Organelle-Specific Pharmaceutical Nanotechnology

ORGANELLE-SPECIFIC PHARMACEUTICAL NANOTECHNOLOGY Edited by Volkmar Weissig Gerard G. M. D’Souza A JOHN WILEY & SONS, I...

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ORGANELLE-SPECIFIC PHARMACEUTICAL NANOTECHNOLOGY Edited by

Volkmar Weissig Gerard G. M. D’Souza

A JOHN WILEY & SONS, INC., PUBLICATION

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ORGANELLE-SPECIFIC PHARMACEUTICAL NANOTECHNOLOGY

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ORGANELLE-SPECIFIC PHARMACEUTICAL NANOTECHNOLOGY Edited by

Volkmar Weissig Gerard G. M. D’Souza

A JOHN WILEY & SONS, INC., PUBLICATION

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Copyright © 2010 by John Wiley & Sons, Inc. All rights reserved Published by John Wiley & Sons, Inc., Hoboken, New Jersey Published simultaneously in Canada No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, recording, scanning, or otherwise, except as permitted under Section 107 or 108 of the 1976 United States Copyright Act, without either the prior written permission of the Publisher, or authorization through payment of the appropriate per-copy fee to the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, (978) 750-8400, fax (978) 750-4470, or on the web at www.copyright.com. Requests to the Publisher for permission should be addressed to the Permissions Department, John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, (201) 748-6011, fax (201) 748-6008, or online at http://www.wiley.com/go/permission. Limit of Liability/Disclaimer of Warranty: While the publisher and author have used their best efforts in preparing this book, they make no representations or warranties with respect to the accuracy or completeness of the contents of this book and specifically disclaim any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives or written sales materials. The advice and strategies contained herein may not be suitable for your situation. You should consult with a professional where appropriate. Neither the publisher nor author shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. For general information on our other products and services or for technical support, please contact our Customer Care Department within the United States at (800) 762-2974, outside the United States at (317) 572-3993 or fax (317) 572-4002. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic formats. For more information about Wiley products, visit our web site at www.wiley.com. