in vitro and in vivo. Adv. Drug Del. Rev. 24, 265-271. 173 Harrigan, P. R., Hope, M . J., Redelmeier, T. E . and Cull is , P. R. (1992) Determination of transmembrane p H gradients and membrane potentials in liposomes. Biophys. J. 63, 1336-1345. Harrigan, P. R., Wong, K . F., Redelmeier, T. E . , Wheeler, J. J. and Cull is , P. R. (1993) Accumulation of doxorubicin and other lipophilic amines into large unilamellar vesicles in response to transmembrane p H gradients. Biochim. Biophys. Acta 1149, 329-338. Hart, S. L . , Harbottle, R. P., Cooper, R., Mi l le r , A . , Will iamson, R. and Coutelle, C. (1995) Gene delivery and expression mediated by an integrin-binding peptide. Gene Ther. 2(8), 552-554. Harvie, P., Wong, F. M . P. and Bally, M . B . (1998) Characterization of l ipid D N A interactions -I - destabilization of bound lipids and D N A dissociation. Biophys. J. 75(2), 1040-1051. Haugland, R. P. (1996) Handbook of Fluorescent Probes and Research Chemicals. (Spence, T. Z. , ed.) Molecular Probes, Eugene, OR. Hirt, B . (1967) Selective extraction of polyoma D N A from infected mouse cell cultures. J. Molec. Biol. 26(2), 365-369. Hofland, H . and Huang, L . (1995) Inhibition of human ovarian carcinoma cell proliferation by liposome-plasmid D N A complex. Biochem. Biophys. Res. Comm. 207(2), 492-496. Hofland, H . E . , Shephard, L . and Sullivan, S. M . (1996) Formation of stable cationic l i p i d / D N A complexes for gene transfer. Proc. Natl. Acad. Sci. USA 93(14), 7305-7309. Holland, J. W. , Hu i , C , Cullis , P. R. and Madden, T. D . (1996) Polyethylene glycol)-lipid conjugates regulate the calcium-induced fusion of liposomes composed of phosphatidylethanolamine and phosphatidylserine. Biochem. 35(8), 2618-2624. Holt, C , Parker, T. G . and Dalgleish, D . G . (1975) Measurement of particle sizes by elastic and quasi-elastic light scattering. Biochim. Biophys. Acta 400(2), 283-292. Hong, K . , Zheng, W. , Baker, A . and Papahadjopoulos, D . (1997) Stabilization of cationic liposome-plasmid D N A complexes by polyamines and poly(ethylene glycol)-phospholipid conjugates for efficient in vivo gene delivery. FEBS Lett. 400(2), 233-237. Hope, M . J., Bally, M . B . , Webb, G . and Cull is , P. R. (1985) Production of large unilamellar vesicles by a rapid extrusion procedure: characterization of size distribution, trapped volume, and ability to maintain a membrane potential. Biochim. Biophys. Acta 812, 55-65. Hughes, J. A . , Aronsohn, A . I., Avrutskaya, A . V . and Juliano, R. L . (1996) Evaluation of adjuvants that enhance the effectiveness of antisense oligonucleotides. Pharmaceut. Res. 13(3), 404-410. Hui , S. W. , Stewart, T. P., Yeagle, P. L . and Albert, A . D . (1981) Bilayer to non-bilayer transition in mixtures o f phosphatidylethanolamine and phosphatidylcholine: implications for membrane properties. Arch. Biochem. Biophys. 207(2), 227-240. 174 Hui , S. W. , Langner, M . , Zhao, Y . - L . , Ross, P., Hurley, E . and Chan, K . (1996) The role of helper lipids in cationic liposome-mediated gene transfer. Biophys. J. 71, 590-599. Jiao, S., Will iams, P., Berg, R. K . , Hodgeman, B . A . , L i u , L . , Repetto, G . and Wolff, J. A . (1992) Direct gene transfer into nonhuman primate myofibers in vivo. Hum. Gene Ther. 3(1), 21-33. Jiao, S., Acsadi, G . , Jani, A . , Feigner, P. L . and Wolff, J. A . (.1992) Persistence of plasmid D N A and expression in rat brain cells in vivo. Exper. Neurol. 15(3), 400-413. Kabanov, A . V . and Kabanov, V . A . (1995) D N A complexes with polycations for the delivery of genetic material into cells. Bioconj. Chem. 6, 7-20. Kabanov, A . V . , Vinogradov, S. V . , Suzdaltseva, Y . G . and Alakhov, V . Y . (1995) Water-soluble block polycations as carriers for oligonucleotide delivery. Bioconj. Chem. 66, 639-643. Ka l in , B . , Sellin, P., von Krusenstierna, S., Schnell, P. O. and Jacobsson, H . (1991) Effect of size fractionation on the distribution of an albumin colloid in the reticuloendothelial system of the mouse. Intl. J. Rad. Applic. Instrumen. - Part B, Nucl. Med. Biol. 18(1), 817-820. Kaneda, Y . , Iwai, K . and Uchida, T. (1989) Increased expression o f D N A cointroduced with nuclear protein in adult rat liver. Science 243, 374-378. Kato, T., Iwamoto, K . , Ando, H . , Asakawa, N . , Tanaka, I., Kikuch i , J. and Murakami, Y . (1996) Synthetic cationic amphiphile for liposome-mediated D N A transfection with less cytotoxicity. Biol. Pharmaceut. Bull. 19(6), 860-863. Kawabata, K . , Takakura, Y . and Hashida, M . (1995) The fate of plasmid D N A after intravenous injection in mice-involvement of scavenger receptors in its hepatic uptake. Pharmaceut. Res. 12(6), 825-830. Kenworthy, A . K . , Simon, S. A . and Mcintosh, T. J. (1995) Structure and phase behaviour of lipid suspensions containing phospholipids with covalently attached poly(ethylene glycol). Biophys. J. 68, 1903-1920. Kichler, A . , Mechtler, K . , Behr, J.-P. and Wagner, E . (1997) Influence of membrane-active peptides on lipospermine/DNA complex mediated gene transfer. Bioconj. Chem. 8, 213-221. Kirby, C , Clark, J. and Gregoriadis, G . (1980) Cholesterol content of small unilamellar liposomes controls phospholipid loss to high density lipoproteins. FEBSLett. Ill, 324-328. Klibanov, A . L . , Maruyama, K . , Torchilin, V . P. and Huang, L . (1990) Amphipathic polyethyleneglycols effectively prolong the circulation time of liposomes. FEBS Lett. 268(1), 235-237. Koltover, I., Salditt, T., Radler, J. O. and Safinya, C. R. (1998) A n inverted hexagonal phase of cationic l iposome-DNA complexes related to D N A release and delivery. Science 281(5313), 78-81. 175 Kondo, T., Ara i , S.-L, Kuwabara, M . , Yoshi i , G . and Kano, E . (1985) Damage in D N A irradiated with 1.2 M H z ultrasound and its effect on template activity o f D N A for R N A synthesis. Rad. Res. 104(3), 284-292. Kricka, L . J. (1991) Chemiluminescence and bioluminescence. Clin. Chem. 37(9), 1472-1481. Kulkarni, S. B . , Betageri, G . V . and Singh, M . (1995) Factors affecting microencapsulation of drugs in liposomes. J. Microencap. 12(3), 229-246. Labat-Moleur, F. , Steffan, A . M . , Brisson, C , Perron, H . , Feugeas, O., Furstenberger, P., Oberling, F. , Brambilla, E . and Behr, J.-P. (1996) A n electron microscopy study into the mechanism of gene transfer with lipopolyamines. Gene Ther. 3(11), 1010-1017. Ladbrooke, B . D . , Will iams, R. M . and Chapman, D . (1968) Studies on lecithin-cholesterol-water interactions by differential scanning calorimetry and X-ray diffraction. Biochim. Biophys. Acta 150(3), 333-340. La i , B . , Cahan, M . A . , Couraud, P. O., Goldstein, G . W . and Laterra, J. (1994) Development of endogenous beta-galactosidase and autofluorescence in rat brain microvessels: implications for cell tracking and gene transfer studies. J. Histochem. Cytochem. 42(7), 953-956. Lasic, D . D . and Templeton, N . S. (1996) Liposomes in gene therapy. Adv. Drug Del. Rev. 20(2-3), 221-266. Lasic, D . D . and Ruff, D . (1998) Cationic liposomes, D N A and gene delivery, in Medical Applications of Liposomes. (Lasic, D . D . , and Papahadjopoulos, D . , eds.) Elsevier, The Netherlands. Laws, G . M . and Adams, S. P. (1996) Measurement of 8-OHdG in D N A by H P L C - E C D : the importance of D N A purity. BioTechniques 20(1), 36-38. Ledley, F. D . (1995) Nonviral gene therapy: the promise of genes as pharmaceutical products. Hum. Gene Ther. 6, 1129-1144. Lee, R. J. and Huang, L . (1996) Folate-targeted anionic liposome-entrapped polylysine-condensed D N A for tumor-cell specific gene transfer. J. Biol. Chem. 271(4), 8481-8487. Lee, R. J. and Low, P. S. (1995) Folate-mediated tumor cell targeting of liposome-entrapped doxorubicin in vitro. Biochim. Biophys. Acta 1233(2), 134-144. Legendre, J . -Y. and Szoka, F. C. (1992) Delivery of plasmid D N A into mammalian cell lines using pH-sensitive liposomes: comparison with cationic liposomes. Pharmaceut. Res. 9(10), 1235-1242. Legendre, J . -Y. and Szoka, F. C. (1993) Cycl ic amphipathic peptide-DNA complexes mediate high efficiency transfection of adherent mammalian cells. Proc. Natl. Acad. Sci. USA 90, 893-897. 176 Legendre, J. Y . Trzeciak, A . , Bohrmann, B . , Deuschle, U . , Kitas, E . and Supersaxo, A . (1997) Dioleoylmelittin as a novel serum-insensitive reagent for efficient transfection of mammalian cells. Bioconj. Chem. 5(1), 57-63. Lepoint, T. and Mul l ie , F. (1994) What exactly is cavitation chemistry? Ultrason. Sonochem. 1(1), S13-S22. Levy, M . Y . , Barron, L . G . , Meyer, K . B . and Szoka, F. C. (1996) Characterization of plasmid D N A transfer into mouse skeletal muscle: evaluation of uptake mechanism, expression and secretion of gene products into blood. Gene Ther. 3, 201-211. Lewis, J. G . , L i n , K . Y . , Kothvale, A . , Flanagan, W . M . , Matteucci, M . D . , DePrince, R. B . , Mook, R. A . Jr., Hendren, R. W. and Wagner, R. W . (1996) A serum-resistant cytofectin for cellular delivery o f antisense oligodeoxynucleotides and plasmid D N A . Proc. Natl. Acad. Sci. USA 93(8), 3176-3181. L i , L . H . and Hui , S. W . (1997) The effect of l ipid molecular packing stress on cationic liposome-induced rabbit erythrocyte fusion. Biochim. Biophys. Acta 1323, 105-116. Liang, X . , Hartikka, J., Sukhu, L . , Manthorpe, M . and Hobart, P. (1996) Novel , high expressing and antibiotic-controlled plasmid vectors designed for use in gene therapy. Gene Ther. 3(4), 350-356. L is , J. T. (1980) Fractionation of D N A fragments by polyethylene glycol precipitation. Meth. Enzymol. 65, 347-353. Litzinger, D . C. and Huang, L . (1992) Phosphatidylethanolamine liposomes: drug delivery, gene transfer and immunodiagnostic applications. Biochim. Biophys. Acta 1113(2), 201-227. L i u , F. and L i u , D . (1996) Serum independent liposome uptake by mouse liver. Biochim. Biophys. Acta 7275, 5-11. L i u , H . W. , Ofosu, F. A . and Chang, P. L . (1993) Expression of human factor I X by microencapsulated recombinant fibroblasts. Hum. Gene Ther. 4(3), 291-301. L i u , J., Shirahama, K . , Miyajima, K . and Kwak, J. C. T. (1998) Interaction of a cationic surfactant to sodium polyphosphates with different degrees of polymerization. Colloid Polymer Sci. 276(1), 40-45. Mack, K . D . , Wei , R., Elbagarri, A . , Abbey, N . and McGrath, M . S. (1998) A novel method for DEAE-dextran mediated transfection of adherent primary cultured human macrophages. J. Immun. Meth. 277(1-2), 79-86. Madden, T. D . , Harrigan, P. R., Tai , L . C. L . , Bally, M . B . , Mayer, L . D . , Redelmeier, T. E . , Loughrey, H . C , Tilcock, C. P. S., Reinish, L . W . and Cull is , P. R. (1990) The accumulation of drugs within large unilamellar vesicles exhibiting a proton gradient: a survey. Chem. Phys. Lipids 56, 37-46. 177 Madden, T. D . , Janoff, A . S. and Cullis , P. R. (1990) Incorporation of Amphotericin B into large unilamellar vesicles composed of phosphatidylcholine and phosphatidylglycerol. Chem. Phys. Lipids 52, 189-198. Manning, G . S. (1978) The molecular theory of polyelectrolyte solutions with applications to the electrostatic properties of polynucleotides. Quart. Rev. Biphys. 11(2), 179-246. Manning, G . S. and Mohanty, U . (1997) Counterion condensation on ionic oligomers. Physica A 247, 196-204. Marquet, M . , Horn, N . A . and Meek, J . A . (1995) Process development for the manufacture o f plasmid D N A vectors for use in gene therapy. BioPharm. 8(1), 26-37. May, S. and Ben-Shaul, A . (1997) D N A - l i p i d complexes - stability of honeycomb-like and spaghetti-like structures. Biophys. J. 73, 2427-2440. Mayer, L . D . , Bal ly , M . B . and Cullis, P. R. (1986) Uptake of adriamycin into large unilamellar vesicles in response to a p H gradient. Biochim. Biophys. Acta 557(1), 123-126. Mayer, L . D . , Bal ly , M . B . , Hope, M . J. and Cullis , P. R. (1985) Uptake of dibucaine into large unilamellar vesicles in response to a membrane potential. J. Biol. Chem. 260(2), 802-808. Mayer, L . D . , Bal ly , M . B . , Loughrey, H . , Masin, D . and Cull is , P. R. (1990) Liposomal vincristine preparations which exhibit decreased drug toxicity and increased activity against murine L1210 and P388 tumors. Cancer Res. 50(3), 575-579. Mayer, L . D . , Hope, M . J. and Cull is , P. R. (1986) Vesicles o f variable sizes produced by a rapid extrusion procedure. Biochim. Biophys. Acta 858(1), 161-168. Mayer, L . D . , Nayar, R., Thies, R. L . , Boman, N . , Cull is , P. R. and Bal ly , M . B . (1993) Identification of vesicle properties that enhance the antitumour activity of liposomal vincristine against murine L1210 leukemia. Cancer Chemother. Pharmacol. 33, 17-24. Mayhew, E . , Lazo, R. and V a i l , W . J. (1984) Preparation of liposomes entrapping cancer chemotherapeutic agents for experimental in vivo and in vitro uses, in Liposome Technology (Gregoriadis, G . , ed.) C R C Press, Boca Raton F L , pp. 19-31. Mayhew, E . , Lazo, R., V a i l , W . J., K ing , J. and Green, A . M . (1984) Characterization of liposomes prepared using a microemulsifier. Biochim. Biophys. Acta. 775(2), 169-174. McKee , T. D . , DeRome, M . E . , W u , G . Y . and Findeis, M . A . (1994) Preparation of asialoorosomucoid-polylysine conjugates. Bioconj. Chem. 5(4), 306-311. Memol i , A . , Palermiti, L . G. , Travagli, V . and Alhaique, F. (1995) Egg and soya phospholipids-sonication and dilaysis: A study on liposome characterization. Int. J. Pharmaceut. 117, 159-163. 178 Meyer, O., Kirpotin, D . , Hong, K . L . , Sternberg, B . , Park, J. W. , Woodle, M . C. and Papahadjopoulos, D . (1998) Cationic liposomes coated with polyethylene glycol as carriers for oligonucleotides. J . Biol. Chem. 273, 15621-15627. Mil ler , D . L . , Thomas, R. M . and Frazier, M . E . (1991a) Single strand breaks in C H O cell D N A induced by ultrasonic cavitation in vitro. Ultrasound in Med. and Biol. 17(4), 401-406. Mil ler , D . L . , Thomas, R. M . and Frazier, M . E . (1991b) Ultrasonic cavitation indirectly induces single strand breaks in D N A of viable cells in vitro by the action of residual hydrogen peroxide. Ultrasound in Med. and Biol. 17(7), 729-735. Mis l ick , K . A . , Baldeschwieler, J. D . , Kayyem, J. F. and Meade, T. J. (1995) Transfection of folate-polylysine D N A complexes: evidence for lysosomal delivery. Bioconj. Chem. 6(5), 512-515. Mita , K . , Zama, M . and Ichimura, S. (1977) Effect of charge density o f cationic polyelectrolytes on complex formation with D N A . Biopolymers 16(9), 1993-2004. Mittal , S. K . , Bett, A . J., Prevec, L . and Graham, F. L . (1995) Foreign gene expression by human adenovirus type 5-based vectors studies using firefly luciferase and bacterial beta-galactosidase gene as reporters. Virol. 210(1), 226-230. Moghimi , S. M . , Porter, C. J., Muir , I. S., Ilium, L . and Davis, S. S. (1991) Non-phagocytic uptake of intravenously injected microspheres in rat spleen: influence of particle size and hydrophilic coating. Biochem. Biophys. Res. Comm. 177(2), 861-866. Mok , K . W . C. and Cullis , P. R. (1997) Structural and fusogenic properties of cationic liposomes in the presence of plasmid D N A . Biophys. J. 73(5), 2534-2545. M o r i , A . , Klibanov, A . L . , Torchilin, V . P. and Huang, L . (1991) Influence of the steric barrier activity of amphipathic poly(ethyleneglycol) and ganglioside G M 1 on the circulation time of liposomes and on the target binding of immunoliposomes in vivo. FEBS Lett. 284(2), 263-266. M o r i , A . , Chonn, A . , Choi , L . S., Israels, A . , Monck, M . A . and Cull is , P. R. (1998) Stabilization and regulated fusion of liposomes containing a cationic lipid using amphipathic polyethylene glycol derivatives. J. Liposome Res. 8, 195-211. Mortimer, I., Tarn, P., MacLachlan, I., Graham, R. W. , Saravolac, E . G . , Joshi, P .B . (1999) Cationic lipid-mediated transfection of cells in culture requires mitotic activity. Gene Ther. 6(3), 403-411. Mount, R. C , Jordan, B . E . and Hadfield, C. (1996) Reporter gene systems for assaying gene expression in yeast. Meth. Molec. Biol. 53, 239-248. Nabel, E . G . , Gordon, D . , Yang, Z . Y . , X u , L . , San, H . , Plautz, G . E . , W u , B . Y . , Gao, X . , Huang, L . and Nabel, G . J. (1992) Gene transfer in vivo with DNA-l iposome complexes: lack of autoimmunity and gonadal localization. Hum. Gene Ther. 3, 649-656. 179 Nabel, G . J., Gordon, D . , Bishop, D . K . , Nickoloff, B . J., Yang, Z . Y . , Aruga, A . , Cameron, M . J., Nabel, E . G . and Chang, A . E . (1996) Immune response in human melanoma after transfer of an allogeneic class I major histocompatibility complex gene with DNA-l iposome complexes. Proc. Natl. Acad. Sci. USA 93(26), 15388-15393. Nabel, G . J., Nabel, E . G. , Yang, Z . Y . , Fox, B . A . , Plautz, G . E . , Gao, X . , Huang, L . , Shu, S., Gordon, D . and Chang, A . E . (1992) Direct gene transfer with DNA-l iposome complexes in melanoma: expression, biologic activity, and lack of toxicity in humans. Proc. Natl. Acad. Sci. USA 90, 11307-11311. Nakanishi, M . , Uchida, T. and Sugawa, H . (1985) Efficient introduction of content of liposomes into cells using H V J (Sendai virus). Exp. Cell Res. 159, 399-409. Needham, D. , Hristova, K . , Mcintosh, T. J., Dewhirst, M . , W u , N . and Lasic, D . (1992) Polymer-grafted liposomes: physical basis for the "stealth" property. J. Liposome Res. 2, 411-430. Nielsen, H . and Andersen, L . P. (1992) Chemotactic activity of Helicobacter pylori sonicate for human polymorphonuclear leucocytes and monocytes. Gut 33(6), 738-742. Nomura, T., Nakahima, S., Kawabata, K . , Yamashita, F. , Takakura, Y . and Hashida, M . (1997) Intratumoral pharmacokinetics and in vivo gene expression of naked plasmid D N A and its cationic liposome complexes after direct gene transfer. Cancer Res. 57(13), 2681-2686. Ohmori, N . , Niidome, T., Wada, A . , Hirayama, T., Hatakeyama, T. and Aoyagi , H . (1997) The enhancing effect o f anionic alpha-helical peptide on cationic peptide-mediating transfection systems. Biochem. Biophys. Res. Comm. 235(3), 726-729. Okada, N . , Miyamoto, H . , Yoshioka, T., Sakamoto, K . , Katsume, A . , Saito, H . , Nakagawa, S., Ohsugi, Y . and Mayumi , T. (1997) Immunological studies of S K 2 hybridoma cells microencapsulated with alginate-poly(L-lysine)-alginate ( A P A ) membrane following allogenic transplantation. Biochem. Biophys. Res. Comm. 230(3), 524-527. Okamoto, T., Mitsuhashi, M . and Kikkawa, Y . (1994) Fluorometric nuclear run-on assay with oligonucleotide probe immobilized on plastic plates. Analyt. Biochem. 221, 202-204. Oku, N . , Yamaguchi, N . , Shibamoto, S., Ito, F. and Nango, M . (1986) The fusogenic effect of synthetic polycations on negatively charged l ipid bilayers. J. Biochem. 100, 935-944. Osaka, G . , Carey, K . , Cuthbertson, A . , Godowski, P., Patapoff, T., Ryan, A . , Gadek, T. and Mordenti, J. (1996) Pharmacokinetics, tissue distribution, and expression efficiency of plasmid [ 3 3 P ] D N A following intravenous administration of DNA/cat ionic l ipid complexes in mice-use of a novel radionuclide approach. / . Pharm. Sci. 85(6), 612-618. Papahadjopoulos, D . , Jacobson, K . , Ni r , S. and Isac, T. (1973) Phase transitions in phospholipid vesicles. Fluorescence polarization and permeability measurement concerning the effect of temperature and cholesterol. Biochim. Biophys. Acta 311, 330-348. Papahadjopoulos, D . and Poste, G . (1975) Calcium-induced phase separation and fusion in phospholipid membranes. Biophys. J. 15(9), 945-948. 180 Papahadjopoulos, D . , Al len , T. M . , Gabizon, A . , Mayhew, E . , Matthay, K . , Huang, S. K . , Lee, K . D. , Woodle, M . C , Lasic, D . D . , Redemann. C. et al. (1991) Sterically stabilized liposomes: improvements in pharmacokinetics and antitumor therapeutic efficacy. Proc. Natl. Acad. Sci. USA 55(24), 11460-11464. Parker, S. E . , Ducharme, D . , Norman, J. and Wheeler, C . J. (1997) Tissue distribution of the cytofectin component of a plasmid DNA/cat ionic l ipid complex following intravenous administration in mice. Hum. Gene Ther. 5(4), 393-401. Parsegian, A . (1969) Energy of an ion crossing a low dielectric membrane: solutions to four relevant electrostatic problems. Nature 221( 183), 844-846. Patel, H . M . , Tuzel, N . S. and Ryman, B . E . (1983) Inhibitory effect o f cholesterol on the uptake of liposomes by liver and spleen. Biochim. Biophys. Acta 761(2), 142-151. Pecheur, E . I., Martin, I., Ruysschaert, J. M . , Bienvenue, A . and Hoekstra, D . (1998) Membrane fusion induced by 11-mer anionic and cationic peptides - a structure-function study. Biochem. 37,2361-2371. Pecora, R. (1972) Quasi-elastic light scattering from macromolecules. Ann. Rev. Biophys. Bioeng. 1, 257-276. Peeters, P. A . , Brunink, B . G. , El ing, W . M . and Crommelin, D . J. (1989) Therapeutic effect o f chloroquine (CQ)-containing immunoliposomes in rats infected with Plasmodium berghei parasitized mouse red blood cells: comparison with combinations of antibodies and C Q or liposomal C Q . Biochim. Biophys. Acta 981(2), 269-276. Perkins, W . R., Dause, R. B . , Parente, R. A . , Minchey, S. R., Neuman, K . C , Gruner, S. M . , Taraschi, T. F. and Janoff, A . S. (1996) Role of l ipid polymorphism in pulmonary surfactant. Science 273(5273), 330-332. Porcher, C , Malinge, M . C , Picat, C. and Grandchamp, B . (1992) A simplified method for determination of specific D N A or R N A copy number using quantitative P C R and an automatic D N A sequencer. BioTechniques 13(1), 106-114. Pratt, W . B . (1994) The Anticancer Drugs, Oxford University Press, New York. Radler, J. O., Koltover, I., Salditt, T. and Safinya, C. R. (1997) Structure of DNA-cat ionic liposome complexes: D N A intercalation in multilamellar membranes in distinct interhelical packing regimes. Science 275(5301), 810-814. Rapaport, D . , Ni r , S. and Shai, Y . (1994) Capacities of pardaxin analogues to induce fusion and leakage of negatively charged phospholipid vesicles are not necessarily correlated. Biochem. 33(42), 12615-12624. Raymond, G . J., Bryant, P. K . , Nelson, A . and Johnson, J. D . (1988) Large-scale isolation of covalently closed circular D N A (cccDNA) using gel filtration chromatography. Analyt. Biochem. 773,125-133. 181 Redelmeier, T. E . , Mayer, L . D . , Wong, K . F. , Bal ly , M . B . and Cull is , P. R. (1989) Proton flux in large unilamellar vesicles in response to membrane potentials and p H gradients. Biophys. J. 56, 385-393. Reimer, D . L . , Kong, S. and Bal ly , M . B . (1997) Analysis o f cationic liposome-mediated interactions of plasmid D N A with murine and human melanoma cells in vitro. J. Biol. Chem. 272(31), 19480-19487. Reimer, D . L . , Kong , S., Monk, M . , Tan, P., Wasan, E . and Bal ly , M . B . (1999) Factors influencing in vivo liposome-mediated transfection of B 1 6 / B L 6 melanoma tumor models: a pharmacodynamic study. J. Pharmacol. Exp. Ther., in press. Reimer, D . L . , Zhang, Y . - P . , Kong, S., Wheeler, J. J., Graham, R. W . and Bal ly , M . B . (1995) Formation of novel hydrophobic complexes between cationic lipids and plasmid D N A . Biochem. 34(39), 12877-12883. Reston, J. T., Gould-Fogerite, S. and Mannino, R. J. (1995) Aspects of cellular physiology that influence DNA-mediated gene transfer i n N I H 3 T 3 cells. Molec. Cell. Biochem. 145(2), 169-175. Roth, J. A . and Cristiano, R. J. (1997) Gene therapy for cancer: what have we done and where are we going? J. Natl. Cancer Inst. 89, 21-39. Sambrook, J., Fritsch, E . I. and Maniatis, T., eds. (1989) Molecular Cloning: A Laboratory Manual, Co ld Spring Harbor, New York. Scherphof, G . L . , Kuipers, F. , Derksen, J. T. P., Spanjer, H . H . and Vonk, R. J. (1987) Biochem. Soc. Trans. i J (Supp l ) , 625-628. Schott, H . (1969) Hydrophile-lipophile balance and cloud points of nonionic surfactants. J. Pharm. Sci. 58(12), 1443-1449. Schwedener, R. A . , Trub, T., Schott, H . , Langhals, H . , Barth, R. F. , Groscurth, P. and Hengartner, H . (1990) Comparative studies of the preparation of immunoliposomes with the use of two bifunctional coupling agents and investigation of in vitro immunoliposome-target cell binding by cytofluorometry and electron microscopy. Biochim. Biophys. Acta 1026(1), 69-79. Seed, B . and Sheen, J . -Y. (1988) A simple phase-extraction assay for chloramphenicol acyltransferase activity. Gene 67(2), 271-277'. Senior, J., Crawley, J. C. W . and Gregoriadis, G . (1985) Tissue distribution of liposomes exhibiting long half-lives in the circulation after intravenous injection. Biochim. Biophys. Acta 839(1), 1-8. Senior, J., Delgado, C , Fisher, D . , Tilcock, C. and Gregoriadis, G . (1991) Influence of surface hydrophilicity of liposomes on their interaction with plasma protein and clearance from the circulation: studies with poly(ethylene glycol)-coated vesicles. Biochim. Biophys. Acta 1062(1), 77-82. 182 Shen, Q., van Beusechem, V . W. , Einerhand, M . P., Hendrikx, P. J. and Valeric-, D . (1991) Construction and expression of an adenosine deaminase:lacZ fusion gene. Gene 98(2), 283-287. Scherphof, G. , Roerdink, F. , Waite, M . and Parks, J. (1978) Disintegration of phosphatidylcholine liposomes in plasma as a result of interactions with high-density lipoproteins. Biochim. Biophys. Acta 542(2), 296-307'. Siegel, D . P. (1986) Inverted micellar intermediates and the transitions between lamellar, cubic, and inverted hexagonal lipid phases. I. Mechanism of the L a lpha—HII phase transitions. Biophys. J. 49(6), 1155-1170. Simon, S. A . and Mcintosh, T. J. (1989) Magnitude of the solvation pressure depends on dipole potential. Proc. Natl. Acad. Sci. USA 86(23), 9263-9267. Sleigh, M . J. (1986) A nonchromatographic assay for expression of the chloramphenicol acetyltransferase gene in eukaryotic cells. Analyt. Biochem. 156(1), 251-256. Smith, J. G . , Wedeking, T., Vemachio, J. H . , Way, H . and Niven, R. W . (1998) Characterization and in vivo testing of a heterogeneous cationic l i p i d - D N A formulation. Pharmaceut. Res. 15(9), 1356-1363. Smith, P. K . , Krohn, R. I., Hermanson, G . T., Mal l ia , A . K . , Gartner, F . H . , Provenzano, M . D . , Fujimoto, E . K . , Goeke, N . M . , Olson, B . J. and Klenk, D . C. (1985) Measurement of protein using bicinchoninic acid. Analyt. Biochem. 150(1), 76-85. Smolarsky, M . , Teitelbaum, D. , Sela, M . and Gitler, C . (1977) A simple fluorescent method to determine complement mediated liposome immune lysis. Immunol. Meth. 15, 255-265. Sokol, D . L . and Gewirtz, A . M . (1996) Gene therapy: basic concepts and recent advances. Crit. Rev. Euk. Gene Expression 6(1), 29-57. Son, K . and Huang, L . (1996) Factors influencing the drug sensitization of human tumor cells for in situ lipofection. Gene Ther. 3(1), 630-634. Sperisen, P., Wang, S. M . , Reichenbach, P. and Nabholz, M . (1992) A PCR-based assay for reporter gene expression. PCR Meth. Appl. 1(3), 164-170. Srienc, F. , Campbell, J. L . and Bailey, J. E . (1986) Flow cytometry analysis of recombinant Saccaromyces cerevisiae populations. Cytometry 7(2), 132-141. Stamatatos, L . , Leventis, R., Zuckermann, M . J. and Silvius, J. R. (1988) Interactions of cationic l ipid vesicles with negatively charged phospholipid vesicles and biological membranes. Biochem. 27(11), 3917-3925. Stamatatos, L . , Leventis, R., Zuckermann, M . J. and Silvius, J. R. (1988) Interactions of cationic lipid vesicles with negatively charged phospholipid vesicles and biological membranes. Biochem. 27(11), 3917-3925. 183 Stegrnann, T. and Legendre, J . -Y. (1997) Gene transfer mediated by cationic lipids: lack of correlation between lipid mixing and transfection. Biochim. Biophys. Acta 1325, 71-79. Sternberg, B . (1998) Ultrastructural morphology of cationic liposome-DNA complexes for gene therapy, in: Medical Applications of Liposomes. (Lasic, D . D . and Papahadjopoulos, D . , eds.) Elsevier, The Netherlands, pp.395-428. Sternberg, B . , Hong, K . , Zheng, W . and Papahadjopoulos, D . (1998) Ultrastructural characterization of cationic l iposome-DNA complexes showing enhanced stability in serum and high transfection activity in vivo. Biochim. Biophys. Acta 1365, 23-35. Sternberg, B . , Sorgi, F. L . and Huang, L . (1994) New structures in complex formation between D N A and cationic liposomes visualized by freeze-fracture electron microscopy. FEBS Lett. 356, 361-366. Struck, D . K . , Hoekstra, D . and Pagano, R. E . (1981) Use of resonance energy transfer to monitor membrane fusion. Biochem. 20, 4093-4099. Strydom, S., Van Jaarsveld, P., Van Helden, E . , Ariatti, M . and Hawtrey, A . (1993) Studies on the transfer of D N A into cells through use of avidin-polylysine conjugates complexed to biotinylated transferrin and D N A . J. Drug Targeting 1(2), 165-174. Stryer, L . (1978) Fluorescence energy transfer as a spectroscopic ruler. Ann. Rev. Biochem. 47, 819-832. Szoka, F. and Papahadjopoulos, D . (1978) Procedure for preparation of liposomes with large internal aqueous space and high capture by reverse-phase evaporation. Proc. Natl. Acad. Sci. USA 75, 4194-4198. Szoka, F. , Olson, F. , Heath, T., V a i l , W. , Mayhew, E . and Papahadjopoulos, D . (1980) Preparation of unilamellar liposomes of intermediate size (0.1-0.2pm) by a combination of reverse phase evaporation and extrusion through polycarbonate membranes. Biochim. Biophys. Acta 601(3), 559-57'1. Szoka, F. C. , X u , Y . H . and Zelphati, O. (1997) How are nucleic acids released in cells from cationic lipid-nucleic acid complexes? Adv. Drug Deliv. Rev. 24(2-3), 291. Tang, M . X . , Redemann, C. T. and Szoka, F . C. (1996) In vitro gene delivery by degraded polyamidoamine dendrimers. Bioconj. Chem. 7, 703-714. Tartakoff, A . M . (1987) The Secretory and Endocytic Paths: Mechanism and Specificity of Vesicular Traffic in the Cel l Cytoplasm. Wiley, New York. Templeton, N . S., Lasic, D . D . , Frederik, P. M . , Strey, H . H . , Roberts, D . D . and Pavlakis, G . N . (1997) Improved D N A - liposome complexes for increased systemic delivery and gene expression. Nature Biotech. 15(7), 647-652. 184 Thierry, A . R., Rabinovich, P., Peng, B . , Mahan, L . C , Bryant, J. L . and Gallo, R. C. (1997) Characterization of liposome-mediated gene delivery: expression, stability and pharmacokinetics of plasmid D N A . Gene Ther. 4(3), 226-237. Thomas, W . H . and Lee, Y . K . (1994) Particles in intravenous solutions: a review. New Zealand Med.J. 80(522), 170-178. Thurnher, M . , Wagner, E . , Clausen, H . , Mechtler, K . , Rusconi, S., Dinter, A . , Birstiel, M . L . , Berger, E . G . and Cotton, M . (1994) Carbohydrate receptor-mediated gene transfer to human T leukaemic cells. Glycobiol. 4(4), 429-435. Tilcock, C. P. (1986) L ip id polymorphism. Chem. Phys. Lipids 40(2-4), 109-125. Tomalia, D . A . and Durst, H . D . (1993) Genealogically directed synthesis: starburst/cascade dendrimers and hyperbranched structures. Top. Curr. Chem. 165, 193-313. Tomalia, D . A . , Naylor, A . M . and Goddard, W . A . (1990) Starburst dendrimers: molecular-level control o f size, shape surface chemistry, topology, and flexibility from atoms to macroscopic matter. Angew. Chem. Int. Ed. Engl. 29, 138-175. Tomita, N . , Higaki , J., Ogihara, T., Kondo, T. and Kaneda, Y . (1994) A novel gene-transfer technique mediated by H V J (Sendai virus), nuclear protein, and liposomes. Ca. Detec. Preven. 18(6), 485-491. Tomlinson, E . and Rolland, A . P. (1996) Controllable gene therapy: pharmaceutics of non-viral gene delivery systems. J. Contr.Rel. 39, 357-372. Truong-Le, V . L . , August, J. T. and Leong, K . W . (1998) Controlled gene delivery by D N A -gelatin nanospheres. Hum. Gene Ther. 9(12), 1709-1717. Truong-Le, V . L . , Walsh, S. M . , Schweibert, E . , Mao, H . Q., Guggino, W . B . , August, J. T. and Leong, K . W . (1999) Gene transfer by gelatin nanospheres. Arch. Biochem. Biophys. 361(1) 41-56. van Dijck, P. W . M . , de Kruijff, B . and van Deenen, L . L . M . (1976) The preference of cholesterol for phosphotidylcholine in mixed phosphotidylcholine-phosphatidylethanolamine bilayers. Biochim. Biophys. Acta 455(2), 576-587. van Dijck, P. W . M . , de Kruijff, B . , Verkleij, A . J., van Deened, L . L . M . and de Grier, J. (1978) Comparative studies on the effects of p H and Ca2+ on bilayers of various negatively charged phospholipids and their mixtures with phosphatidylcholine. Biochim. Biophys. Acta 512(1), 84-96. Verkleij, A . J., van Echteld, C. J., Gerritsen, W . J., Cull is , P. R. and de Kruijff, B . (1980) The lipidic particle as an intermediate structure in membrane fusion processes and bilayer to hexagonal HII transitions. Biochim. Biophys. Acta 600(3), 620-621. 185 Verma, R. S., Giannola, D . , Shlomchik, W . and Emerson, S. G . (1998) Increased efficiency of liposome-mediated transfection by volume reduction and centrifugation. Biotech. 25(1), 46-49. Vidal-Naquet, A . , Gossage, J. L . , Sullivan, T. P., Hayes, J. W. , Gilruth, B . H . , Beissinger, R. L . , Sehgal, L . R. and Rosen, A . L . (1989) Biomater. Art. Cells Art. Organs 17(5), 531-552. V i l e , R. G . and Hart I. R. (1994) Targeting of cytokine gene expression to malignant melanoma cells using tissue-specific promoter sequences. Ann. Oncol. 5(Suppl. 4), 59-65. Vogel , K . , Wang, S., Lee, R. J., Chmielewski, J. and Low, P. S. (1996) Peptide-mediated release of folate-targeted liposome contents from endosomal membranes. J. Amer. Chem. Soc. 118(1), 1581-1586. Vo-Quang, T., Malpiece, Y . , Buffard, D . , Kaminski , P .A . , V ida l , D . and Strosberg, A . D . (1985) Rapid, large-scale purification of plasmid D N A by medium or low pressure gel filtration. Application: construction of thermoamplifiable expression vectors. Biosci. Rep. 5, 101-111. Wagner, E . , Cotton, M . , Foisner, R. and Birnstiel, M . L . (1991) Transferrin-polycation-DNA complexes: the effect of polylysine on the structure of the complex and D N A delivery to cells. Proc. Natl. Acad. Sci. USA 55(10), 4255-4259. Wagner, E . , Zenke, M . , Cotton, M . , Beug, H . and Birnstiel, M . L . (1990) Transferrin-polycation conjugates as carriers for D N A uptake into cells. Proc. Natl. Acad. Sci. 57(9), 3410-3414. Wasan E . K . , Reimer D . L . and Bal ly , M . B . (1996) Plasmid D N A is protected against ultrasonic cavitation-induced damage when complexed to cationic liposomes. J. Pharm. Sci. 55(4), 427-433. Wasan, E . K . and Bal ly , M . B . (1998) Cationic l iposome/DNA complexes for gene therapy: development of salt-stable formulations and characterization of transmembrane ion gradients. AAPSPharmSci i ( l )(Suppl) Abstract 2464. <http://www.aaps.org/publications.html>. Wasan, E . K . , Fairchild, A . and Bally, M . B . (1998) Cationic liposome-plasmid D N A complexes used for gene transfer retain a significant trapped volume. J. Pharm. Sci. 57(1), 9-14. Wasan, K . M . and Cassidy, S. M . (1998) Role of plasma lipoproteins in modifying the biological activity of hydrophobic drugs. J. Pharm. Sci. 57(4), 411-424. Webb, M . S., Harasym, T. O., Masin, D . , Bal ly , M . B . and Mayer, L . D . (1995) Sphingomyelin-cholesterol liposomes significantly enhance the pharmacokinetic and therapeutic properties of vincristine in murine and human tumour models. Brit. J. Cancer 72(4), 896-904. Webb, M . S., Saxon, D . , Wong, F. M . P., L i m , H . J., Wang, Z . , Bal ly , M . B . , Choi , L . S. and Cullis , P. R. (1998) Comparison of different hydrophobic anchors conjugated to poly(ethylene glycol)-effects on the pharmacokinetics of liposomal vincristine. Biochim. Biophys. Acta-Biomembranes 1372, 272-282. Weber, M . , Moller , K . , Welzeck, M . and Schorr, J. (1995) The effects of lipopolysaccharide on transfection in eukaryotic cells. BioTechniques 19, 930-940. 186 Weinstein, J. N . , Klausner, R. D. , Innerarity, T. L . , Ralston, E . and Blumenthal, R. (1981) Phase transition release, a new approach to the interaction of proteins with l ipid vesicles. Application to lipoproteins. Biochim. Biophys. Acta 647(2), 270-284. Weintraub, H . , Cheng, P. F. and Conrad, K . (1996) Expression of transfected D N A depends on D N A topology. Cell 46, 115-122. Wheeler, C. J., Sukhu, L . , Yang, G . , Tsai, Y . , Bustamente, C , Feigner, P. L . , Norman, J. and Manthorpe, M . (1996) Converting an alcohol to an amine in a cationic l ipid dramatically alters the co-lipid requirement, cellular transfection activity and the ultrastructure of DNA-cytofectin complexes. Biochim. Biophys. Acta 1280, 1-11. Wicks, I. P., Howell , M . L . , Hancock, T., Kohsaka, H . , Olee, T. and Carson, D . A . (1995) Bacterial lipopolysaccharide copurifies with plasmid D N A : implications for gene therapy. Hum. Gene Ther. 6, 317-323. Wilschut, J. and Papahadjopoulos, D . , (1979) Ca 2 + -induced fusion of phospholipid vesicles monitored by mixing of aqueous contents. Nature 281, 691-692. Wilschut, J., Scholma, J., Bental, M . , Hoekstra, D . and Ni r , S. (1985) Ca 2 + -induced fusion of phosphatidylserine vesicles: mass action kinetic analysis of membrane l ipid mixing and aqueous contents mixing. Biochim. Biophys. Acta 821, 45-55. Winegar, R. A . , Monforte, J. A . , Suing, K . D . , Oloughlin, K . G . , Rudd, C. J. and Macgregor, J. T. (1996) Determination of tissue distribution of an intramuscular plasmid vaccine using P C R and in situ D N A hybridization. Hum. Gene Ther. 7(17), 2185-2194. Wive l , N . A . and Wilson, J. M . (1998) Methods of gene delivery. Hem.-One. Clinics N. Amer. 12(3), 483-513. Wolfert, M . A . , Schacht, E . J., Toncheva, V . , Ulbrich, K . , Nazarova, O. and Seymour, L . W . (1996) Characterization of vectors for gene therapy formed by self-assembly of D N A with synthetic block co-polymers. Hum. Gene Ther. 7(17), 2123-2133. Wong, F. M . P, Brooks, D . and Bal ly , M . B . (1999) Arch. Biophys., in press Wong, F. M . P., Reimer, D . L . and Bal ly , M . B . (1996) Cationic l ipid binding to D N A : characterization of complex formation. Biochem. 35, 5756-5763. Wong, T. K . , Nicolau, C . and Hofschneider, P. (1980) Appearance of (3-lactamase activity in animal cells upon liposome-mediated gene transfer. Gene 10, 87-94. Woodle, M . C. and Lasic, D . D . (1992) Sterically stabilized liposomes. Biochim. Biophys. Acta 1113, 171-199. Wu , G . Y . and W u , C. H . (1988a) Evidence for targeted gene delivery to Hep G2 hepatoma cells in vivo. Biochem. 27(3), 887-892. 187 Wu, G . Y . and Wu, C. H. (1988b) Receptor-mediated gene delivery and expression in vivo. J. Biol. Chem. 263(29), 14621-14624. X u , Y . H . and Szoka, F . C. (1996) Mechanism of D N A release from cationic l iposome/DNA complexes used in cell transfection. Biochem. 55(18), 5616-5623. Yoshimura, K . , Rosenfeld, M . A . , Nakamura, H . , Scherer, E . M . , Pavirani, A . , Lecocq, J. P. and Crystal, R. G . (1992) Expression of the human cystic fibrosis transmembrane conductance regulator gene in the mouse lung after in vivo intratracheal plasmid-mediated gene transfer. Nucl. Acids Res. 20(12), 3233-3240. Zabner, J., Fasbender, A . J., Moninger, T., Poellinger, K . A . and Welsh, M . J. (1995) Cellular and molecular barriers to gene transfer by a cationic lipid. J. Biol. Chem. 270(32), 18997-19007. Zauner, W. , Kichler, A . , Schmidt, W. , Mechtler, K . and Wagner, E . (1997) Glycerol and polylysine synergize in their ability to rupture vesicular membranes: a mechanism for increased transferrin-polylysine-mediated gene transfer. Exper. Cell Res. 232(1), 137-145. Zeisig, R., Shimada, K . , Hirota, S. and Arndt, D . (1996) Effect o f sterical stabilization on macrophage uptake in vitro and on thickness of the fixed aqueous layer of liposomes made from alkylphosphocholines. Biochim. Biophys. Acta 1285(2), 237-245. Zelphati, O. and Szoka, F. C. (1996) Intracellular distribution and mechanism of delivery of oligonucleotides mediated by cationic lipids. Pharmaceut. Res. 13(9), 1367-1372. Zelphati, O., Liang, X . W. , Hobart, P. and Feigner, P. L . (1999) Gene chemistry: Functionally and conformationally intact fluorescent plasmid D N A . Hum. Gene Ther. 10(1), 15-24. Zelphati, O., Uyechi, L . S., Barron, L . G . and Szoka, F. C. (1998) Effect o f serum components on the physico-chemical properties of cationic lipid/oligonucleotide complexes and on their interactions with cells. Biochim. Biophys. Acta - Lipids & Lipid Metabolism 1390(2), 119-133. Zenke, M . , Steinlein, P., Wagner, E . , Cotton, M . , Beug, H . and Birnstiel, M . L . (1990) Receptor-mediated endocytosis of transferrin-polycation conjugates: an efficient way to introduce D N A into hematopoietic cells. Proc. Natl. Acad. Sci. USA 57(10), 3655-3659. Zhang, Y . - P . , Reimer D . L . , Zhang, G . , Lee, P. H . and Bal ly , M . B . (1996) Self-assembling D N A - I i p i d particles for gene transfer. Pharmaceut. Res. 14(2), 190-196. Zheng, S., Zheng, Y . , Beissinger, R. L . and Fresco, R. (1992) Liposome-encapsulated hemoglobin processing methods. Biomater. Art. Cells Immobiliz. Biotech. 20(2-4), 355-364. Zheng, S., Zheng, Y . , Bessinger, R. L . and Fresco, R. (1994) Microencapsulation of hemoglobin in liposomes using a double emulsion, film dehydration/rehydration approach. Biochim. Biophys. Acta 1196(2), 123-130. Zhou, X . , Klibanov, A . and Huang, L . (1991) Lipophile polylysines mediate efficient D N A transfection in mammalian cells. Biochim. Biophys. Acta 1065, 8-14. 188 Zuidam, N . J. and Barenholz, Y . (1998) Electrostatic and structural properties of complexes involving plasmid D N A and cationic lipids commonly used for gene delivery. Biochim. Biophys. Acta 1368(1), 115-128. 189 Appendix Analytical Methods for the Analysis of Lipid-Based DNA Carriers* A.l Introduction The preparation of materials and the means used to analyze them have a direct effect on the physical and/or biological behavior of the lipoplex components, and therefore the methodology chosen for their study can influence the interpretation of their effects. While not every technique mentioned in this Appendix was used in the experimental section of this thesis, current general procedures used for the biophysical and biological characterization lipid-based D N A carriers, and their limitations, are described here to emphasize why the present work of basic physical characterization of the carriers is so crucial. Understanding how the lipid and D N A components interact with each other and their external environment is necessary for fully appreciating how they are interacting with cells or whole organisms. A.2 Materials Materials for the preparation and analysis of lipoplexes as used in this thesis were listed in Chapter 2. A.2.1 Additional Cationic Lipids D O T M A (N-[l-(2,3 dioleyloxy)-propyl]-N, N , N -trimethylammonium chloride) is available in combination with the neutral helper lipid dioleoylphosphatidylethanolamine (DOPE) in Lipofectamine™ reagent (Gibco-BRL, Grand Island, N Y ) . D O S P A (2,3-dioleyloxy-N[2-(sperminecarboxyamido)ethyl]-N,N-dimethyl-l-propanaminium trifluoroacetate) is available with D O P E in Lipofectin™ reagent (Gibco-BRL). D O T A P (dioleoyl-l,2-diacyl-3-trimethylammonium propane) can be obtained from Avanti Polar Lipids (Alabaster, A L ) or as methyl sulfate salt from Sigma (St. Louis, M O ) . D D A B * Adapted from: Wasan, E.K., Reimer, D.L., Harvie, P., Kong, S, Wong, F.M.P. and Bally, M.B. (1999) "Targeted Gene Transfer: a Practical Guide Based on Experience with Lipid-Based Plasmid Delivery Systems" in: Methods in MolecularMedicine Vol. 25 Drug Targeting, Ch.15. Francis, G.E. and Delgado, C , eds. Humana Press, Totowa, NJ. 190 (dimethyldioctadecyl ammonium bromide) can be purchased from Sigma or Avanti Polar Lipids, or formulated with D O P E in Lipofectace™ reagent (Gibco-BRL) . D M R I E (dimyristyloxypropyl-3-dimethyl-hydroxyethyl ammonium, G i b c o - B R L ) and D C - C h o i (cholesteryl 3p-[N-dimethylaminoethyl] carbamate are also available from Sigma. D O G S (dioctadecylamidoglycyl spermidine, Transfectam™) can be purchased from Promega (Madison, WI). D O D A C (dioleyldimethylammonium chloride) was generously provided by Inex Pharmaceuticals (Vancouver, B C , Canada). A.3 The Active Components A.3.1 Plasmids Gene transfection experiments often require large quantities of plasmid of high purity. Large amounts of D N A can be obtained by transforming a host cell such as the bacteria Escherichia coli (E. coli) with these small extrachromosomal pieces of circular D N A (the plasmids). It is important that the plasmid contain a gene that encodes for antibiotic resistance (such as ampicillin or tetracycline), or another selection marker, which is used to ensure that the bacteria growing in the presence of the drug retain the plasmid of interest. Following growth of the bacterial culture and subsequent amplification of the plasmid, the bacteria are lysed and the plasmid recovered by routine isolation techniques. A.3.1.1 Plasmid Isolation and Purification Routine techniques for culturing bacteria carrying plasmid can be found in standard manuals (Sambrook et al, 1989, Ausubel et al, 1997) and the methods used in this thesis were described in Chapter 2. There are many strains of bacteria now available, which have been genetically characterized for specific uses, and culture conditions may vary depending on the strain chosen. Plasmid is routinely recovered from bacteria by an alkaline lysis procedure, which lyses the bacterial cell while maintaining bacterial D N A attachment to the cell wall . This procedure enables subsequent precipitation of bacterial D N A and cellular debris, leaving a crude preparation enriched in plasmid. A plasmid 191 D N A purification kit provided by Qiagen was used in this thesis that utilizes the alkaline lysis method for harvesting, and anion exchange column chromatography for rapid purification. Detailed instructions are provided in the kit by the manufacturer, which are adequate for purification of laboratory-use plasmid D N A . There are several other methods available for purification of plasmid from the crude nucleic acid preparation once the bacterial cells have been lysed and a crude preparation of plasmid has been obtained. These include cesium chloride ultracentrifugation, gel filtration/size exclusion chromatography, anion exchange (used in the Qiagen kit mentioned above), and high performance liquid chromatography (HPLC) . They vary in the time, expense and equipment required and in the purity o f the plasmid produced. Purity is important to generate high levels of transfection in a reproducible manner. Potential contaminants include: bacterial genomic D N A , R N A , protein, endotoxin, chemical residues, trace metals and undesirable counterions (Marquet et al, 1995). Depending on the user's endpoint, concern regarding any of these contaminants w i l l vary. For example, i f the plasmid product is intended for human clinical trials, strict quality control over of all these factors, among others, must be addressed. Lesser concern is warranted for use in molecular biology studies, however, one should bear in mind that even cell culture transfection assays might be strongly affected by the quality o f the D N A (Weber et al, 1995). The cesium chloride gradient method using ultracentrifugation (Sambrook et al, 1989) yields high-quality D N A but requires considerable time to perform, is more difficult to scale up and requires the use of ethidium bromide, a known carcinogen. The ethidium bromide is typically removed from the purified plasmid by solvent extraction and the cesium chloride by dialysis. Techniques have been developed in which the cesium chloride and ethidium bromide can be removed from the D N A in one rapid step on a desalting column (see Materials). The impure D N A is first pretreated with 5 M guanidine thiocyanate and 5% P-mercaptoefhanol, 192 briefly heated at 65°C for 5 min, which allows complete removal of the C s C l upon column chromatography (Chang and Mattison, 1996). Anion exchange chromatography takes advantage of the affinity difference of plasmid D N A vs. proteinaceous cellular debris for the column matrix. A recent study comparing plasmid purified by anion exchange chromatography vs. C s C l density gradient showed that the two methods produced plasmid that was equivalent in transfection ability (Davis et al, 1996). In the studies detailed in this thesis, no obvious difference was noted between plasmids prepared by the two methods. (Usually plasmid was prepared by the anion exchange method.) Clearly, both methods yield highly purified D N A . The report by Davis did show, however, that the anion exchange method yielded plasmid with significantly higher levels o f endotoxin. Endotoxin is bacterial cell wall lipopolysaccharide, which is toxic to cells and may have an impact on the transfection of some cell lines. Its presence is also unacceptable in human gene therapy products due to the potential for anaphylactic reactions. Endotoxin is the agent responsible for inducing a state of shock in patients with systemic gram-negative bacterial infections. Because endotoxin behaves much like large plasmid on a size exclusion or anion exchange column due to its physical and chemical characteristics, it can be difficult to separate from plasmid D N A (Wicks et al, 1995). Endotoxin removal can be accomplished by several methods, such as treatment with Triton X - l 14 or polymixin B chromatographic resin. Triton X -114, when brought to its cloud point temperature of 20°C, w i l l solubilize the highly lipophilic endotoxin but not plasmid. Polymixin B binds with high affinity to endotoxin but not to D N A . Polymixin B is available bound to a polymer, which is used as a slurry to perform the extraction, usually by mixing with the crude D N A extract overnight or it can be used in a column. Unfortunately, these two methods are reported to not work well in the presence of C s C l (Davis et al, 1996). The anion exchange columns used in the Qiagen and Nucleobond plasmid preparation kits also remove some endotoxin, but not as much as by the aforementioned methods 193 (Cotton et al, 1994). Commercially available regeneratable resin with immobilized ligand can be used to remove endotoxin to trace levels. However, i f clinical use of the D N A is intended, methods must be in place to quantify levels of released ligand in the final product. Gel filtration or size exclusion isolation of plasmid from the crude nucleic acid preparation offers the advantages of good separation of plasmid from genomic D N A and R N A and a simple, inexpensive procedure (Bywater et al, 1983, Vo-Quang et al, 1985). The use of Sephacryl S-1000 Superfine matrix allows excellent separation of plasmid from R N A . When this separation is optimal, the use of RNase can be omitted from the isolation procedure, which is important for clinical considerations. This method isolates predominantly supercoiled D N A (covalently closed circular D N A ) (Raymond et al, 1988). This method of purification may be quite useful for studies investigating which conformation of the D N A is the most effective in transfecting cells; some studies have suggested that supercoiled D N A may be the most effective conformation (Weintraub et al, 1996). A disadvantage of using large size exclusion columns to effect complete separation of supercoiled D N A is that the sample may become quite diluted in the process, requiring an additional precipitation step and subsequent loss of product. Differential separation of plasmid forms (supercoiled, relaxed, linear) can also be accomplished using poly(ethylene glycol)6ooo (PEG) as the precipitating agent rather than ethanol or isopropanol. The technique is effective because a certain threshold concentration of P E G is required to precipitate nucleic acid of a given size. R N A and genomic D N A , therefore, can also be removed. Lower molecular weight nucleic acids require higher P E G concentrations to precipitate. To isolate plasmid, P E G (5-12%) is added to the crude nucleic acid preparation and incubated overnight, followed by centrifugation at 8000 x g for 5 min. Recovery is reduced i f the initial D N A concentration is low. Removal of the P E G , i f required, can be accomplished by D E A E cellulose chromatography, gel electrophoresis or C s C l density gradient (Lis, 1980). 194 Once the purified plasmid has been obtained, it is necessary to evaluate the purity of the sample and to determine the quantity and concentration obtained from the extraction procedure. Using a spectrophotometer, an optical density profile should be performed (between 230 and 340 nm). The profile should generate a peak at OD260 where D N A absorbs maximally. A simple measure of D N A purity is to determine the ratio of OD 26o/OD 2go. A pure D N A sample w i l l have a ratio of 1.8-2.0 (Ausubel et al, 1997). Ratios lower than 1.8 raise concerns about high levels of contamination with R N A or protein. A spectrophotometric scan (absorbance 230-340 nm) is also useful to evaluate protein contamination since protein scatters light much more effectively at 230 nm and the presence of protein may shift the absorbance of D N A to slightly greater than 260 nm (Laws and Adams, 1996). To determine the concentration of the D N A sample, an O D reading at 260 nm is taken and the concentration determined (one OD26o = 50 pg D N A / m l ) (Sambrook etal, 1989). Following quantification, the sample is dialyzed to remove impurities, such as salts and other contaminants, which can affect the behavior of the D N A in its interaction with the carrier and its transfection efficiency. For example the entire D N A sample may be dialyzed against 4 L of sterile distilled water overnight at 4°C. One hour after the start of dialysis 1 L of the water should be replaced with fresh sterile water. After the overnight dialysis, the dialysis bag is transferred to a fresh beaker of sterile water and dialyzed another 1 h at 4°C. The OD26o of the post-dialysis sample at an appropriate dilution in water is checked and concentration and recovery are calculated. For laboratory use, such as in vitro transfection, it is convenient to dilute this stock D N A solution to 1 mg/ml aliquots, which may be stored at - 2 0 ° C until use. A s an additional precaution, it is important to ensure that the plasmid that has been isolated, or that has been stored for an extended period, contains the expected insert. The plasmid should be digested with a restriction enzyme known to excise the insert and the resulting fragments evaluated following agarose gel electrophoresis. 195 A.3.1.2 Preparation of Radiolabeled Plasmid Radiolabeled plasmid can be utilized for studies involving animal pharmacokinetics, subcellular distribution during transfection, or biochemical/biophysical analyses in which one wishes to follow the fate of the plasmid D N A . For these purposes, 3 2 P - or 3H-labeled plasmid may be used. While 3 2 P provides excellent sensitivity, the 3 2P-labeled plasmid is more susceptible to radiation-induced damage and, i f prepared by nick translation, to exonuclease attack. 3 H-labeling of the plasmid provides a safer alternative (for both plasmid and user) with a high degree o f sensitivity for detection. The preparation of tritiated plasmid, described in Chapter 2, was modified from standard protocols due to the need to induce the bacteria to incorporate large quantities of radiolabeled nucleotide (for example, tritiated thymidine) during log phase growth. The radioactive nucleotide is incorporated not only into the plasmid, but also into the host genome, the latter o f which is a larger percentage of the total radiolabeled D N A . After plasmid isolation and purification, the 3 H-plasmid should be stored at -70°C to minimize radiation-induced breaks. It may be used for up to a year, providing that the quality of the plasmid is checked regularly (e.g. approximately every 2 months) by agarose gel electrophoresis. A.3.1.3 Preparation of Fluorescently Labeled Plasmid Fluorescently labeled plasmid can be quite useful for cellular uptake and distribution studies (Thierry et al, 1997). The main concern with fluorescently tagged D N A is that the presence of the dye molecules may interfere with the interactions between the D N A and the carrier, or the D N A and cellular components. O f equal importance, the binding of the dye to the D N A must be irreversible, so that the dye molecules do not dissociate from the D N A once internalized by the cell (Zelphati et al, 1999) and yet not interfere with transcription. A number of fluorescent DNA-intercalating dyes are available for D N A labeling, such as ethidium bromide, ethidium monoazide, the T O - P R O series (Figure A . l ) , various other cyanine dyes and many others (Haugland, 1996). For example, the membrane impermeable compound 196 Y O - Y O (Figure A.2) labels plasmid extremely well , giving a strong signal for either direct fluorescence measurements (Okomoto et al, 1994), fluorescence-activated cell sorting (FACS) (Kichler, 1997), or for microscopy of cells that have taken up the labeled plasmid D N A (Arndt-Jopvin and Jovin, 1989). This dye binds to D N A by intercalation (high affinity) and by ionic interactions (low affinity). Y O - Y O is supplied as a 1 m M solution in D M S O , which is added to the plasmid at approximately 1:100 dye:base pair ratio. A t this ratio, sensitivity is high and DNA/liposome complex formation is compromised very little. After incubation of plasmid and Y O - Y O dye at 50°C for 2-4 h, the plasmid is dialyzed against sterile distilled water at 4°C for 4-5 h with several changes of the water, or overnight. Dialysis removes the D M S O and any potential unbound Y O - Y O dye. After requantification of D N A concentration as described in the above sections, the plasmid is ready for use. Fluorescence can be read with excitation at 491 nm, and emission at 509 nm (Haugland, 1996). It is important to note, however, that a significant portion of the Y O - Y O is readily displaced from the D N A during preparation of lipid-based D N A delivery systems (Wong et al, 1999). The behavior of the Y O - Y O / D N A complex after cellular uptake is yet uncharacterized in this context. TO-PRO-1 Figure A . l DNA Binding Dye TO-PRO-1, Quinolinium, 4-[(3-methyl-2(3H> benzothiazolylidene)methyl] -1 - [3 -(trimethylammonio) propyl] -, diiodide YO-YO-1 Figure A.2 DNA Binding Dye YO-YO-1, Quinolinium, 1,1'-[1,3-propanediylbis[(dimethyliminio)-3,l-propanediyl]]bis[4-[(3-methyl-2(3H)-benzoxazolylidene)methyl]]-,tetraiodide 197 A.3.2 Cationic Liposomes for Gene Transfer Lipoplexes are taken up by most cells in vitro or in vivo much more efficiently than naked D N A . For preparation of gene transfer vehicles the liposomes are pre-formed, then mixed in a defined ratio with the nucleic acid for rapid, spontaneous formation of lipoplexes. Several methods are available for the preparation of liposomes, which are reviewed briefly in section 1.2.1, and other lipid-based D N A transfer formulations. Prepared, quality control-tested liposomes are available from commercial sources, as listed above, but they remain quite expensive. A method was described in Chapter 2 for the preparation of cationic liposomes by extrusion, a process that can be performed reproducibly in the laboratory with a minimum of equipment (a vortex mixer, a source of vacuum, and an extrusion device).- Liposomes may be radiolabeled with trace amounts (~1 pCi/ml) o f H - or 1 4C-cholesteryl hexadecyl ether (CHE) . Although it is well-established that C H E is an excellent lipid marker for following the fate of conventional liposomes in vivo because it is non-exchangeable and non-metabolizable, it is not clear whether this l ipid marker is adequate for following the fate of cationic lipoplexes used for gene transfer. For this reason, it may be preferable to use radiolabeled D O P E and/or radiolabeled cationic l ipid, particularly i f in vivo disposition is under study. The latter option is considerably more expensive than using radiolabeled C H E . A.3.2.1 Preparation of Cationic Liposomes The l ipid solution is reduced in volume under a stream of N 2 gas, and dried to a thin film for several hours under vacuum. The lipid films are hydrated by adding an aqueous solution while agitating, such as with a vortex mixer, to produce multilamellar vesicles ( M L V s ) . If D O P E is present, the hydrating solution should be of low ionic strength (<10 m M NaCl) or nonionic (such as 300 m M lactose or sucrose. The hydrating solution should be 0.22 pm filtered prior to use. Concentrated salt solutions (e.g. >100 m M NaCl) tend to cause rapid aggregation of DOPE-containing liposomes, or may prevent 198 proper hydration of the lipid film, required for liposome formation. This step should be performed at a temperature above the gel-to-liquid crystalline transition temperature of the lipids, however, i f D O P E is present hydration should be done on ice to avoid bilayer-to-hexagonal phase transition. The hydrated l ipid solution is then passed 10 times at room temperature through an extrusion device containing three polycarbonate membranes stacked together (0.08-1.0 um. The resulting large unilamellar vesicles ( L U V s ) have a mean diameter of 100-140 nm as determined by quasi-elastic light scattering (QELS) . This diameter can be modified by using membranes of different pore sizes. Final l ipid concentration may be determined by use of the radiolabeled l ipid and/or by an assay of phospholipid phosphorous. Liposomes should be stored at 4°C and used within 1 month. Extruded vesicles should not be frozen. Cationic l ip id /DOPE vesicles w i l l naturally begin to coalesce within 6 months to 1 year. A3.2.2 Formation of Lipoplexes Plasmid to be used for making lipoplexes is prepared in water. Liposomes ( L U V s but not M L V s ) and plasmid D N A should be centrifuged (7,000-10,000 x g for 2 min) prior to use to pellet any debris or aggregates. Large unilamellar liposomes and plasmid D N A w i l l not be pelleted by this procedure (Bally et al, 1997). Complexes are prepared with plasmid D N A and liposome components cooled on ice. Liposomes and D N A are diluted separately in dust-free tubes using the same type of solution that was used to prepare the liposomes. Typically, liposomes were used at a final dilution of 0.5-1 m M total lipid, and D N A concentrations of 5-500 pg/ml in a total volume of 0.2-10ml. Diluted plasmid D N A is pipetted gently into an equal volume of diluted liposomes, keeping the mixture on ice i f D O P E is present. There has been considerable debate in the field about the relative importance of the mixing rate during lipoplex preparation, but evidence is conflicting. In this laboratory it has been observed that flocculent aggregation is less likely i f D N A is added slowly to the liposomes and mixed immediately by gentle pipetting rather than by rapid vortex mixing. Other investigators promote the use of rapid mixing techniques to limit nucleation of particles (Lasic, 199 1998). Dust and debris in the components must also be scrupulously avoided. Flocculent aggregation may be related to the presence of impurities in the plasmid D N A , which can be avoided by careful, standardized D N A purification procedures and quality control testing prior to use. Complex formation is rapid (Zuidam and Barenholz, 1998); typically, complexes are allowed to form for about 30 minutes. The particle size of the resultant complexes varies with the ratio of l ipid to D N A , however, the minimum diameter is usually 200-400 nm, a size which is 2 to 5 times larger than the original 100 nm liposomes prior to D N A addition. The greatest aggregation occurs when the complex is approximately charge-neutral (Wasan et al, 1996). Complexes should be used within a few hours of preparation because transfection efficiency is reduced i f complexes are stored (Holland et al, 1996) and further aggregation can occur over time. A. 3.3 Characterization of Lipid-Based Gene Transfer Formulations Parameters which are believed to be important for lipoplex-mediated transfection include particle size and morphology, fusogenic behavior (with self and target membranes), D N A protection and stability in buffer or serum. Several assays for each of these parameters are discussed below. A. 3.3.1 Measuring Particle Size and Aggregation State The use of Q E L S and measurements of turbidity or sedimentation was addressed in Chapter 2 (sections 2.3.2.1 and 2.5.1). Particle size can also be estimated by electron microscopy. Freeze fracture E M requires fairly high l ipid concentrations (preferably >10 m M ) , which may not be what is used in actual working gene therapy formulations, or may be difficult to prepare (e.g. for those containing DOPE) . Additionally, the use of glycerol in freeze-fracture sample preparation can induce vesicle shrinkage due to osmotic forces. Despite these limitations, electron microscopy can provide further structural information. A n interesting variety o f structures such as long strands or globules or sheets have been described using this technique (Sternberg et al 1994, Gustaffson et al, 1995, Radler et al, 1997). Freeze-fracture E M of l i p i d / D N A particles has not been very 200 informative, presumably because the lipid in the particles is not in a typical bilayer configuration. Negative staining E M of l i p i d / D N A particles yields an image of spherical particles (Zhang et al, 1997) . Another microscopy technique that has been applied to l ipid carriers for plasmid D N A is cryogenic transmission E M ( C T E M ) , which has the advantage of rapid sample freezing, but it is technically challenging to perform. A detailed discussion of C T E M analysis may be found in Chapter 3. In addition to the basic sedimentation assay outlined in Chapter 2 (section 2.5.1.1), centrifugation can also be done with a density gradient in order to fractionate the lipoplexes into subpopulations. Useful gradient materials include sepharose, sucrose, dextrose, F ico l l , etc. This can be used, for example, to determine stability of the l i p i d - D N A complex under various experimental conditions such as when polyanions are added (Szoka et al, 1996, Ferrari et al, 1998) . Caution is warranted, however, to ensure that the gradient materials are not altering lipoplex size, structure or zeta potential (potential difference between lipoplex surface and the bulk phase, which is related to surface charge). Subpopulations within the lipoplex sample may vary between each other in their ability to transfect (Verma et al, 1998, Smith et al, 1998). Excess l ipid in the unfractionated sample may be responsible for much of the lipoplex toxicity but may be necessary for maximal transfection. This may be due to the need for an overall positive charge on the lipoplex for greatest uptake, but the resulting membrane perturbations may be partly responsible for the toxicity. A.3.3.2 Lipid Mixing Potential In order for l ipid carriers to be taken up by cells, there must be interaction between the cellular membrane and the particle. Lipoplexes are believed to be taken up primarily by endocytosis (Friend et al, 1996), with direct membrane fusion as a potential alternative route (Hui et al, 1996). Once inside the cell, the particle is thought to interact with anionic endocytic membranes in order to escape the endosome and avoid degradation (Szoka et al, 1996, Szoka et al, 1997). The D N A must reach either the cytoplasm 201 or the nucleus for activation. These processes imply that the particle itself must possess a certain critical degree of " l ip id mixing potential," or ability to interact or jo in with these cellular membranes. This property can be measured with model membrane systems, as described in Chapter 2 (section 2.5.4). One should bear in mind that the initial l ipid mixing potential of the liposome itself might be significantly reduced once it has bound plasmid D N A , or undergone further interactions in the course of cellular transfection. This issue is addressed in Chapter 6. Lipid-mixing assays developed in this thesis were based on earlier work using the resonance energy transfer technique (RET) with conventional liposomes as model membranes (Struck et al, 1981). Fluorescently labeled lipids, such as N B D - P E [N-(7-nitrobenz-2-oxa-l,3-diazol-4-yl)-l,2-dihexadecanoyl-5«-glycero-3-phosphoethanolamine)] and rhodamine-PE (Rh-P E : Lissamine-rhodamine phosphatidylethanolamine) are incorporated into the l ipid mixture used to make the liposomes or l i p i d / D N A particles. The fluorescence of N B D - P E molecules is quenched when close to rhodamine-PE molecules due to resonance energy transfer from N B D to rhodamine (Fung and Stryer, 1978, Stryer, 1978). When l ipid mixing occurs, the rhodamine component is physically moved away from the N B D by l ipid dilution, and the N B D then spontaneously emits its energy as fluorescence. N B D - P E and rhodamine-PE are used in low concentrations in the l ipid formulation (0.5-1% each), to minimize functional changes in the l ipid membrane or particle to be studied. To run a lipid-mixing assay, the fluorescent liposomes are mixed with nonfluorescent liposomes of otherwise identical composition (in a.ratio of 1:1 to 1:10). A ratio as high as 1:1 gives a strong fluorescence signal, which is useful when the effects of light scattering produce a noisy background. However, a ratio as low as 1:10 may be more sensitive because the dilution of the fluorescent lipids w i l l be greater upon membrane fusion, producing a bigger percent change in the fluorescence signal. To begin the assay, the baseline fluorescence (T 0) of a dilute suspension of liposomes or l i p i d / D N A particles (0.5-1 m M ) is measured over time with a 202 fluorometer. For the NBD/rhodamine system, excitation A= 465 run, emission X= 535 nm (Haugland, 1996), and an emission wavelength cutoff filter at about 530 nm should be used to minimize the effects of light scattering, which can be significant. I f no cutoff filter is available, controls must be included in which the experimental conditions are the same except no fluorescent lipids are used (Holland et al, 1996). These control values can then be subtracted from the experimental values as background. The next step is to add the component that induces l ipid mixing, such as a salt solution, D N A , another kind of liposome/membrane/cell while continuously recording the fluorescence (Tf). Continuous stirring is preferred, but not all fluorometers are equipped with such a device. If it not available, particular care must be taken to completely mix all components at each step prior to collecting data. Otherwise, a sample that exhibits a slow rate of l ipid mixing and one that is poorly mixed may be indistinguishable. The ratio of the inducing component to the l ipid vesicles or particles w i l l vary according to the experimental goals. Similar to anionic liposomes such as PS/PC (phosphatidylserine/phosphatidylcholine), which undergo rapid l ipid mixing in the presence of 1 m M C a 2 C l (Diizgiines et al, 1981), cationic liposomes such as D O D A C / D O P E or D O D A C / D O P C undergo rapid l ipid mixing when D N A is added. The reaction is complete within a few minutes (Stamatatos et al, 1988, Wasan et al, 1998). L i p i d / D N A particles or cationic liposomes containing D O P E also undergo rapid l ipid mixing with model membranes such as D O P S / D O P C (Xu and Szoka, 1996, Harvie et al, 1997). Finally, maximal l ipid mixing is simulated by the addition o f a nonionic detergent solution, such as 0.1-3% Triton X-100 or 35 m M O G P to disperse the lipids into monomers and/or mixed micelles. The fluorescence reading at this point is taken to be the maximal fluorescence ( T m a x ) . It can be difficult to solubilize lipoplexes with Triton X-100, but heating the samples to the cloud point of the detergent (about 100°C) (Schott, 1969) followed by vortex mixing helps. The fluorescence emitted after detergent addition w i l l only be the true maximum 203 i f the concentration of the fluorescent lipids is low enough that no resonance energy occurs, either within a micelle, between dispersed monomer lipids or between different micelles. The ideal detergent w i l l have no effect on the fluorescence properties (i.e. no quenching). These ideal conditions are difficult to meet experimentally. In the R E T experiments described in Chapters 5 and 6, the detergent concentration was titrated to maximize the decrease in R E T and to minimize quenching of N B D fluorescence. Some researchers prepare a mock total l ipid mixing control, in which one-tenth of the usual fluorescent l ipid (i.e. 0.05-0.1%) is included in the liposome preparation (Holland et al., 1996). Theoretically, the fluorescence of the mock fusion control should be the same as fluorescent plus nonfluorescent liposomes in a 1:10 ratio that exhibit 100% lipid mixing (and thus this value could be used as T m or factored into the equation below). However, in practical application, the efficiency of fluorescent lipid incorporation into liposomes tends to be variable, and the distribution of the fluorescent lipid in the two faces of the liposomal bilayer may not be equivalent. A n additional concern is that lipid exchange may occur. These factors are difficult to control. The average fluorescence readings at each step are then converted to percent of maximal l ipid mixing by the following formula: (Tf -T 0 ) / ( T m - T 0 ) . The initial fluorescence is subtracted from both T m and Tf in order to account for the variability in the actual fluorescence readings from sample to sample and between different batches of liposomes. A.3.3.3 Carrier-Cell Membrane Lipid Mixing Studies of the interaction between liposomes and model membranes have been used in an attempt to understand what factors are important in carrier-cell interactions. Vesicle aggregation, aqueous contents mixing or leakage and lipid mixing assays in the presence and absence of the fusion promoter (such as ions) are the most frequently examined phenomena. Contents mixing analysis demonstrates more clearly than lipid mixing that complete fusion of liposomes has occurred (Wilschut and Papahadjopoulos, 1979). More recently, l ipid mixing assays are being used to study the interaction between lipid-204 based D N A carriers and cellular membranes (Bailey and Cull is , 1997). Liposome aqueous contents mixing assays, while more definitive, are difficult to perform for DOPE-containing cationic liposomes because the combination o f fluorescent molecules typically used are salts o f strong acids, such as A N T S / D P X (l-aminonaphthalene-3,6,8-trisulfonic acid / N , N ' - r -xylylenebis(pyridinium bromide) (Smolarsky et al, 1977) or TbCf j /DPA (terbium chloride / dipicolinic acid) (Ellens et al, 1994). These compounds induce Hu phase formation of D O P E , resulting in the inability of the liposome to maintain a bilayer configuration. A. 3.3.4 DNA Protection For gene transfer to be effective, the D N A must reach its site of action intact. One reason that naked D N A is ineffective for gene transfer to most tissues [except muscle (Acsadi et al, 1991)] is that it is rapidly degraded by nucleases in the plasma compartment and interstitial spaces. Once taken up by the cells, D N A can also be degraded by intracellular nucleases in lysosomes. This terminal, degradative component of the endosomal system (Bainton et al, 1981) is the compartment into which transfected D N A is transported after gaining access to the cell (Friend et al, 1996). Intracellular nuclease-mediated degradation is detrimental to both in vitro and in vivo transfection efforts. It is therefore essential that the D N A be protected until it is released into the cytoplasm. This can be achieved by binding the D N A to cationic liposomes, by l i p i d - D N A particle (LDP) formation or by D N A condensation, such as with cationic polymers like polylysine, thus preventing accessibility of nucleases to labile bonds of the D N A structure. A n additional strategy is to include a component in the formulation that w i l l promote endocytic escape, such as a fusogenic lipid (DOPE) or a fusogenic peptide (Dzau et al, 1996). Below are described three complementary assays for determining the level of D N A protection, which are useful for comparing formulations. Relatively greater protection, however, does not necessarily correlate directly to increased competency in transfection (Harvie et al, 1998). For example, L D P s or lipoplexes prepared using cationic lipid and phosphatidylethanolamine (PE) are less stable than formulations 205 prepared with phosphatidylcholine (PC); however, PE-containing L D P s or lipoplexes are more effective for in vitro transfection of cells than PC-containing L D P s . The more important factor in transfection efficiency may be the relative fusogenicity of P E vs. P C . The P E headgroup may allow dissociation of the cationic l ipid from D N A more effectively than P C . It may be speculated that i f the D N A is too tightly bound, it cannot escape the carrier and reach the nucleus for transcription. A. 3.3.4.1 Fluorescent Dye Labeling of Plasmid D N A accessibility can be determined by how well the D N A binds fluorescent dyes. TO-PRO-1 (Figure A . l ) is a cyanine dye that fluoresces only when bound to nucleic acid. It is more sensitive for fluorescence detection than ethidium bromide and binds stably to the D N A . The relative degree of protection of the D N A can be quantified. TO-PRO-1 (final concentration 1 p M ) is added to formed complexes or particles containing 1 pg plasmid in 0.6 ml . A reference standard consisting of 1 pg D N A in water is also prepared and TO-PRO-1 added. Spectrofluorometric readings are taken after a 5 min incubation of the experimental samples and the reference standard with TO-PRO-1 [excitation A=509 nm, emission A=533 nm] (Haugland, 1996). TO-PRO-1 is light sensitive, so all experiments using this dye should be carried out under reduced light conditions. A value for comparison between samples can be calculated, called the "condensation index," CI = (I0-I) / 1 0 , where I is fluorescence of the sample in the presence of T O - P R O - 1 , and I 0 is fluorescence of the D N A reference (Wong et al, 1996). Under conditions where D N A is fully condensed, such as when bound to polylysine, the CI index approaches 1.0. TO-PRO-1 is reported to be membrane impermeant, therefore, the D N A could be protected from dye binding by the lipids and yet still not be condensed in the rigorous sense of the term (Bloomfield et al, 1991). A s an alternative, the "dye exclusion index," D E I = (If - L) / If x 100%, may be calculated. I; is the fluorescence of the l i p i d / D N A complexes or particles in the presence of TO-PRO-1 and If is the fluorescence of 206 the sample after solubilization with 35 m M O G P . The D E I is the percentage of the dye which is excluded from binding that potentially could bind. A33.4.2 DNase I Sensitivity DNase I is now routinely used to evaluate the stability of D N A against enzymatic degradation in the context of a carrier system to provide an indication of the potential in vivo usefulness of the carrier. The more protected and/or condensed that the plasmid D N A is by the components of the carrier, the less that DNase I w i l l be able to degrade the plasmid (Crook et al, 1997, Harvie et al, 1998). This method was described in Chapter 2 (section 2.6.2). Electrophoresis of the D N A sample on a 0.8% agarose gel w i l l then reveal the amount of degradation that has occurred. Well-protected D N A w i l l appear in a banding pattern that is similar to untreated plasmid, although a slightly higher proportion of the relaxed form is consistently observed. Degraded control plasmid w i l l appear as a smear of low M W fragments. A3.3.43 Serum Stability It has been demonstrated that in vitro transfection efficiency using most cationic lipids is reduced in the presence of fetal bovine serum (5-10%-typical tissue culture conditions) (Lewis et al, 1996). Serum contains proteins (albumin, alpha-2 macroglobulin, lipoproteins, IgG, complement, heparin, etc.) and nucleases that can interact with the lipoplexes and destabilize or degrade them (Zelphati et al, 1998). The stability of D N A in serum therefore determines in part the degree of protection afforded to the D N A by the l ipid carrier. To test serum stability, lipoplexes or L D P s (40 pg D N A / m l and 100 nmoles lipid/ml) containing 2 pg D N A total in 35-50 pi are added to 2 volumes of sterile water or 2 volumes of serum (such as normal mouse serum). The serum is centrifuged at 600 x g for 2 min prior to use to settle any precipitates. After mixing, incubation proceeds for 2 h at 37°C, sufficient time to fragment naked plasmid to small pieces, which can be detected by gel electrophoresis. For ease of handling, sample volumes are brought up to 300 pi with water. One volume of Tris-buffered phenol is added followed by vigorous vortex mixing. Centrifugation at 600 x g for 15 min 207 enhances separation of the sample into two phases. The upper aqueous phase containing the D N A is removed to a clean tube, avoiding the protein precipitate at the interface. The D N A is ethanol-precipitated by standard techniques, and evaluated by 0.8% agarose gel electrophoresis (80). The results are typically similar to those of the DNase I stability assay, where protected plasmid appears with its characteristic bands and degraded plasmid is seen as a low M W smear. A.3.4 Analysis of Cell Delivery The process by which the transgene is taken up into a cell and the gene product is expressed is complicated and not fully understood. Understanding the mechanisms involved in this process is imperative for the development of nonviral D N A delivery systems with high transfection efficiency. This has recently become the subject of intensive investigation and a number of critical barriers to transfection have now been identified (Zabner et al, 1995, Lasic and Templeton, 1996), including entry across the cell membrane, escape from the endosomal/lysosomal system, and nuclear entry. Endocytosis is believed to be the primary mode of cellular internalization of lipid-based D N A complexes. Receptor-mediated uptake is thought to be the major route of cellular entry with targeted vectors bearing peptide ligands (Wu and W u , 1998a, 1998b, Hart et al, 1995), carbohydrate ligands (Thurnher et al, 1994, Erbacher et al, 1996) or antibodies (Buschle et al, 1995, Ferkol et al, 1996). The first intracellular destination of the complex is the endosomal system. For D N A to be an effective transfection agent, it must escape the endocytic pathway, or it w i l l eventually be degraded when delivered to lysosomes (which contain nucleases and lipases). A n event that has been recently recognized to be of fundamental importance is that the plasmid D N A must dissociate from the carrier once it is internalized by the cell for transcription to occur. It is unclear whether this event occurs within the endosome itself, releasing the D N A in an unbound form into the cytoplasm (Lasic and Templeton, 1996, Zelphati and Szoka, 1996), or whether it occurs at the nuclear membrane (Friend et al, 1996) or in the nucleus itself (Labat-Moleur et al, 1996). Most evidence to date suggests that endosomal release is the most common mechanism. 208 The biochemical characteristics of the carrier play a critical role in the timing and efficiency of that release process. It is likely that cell-specific properties, yet be determined, also affect lipoplex uptake and breakdown. It was thought that dissociation of the nucleic acid and l ipid could occur within the endosome due to destabilization of the lipoplex under low p H conditions. The freed lipid would then have to destabilize the endosomal membrane sufficiently for the plasmid to diffuse into the cytoplasm, at a rate greater than that of plasmid degradation (endonuclease attack or base hydrolysis). This scenario seems rather unlikely because of the strength of the polyvalent binding between cationic lipids and D N A . Experimental evidence now suggests an alternative mechanism (Xu and Szoka, 1986, Harvie et al, 1998, Wasan et al, 1999). Lipoplexes are able to go through lipid mixing interactions with anionic l ipid model membranes. It is therefore possible that during these l ipid rearrangements and ion-pairing interactions, the D N A is released from the complex into the cytoplasm. This mechanism requires that some of the lipids in the carrier are fusogenic, i.e. readily able to form the nonbilayer intermediates generated during typical membrane-membrane fusion. The low p H o f the endosomal compartment is a condition that can induce D O P E to form nonbilayer structures (Hui et al, 1981, Siegal et al, 1986). Fusogenic peptides that change conformation at the p H of the endosomal compartment and disrupt endosomal membranes have been used with lipoplexes (Vogel et al, 1996). This mechanism assumes that the lipid-lipid associations during and immediately after the fusion event are stronger than those between the cationic l ipid and D N A , to allow D N A to escape. A third possible mechanism is that the entire lipoplex achieves endosomal release and approaches the nuclear pore. A t that point the D N A may be released or specific uptake mechanisms may permit the D N A to enter the nuclear pore complex. Confocal microscopy studies often show the lipid component segregated to the area just outside the nucleus while the polynucleotides are shown inside the nucleus (Zelphati et al, 1998). 209 Regardless, the plasmid D N A must gain entry to the nucleus in an intact form, where the cell's own machinery can eventually produce a functional protein. The mechanism by which plasmid enters the nucleus, whether by active transport or by diffusion, is not clear, however it is believed to be a major barrier to transfection (Zabner et al, 1995). Several investigators have utilized nuclear localization signal peptides (NLS) to facilitate nuclear targeting, with modest improvements in activity (Kaneda et al, 1989, Tomita et al, 1994). In order to elucidate the mechanism of gene transfer and expression, studies must be done to track the tissue, cellular and subcellular localization of both the active agent, in this case D N A , and the carrier components (cationic and associated helper lipids). In this section, a few techniques are described that are useful to gain a better understanding of the mechanisms of delivery of plasmid and carrier lipids. Many of the methods described herein can yield valuable information, however, they have limitations; new approaches to evaluate the pharmacokinetic and pharmacodynamic behavior of "gene medicines" must be developed. A.3.4.1 Plasmid and Lipid Delivery To achieve successful transfection, it is important that the D N A and the carrier are taken up by the target cells. Assessing the delivery process w i l l help determine whether uptake of a particular l i p i d / D N A formulation is a limiting factor in the transfection process. Analysis of D N A and l ipid delivery w i l l also help in understanding whether internalization is a limiting factor for a cell line that is difficult to transfect. A non-transfecting cell line may exhibit D N A uptake, but no transfection (Fasbender et al, 1997, Reimer et al, 1998). One type of carrier may work very wel l in transfecting cells and a similar one not at all. Moreover, the study of intracellular pharmacokinetics can reveal important characteristics about how these elements behave during the transfection process. Although there are several methods available to evaluate D N A and l ipid delivery, caution should be exercised in choosing the approach to be used. For example, many researchers have used plasmid that is bound to gold particles or tagged by fluorescent molecules. Although this 210 approach is convenient for microscopy studies, there is the potential for the tagging molecules themselves to interfere with the interaction between the D N A and the carrier, or the carrier/DNA complex and cellular components. Radioactive tags can provide excellent sensitivity for following the fate of plasmid or l ipid through the cell, but they must be non-exchangeable and chemically unreactive. Even so, one cannot distinguish between the radioactive parent compound and its chemical or metabolic degradation products. A s discussed above, the DNA/carrier complex is processed in endosomes, therefore, analysis of delivery to subcellular components may be required, which involves cellular fractionation. In addition, isolation of cell nuclei, which may be simpler to perform than more extensive subcellular fractionation, may be helpful in the study of the intracellular processing o f plasmid/carrier complexes. These procedures must be executed under well-controlled conditions to avoid contaminating the fractions (Tartakoff, 1987) in order to determine accurately the intracellular distribution of the carrier and D N A . A detergent-based lysis procedures for example, may be inappropriate due to its potential to alter the lipid component's distribution. Separation of cell nuclei and cationic lipoplexes can be difficult even with non-detergent methods. Highly specific markers for intracellular compartments (early endosomes, late endosomes, lysosomes, nuclei, etc) must be used to identify the fractions. . A combination of techniques is best employed to ensure an accurate assessment of intracellular distribution. A.3.4.1.1 Quantification of Plasmid Isolated from Transfected Cells There are several ways to determine the quantity of plasmid in cells following transfection. A simple procedure is presented here in which radiolabeled plasmid complexed to cationic lipids is used to transfect cells, followed by quantification of the plasmid at various timepoints (Reimer et al, 1997). Cells are grown at an appropriate concentration such that they w i l l be 70-80% confluent at the time of transfection. The serum-containing medium is removed and replaced with serum-free medium (80 pi in the case of 48 well plates). Lipoplexes in which the plasmid is 211 radiolabeled (see section 2.3.1.2) are added to the cells at the desired concentration, usually 0.5-1 pg D N A / w e l l in 20 pi of complexes. Incubation proceeds at 37°C for 1, 2, 3 or 4 h, at which time the experiment is terminated. It is important to include a parallel control at 4°C, where cellular metabolism and D N A uptake are inhibited. This avoids using metabolic inhibitor compounds to disrupt endosomal processing and serves as a valuable control to evaluate D N A internalization. The supernatant is removed and the cells are washed with serum-free medium that is pooled with supernatants, to determine the amount of plasmid that is not associated with the cells. The cells are then lysed in 300 pi of 10 m M T r i s - H C L (pH 8) containing 0.5% SDS, 3 m M M g C f i , 10 m M N a C l . The lysate is transferred to clean tubes and the wells rinsed with 300 p i lysis buffer, which is pooled with the corresponding samples. The amount of tritium in each sample is determined by scintillation counting and the amount of cell-associated D N A is calculated by subtracting the amount of D N A associated with the cells at 4°C from that obtained at 37°C. One of the concerns with this method is that it is difficult to determine whether the plasmid associated with the cells is in fact internalized. The plasmid may be only associated externally with the cell membrane or simply adhering nonspecifically to the tissue culture plate. To address this problem the cells can be briefly protease-treated prior to collection. Adherent cells are subjected to trypsin (0.25% at 4°C) and diluted with 400 p i HEPES-buffered saline (HBS, 25 m M H E P E S , 150 m M NaCl) . Cell-associated D N A is separated from non-associated by centrifugation. A density gradient is prepared from 1.077 and 1.022 (NycoPrep, Sigma). Samples are centrifuged at 400 x g for 30 min at ambient temperature. Cells are collected at the interface and cell-associated D N A evaluated for radioactivity. Other methods for quantifying the amount of D N A delivered to cells often require the use of D N A isolation procedures. Radiolabeled plasmid can be isolated from transfected cells using a modified Hirt lysis procedure (Hirt, 1967). This procedure selectively precipitates host 212 genomic D N A using a high salt concentration and cold SDS precipitation to precipitate genomic D N A , leaving plasmid in the supernatant for further purification and analysis. In the modified Hirt procedure, N a C l is added to the cell lysates (0.6% SDS, 10 m M E D T A ) to a final concentration of 1 M . The samples are incubated at 4°C overnight to promote precipitation. Centrifugation at 10, 000 x g for 15 min settles out a pellet of crosslinked genomic D N A and cell debris. The supernatant containing the plasmid is extracted using equal volumes of phenol/chloroform (1:1). The purified plasmid D N A is precipitated with 2 volumes of ethanol or 0.7 volumes isopropanol, centrifuged (10,000 x g for 30 min), and the resulting pellet dissolved in T E buffer (10 m M Tr i s -HCl , 1 m M E D T A , p H 8). The plasmid is then subjected to agarose gel electrophoresis. Since the plasmid has been radiolabeled, the bands on the gel can be excised and radioactivity measured by liquid scintillation counting. After transferring the gel slices to scintillation vials, the gel is melted in 500 pi of water or tissue solubilizer in a boiling waterbath. Subsequently the samples are mixed with the scintillation fluid and radioactivity counted. If the plasmid has not been radiolabeled, the amount of D N A taken up following transfection can be evaluated using "dot blot" analysis. Total D N A (genomic and plasmid) can be isolated from cells following transfection in vitro and in vivo using standard techniques (Sambrook et al, 1989). Alternatively, total D N A can be extracted from cells or tissue homogenates using D N A z o l following the manufacturer's instructions, or by the modified Hirt lysis procedure described above. DNazol is a guanidine detergent-based lysis solution, which selectively precipiates D N A (Cox, 1968). Following isolation and purification of D N A , the resolubilized D N A may be applied, to a dot blot apparatus for quantification. Plasmid D N A • 32 35 bound to the nylon membrane used in this procedure can be detected using J Z P , J J S or digoxigenin random-prime labeled plasmid (Sambrook et al, 1989, Ausubel et al, 1997). Plasmid is quantified by comparison with known standards after X-ray fi lm exposure or directly using a phosphoimager. Although the dot blot technique is excellent for quantification of 213 plasmid, a concern is that the probe w i l l hybridize to degraded fragments of the plasmid as well as intact plasmid, so that one cannot distinguish between the two. The polymerase chain reaction (PCR) is another routinely used option that eliminates concerns about using radiolabeled plasmid. P C R allows detection of very low levels of transfected D N A (Kawabata et al, 1995) after in vivo administration. It can be adapted to the analysis of histological sections of tissues (Osaka et al, 1996). Limitations to this technique include the requirement for stringent controls to avoid contamination and the inability to evaluate plasmid integrity. After in vivo lipoplex administration, plasmid can be quantified as described above by extraction from blood or tissues. The distribution, half-life and other pharmacokinetic parameters of the plasmid can then be calculated i f recovery is adequate. The simplest method to follow D N A in vivo is to utilize radiolabeled D N A and to assess radioactivity in tissue homogenates after transfection (Levy et al, 1996, Winegar et al, 1996, Reimer et al, 1999). One must be careful to account for the total dose in order to make analysis o f the data meaningful. Upon systemic administration, for example, larger organs such as the liver may take up more of the total dose, but may not account for the largest proportion when normalized to organ weight. A3.4.1.2 Analysis of Plasmid Integrity Delivery of the plasmid to the target cells is only useful i f the plasmid remains intact during the process and it is in a transcription-ready form after cellular internalization. A s indicated earlier, it is not clear yet which plasmid conformation(s) can be defined as the "active species." A n y analysis of plasmids isolated from cells or tissues should take this into consideration; bearing in mind that the extraction procedure itself could possibly affect D N A topology. In addition, it is unknown what quantity of D N A must reach the nucleus intact in order for transfection to be successful. The most widely used approach for the detection and analysis o f D N A is Southern analysis (Ausubel et al, 1997). This technique is useful because it allows not only detection of 214 the specific plasmid, but also assessment of degradation and conformation of the transfected plasmid. The method can be applied to D N A samples from in vitro or in vivo transfection. Southern analysis is sensitive enough to detect and quantify small amounts of plasmid D N A against a large background of genomic D N A (Porcher et al., 1992). In the procedure, total D N A is isolated from washed cells or frozen-powdered tissues by standard techniques, such as with T r i s /EDTA/SDS extraction (10 m M Tr i s -HCl , 0.1 m M E D T A - p H 8, 20 pg/ml pancreatic RNase, 0.5% SDS), followed by proteinase K treatment (100 pg/ml) and phenol/CHCl3 extraction of the D N A . A n alternative method involves lysing the cells with 7.5 volumes 6 M guanidine, 0.1 M sodium acetate, followed by ethanol precipitation then rehydration in T E buffer (Sambrook et al., 1989). Following D N A extraction, a restriction digest is performed, using one or more restriction enzymes that w i l l not cleave the plasmid. This digestion w i l l result in a uniform smear of genomic D N A upon gel electrophoresis and thus uninterrupted migration of the plasmid. After gel electrophoresis, the D N A fragments are transferred to a charged nylon membrane. D N A probes complementary to the sequence of interest are hybridized to the D N A on the membrane. The probes can be radioactively labeled, or bound to other molecules that can be easily detected, such as digoxigenin. Radiolabeled probes can detect as little as 0.1 pg of D N A on a Southern blot. The digoxigenin chemiluminescence system, available in kit form, has the advantage of not requiring radioactive materials. Detection is done via exposure of x-ray film, and sensitivity is on the order of 10 ng of plasmid. These high levels of sensitivity are important in order to detect low levels of D N A delivered to cells or tissues. If even greater sensitivity is required, P C R can be used to amplify plasmid from transfected cells or tissues (Sperisen et al, 1992) for a qualitative assessment of the presence of the transfected plasmid. P C R , like Southern analysis, requires specific intact D N A sequences for detection of the plasmid, so these methods may be more appropriate than 215 analyzing for the presence of radiolabeled D N A . P C R , however, can also amplify degraded plasmid; Southern blotting provides a more thorough analysis ( D N A quality as well as quantity). A.3.4.1.3 Analysis of Lipid Delivery The interaction of carrier l ipid molecules and cellular components is an essential factor in the delivery and release of the plasmid D N A prior to transcription. The carriers' physico-chemical properties that promote optimal transfection are still being investigated. A n analysis of carrier l ipid disposition within the cell, however, coupled with plasmid analysis and transfection data w i l l provide valuable information. Assessment of l ipid delivery to transfected cells or tissues can be accomplished in several ways, all of which suffer from serious problems. The lipid components of the carrier system can be labeled with fluorescent or radioactive moieties. Fluorescently labeled l ipid taken up by cells can be visualized by microscopy or quantified by flow cytometry (Schwedener et al, 1990). Radioactivity due to the labeled lipid in transfected tissue cultured cells can be assessed directly by scintillation counting. It should be noted that the distribution of labeled l ipid components, whether tagged by a radioisotope or a fluorescent marker, may not necessarily be representative of the intact l iposome/DNA complex or l i p i d / D N A particle. The labeled component must not affect the carrier assembly. When these systems are mixed with serum (in vitro or following intravenous injection) or are in contact with other membranes (cell plasma membrane) or lipid-rich proteins (lipoproteins) there is the potential for the tagged lipid component to exchange out of the lipoplex. The problem is compounded by issues related to metabolism and degradation. These concerns have been well addressed for liposomes used as carriers of small molecules where C H E has proven to be a non-exchangeable, non-metabolizable liposomal lipid marker. Importantly, C H E as the lipid marker only tracks the fate of the liposome, not the associated small molecule. For these reasons, one should always attempt to measure both l ipid component as well as associated active agents (Bally et al, 1993). 216 In the case of lipoplexes, the lipid markers that have been used are often the same as those used for conventional liposomes (i.e., C H E ) . This may not be appropriate. Radiolabeled lipids have been used effectively for investigating the biodistribution of lipid-based D N A carriers (Parker et al, 1997). Ideally, carrier cationic lipid and plasmid D N A quantification should be performed on the same tissues within in the same experiment. It may also be useful to analyze the fate of the neutral l ipid components of the carrier. Radiolabeled D O P E is available, for example. It may also be useful, in certain types of analyses such as pharmacokinetic and biodistribution studies, to label both the cationic and neutral components of the l ipid carrier. Without some kind of tag, however, the process of detection may be more complex due to the need to efficiently extract the carrier lipid(s) from cells or tissues prior to analysis. This may be difficult due to the presence of endogenous lipids. For cationic lipids such as D O T A P or D O D A C , quantification of tissue levels such as by H P L C analysis may be best performed by those specializing in l ipid analysis (Northern Lipids, Vancouver, B C , Canada). A.3.5 Expression of the Transgene The endpoint of any gene therapy strategy is expression of the transgene in the tissue of interest. Following successful cellular uptake of the gene carrier system, the transcription and translation machinery of the cell produces the protein product from the transgene (Figure 2.1). The number and specificity of the interactions that must occur between the gene carrier, the exogenous plasmid D N A , and components of the target cell in order for the gene product to be expressed is staggering. Control over the specificity, onset, and duration of this expression is still a major hurdle, largely due to the many variables involved. For example, without integration into the genome, which is currently achievable with retroviruses, expression w i l l necessarily be transient. Loss of the transgene w i l l inevitably occur as cell division progresses. However, in certain scenarios transient expression may be desirable, such as the treatment of cancer with toxin-producing genes. Restriction of expression can be achieved with tissue-specific gene promoters, but currently those in use for gene therapy 217 applications are few in number (Vile and Hart, 1994, Cowan et al, 1996, Gerloni et al, 1997). Inducible promoters are also an attractive option, giving the investigator the ability to turn expression of the gene on and off as desired or in response to a biological signal (Hallahan et al, 1995, Liang et al, 1996, Delort and Capecchi, 1996). These promoters, however, may not generate the high level o f expression achievable with nonspecific promoters, such as the cytomegalovirus ( C M V ) , simian virus (SV-40) or Rous sarcoma virus (RSV) promoters commonly in use today. Promoter inactivation by the host cell, which limits expression duration, is another major limitation to effective gene therapy. A.3.5.1 Evaluation of Transfection Using Reporter Genes "Reporter genes", also known as "marker genes," evaluate the ability o f a gene carrier system to deliver exogenous D N A and achieve expression in vitro or in vivo (Mount et al, 1996). These genes encode protein products, usually enzymes, for which there are simple biochemical assays, such as B-galactosidase, chloramphenicol acetyltransferase ( C A T ) and luciferase. The plasmids encoding green fluorescent protein (pGFP, available in several forms from Clontech) have recently become popular for fluorescence microscopy studies. G F P , a stable protein endogenous to the jellyfish Aequorea victoria, does not require a substrate to fluoresce (Heim et al, 1994). The disadvantage of G F P is that it is difficult to quantify the amount of G F P produced by measuring fluorescence. These four are currently the most commonly used reporter genes. They share the advantage of providing a simple measure of the effectiveness o f a gene therapy vector, in terms of expression of a product. Using any reporter gene, however, the actual level of expression may vary greatly between experiments, between cells on a tissue culture plate, or between animals within the same experiment. One must be cautious about how to make comparisons between groups or between different gene therapy studies that are utilizing reporter gene expression as a measure of success. The reporter genes are not interchangeable in a given vector; i.e., the level of expression driven by a particular promoter may vary according to the reporter gene (Mittal et 218 al, 1995). Another consideration to bear in mind is that all three o f these reporter gene products remain cytoplasmic, and may not be representative of the efficacy of a secreted therapeutic gene product, which could produce a "bystander effect." A.3.5.1.1 P-galactosidase (EC 3.2.1.23) P-gal is an enzyme found in many bacterial and eukaryotic cell types that converts o-nitrophenyl (3-D-galactoside to o-nitrophenol and D-galactose. It w i l l also convert 5-bromo-4-chloro-3-indoyl p-D-galactoside ("X-gal") to form a blue colored substance that can be assayed spectrophotometrically. The assay for the final product is reasonably rapid and simple to perform. The main advantage and use of P-gal as a reporter gene is that individual transfected cells can be seen under a light microscope due to the deposition of the blue product in the cytoplasm. X-ga l can be added to the cells during histochemical analysis or it can be incorporated into agar tissue culture plates (Hanna et al, 1984) for easy visualization of the transfected (blue) cells. Transfection efficiency and the percentage of cells expressing the transgene can be calculated from counting the blue vs. clear cells. Fluorescence-activated cell-sorting (FACS) analysis can also be adapted for analysis of cells transfected with P-gal (Srienc et al 1986). The disadvantage of P-gal is that quantification of the blue product, based on the optical density of cell lysates at 570 nm, is not very accurate or sensitive. There is also the concern that endogenous P-gal may interfere with interpretation of transfection results (Lai et al, 1994). In an in vivo transfection setting, typically one w i l l use P-gal as the reporter gene to determine the cell types being transfected and to estimate efficiency. This is coupled with the use of a different reporter gene, such as luciferase or C A T as described below, to perform the actual quantification of the amount of transgene product produced (Jiao et al, 1992, Yoshimura et al, 1992). When using P-gal as a histological marker it is important to control for the background by including tissues obtained from animals injected with a p-gal-null plasmid. 219 A3.5.1.2 Chloramphenicol Acetyltransferase C A T (EC 2.3.1.28) is a bacterial enzyme that catalyzes acetylation of chloramphenicol with the use of acetyl-S-Coenzyme A (CoA). When the bacterial gene is inserted into a plasmid expression vector appropriate for transfecting eukaryotic cells, the C A T enzyme can be quantified by measuring its activity (Gorman et al, 1992). One assay for enzyme activity in cell or tissue homogenates employs a radioactive substrate, either 1 4C-chloramphenicol, or 1 4 C-acetyl from labeled acetyl C o A (Sleigh, 1986). Tissues must first be heat-inactivated and treated with paraoxan (diethyl p-nitrophenyl phosphate) to avoid contributions from other acetylating enzymes present in the tissue. Analysis is performed by liquid scintillation counting of the radioactive product after mixed-xylene extraction. The activity of the cells or tissue expressing C A T is compared to the known activity of a C A T standard. This assay provides better sensitivity than P-gal, however, it involves the use of radioactive materials and the assay procedure is lengthy and labor-intensive. When reporting C A T activity, one should convert the transferred cpm of radioactivity to units of C A T activity based on the standard. It has also been argued that units should be converted into mg of product to obtain a value that can be compared between different research laboratories. A3.5.1.3 Luciferase (EC 1.13.12.7) Luciferase is a light-producing enzyme from the American firefly Photinus pyralis. The luciferase gene has been cloned and inserted into vectors for expression in bacterial or eukaryotic cells (de Wet et al, 1985). Luciferase has no mammalian endogenous counterpart, so there is no background interference (Chapman et al, 1992). Luciferase utilizes A T P as a substrate to produce light, which can then be measured in a simple luminometer (de Wet et al, 1987, Brasier et al, 1989). Less accurately, the light emission can be measured by scintillation counting or by exposure to photographic fi lm (Kricka et al, 1991). The assay to measure luciferase activity is described in Chapter 2 (section 2.7.4). The more sophisticated luminometers can analyze the kinetics of the reaction and assess luciferase activity based on the integrated area of steady-state light emission vs. time. 220 A.3.5.2 Gene vs. Message vs. Product While we are necessarily most interested in the final expression of the transgene, its time of onset and duration of action, it may be necessary to perform more fundamental investigations when expression is less than optimal. It is important to assess D N A delivery to the target cell population. It has been noted that while plasmid delivery is necessary for transfection, it is not sufficient in itself. The interaction between the carrier and the plasmid D N A is important for not only delivery and protection of the D N A , but also release within the appropriate intracellular compartment. Thus, the study of the intracellular uptake, distribution, and degradation of both plasmid and carrier are useful in the effort to improve our understanding of nonviral D N A transfection of cells. While the discussion here has focused on lipid carriers, the same principles apply to other types of nonviral carriers. Analysis o f the disposition of targeted vectors and multi-component delivery systems may be more complex, but no less worthwhile. Another issue is the transcription of the delivered gene into m R N A , in particular the time of its appearance in the target cells and the stability of the message. This may be influenced not only by the properties of the transfected cell, but also by the sequence itself. Northern blot analysis (Sambrook et al, 1989) can reveal when transcription of the transferred gene is beginning and how long it continues, whether less than 24 h or for months. Analysis of the gene product, the marker or therapeutic protein of interest, is the ultimate step and care must be taken to chose a detection method that is both sensitive and free of background. When D N A delivery, carrier delivery and stability, m R N A expression and protein expression data are examined together, this can give information about which steps in the cellular process leading to expression are rate-limiting. The greatest limitation to improving nonviral D N A delivery systems may be in not knowing where the bottleneck is in the process from plasmid uptake to functional protein production. 221 A .4 Summary Gene therapy strategies are being aggressively tested in the clinic, an indication of the immense hope being placed on this treatment approach. It is clear, however, that the pharmaceutical development of drugs consisting of oligonucleotides or plasmids has not been fully realized yet. Furthermore, our understanding of these agents as therapeutic molecules is very limited. It is believed that carriers w i l l be required in order to facilitate efficient D N A delivery to target cells. Issues that must be addressed in the development of D N A as a drug are similar to those encountered in the development of small molecules. Interest may arise from a novel mechanism of action and/or an activity that is better than that achieved with drugs already known to be active in the particular target disease. A pharmaceutically viable formulation must be defined, one that is amenable to manufacturing processes that can be validated, resulting in a product that is suitable for use in humans. It is anticipated that Phase I studies, designed to assess toxicity, and Phase II studies, designed to assess efficacy, w i l l progress in the absence of established mechanism(s) of action. However, it w i l l be critical to the pharmaceutical development of D N A as a drug to define the pharmacokinetic and pharmacodynamic characteristics of the D N A . It is l ikely that the carrier formulations used to deliver the D N A w i l l also be considered active drug components, therefore, methodologies must be further developed in order to accurately and reproducibly measure the biological fate of D N A delivery systems. 222 

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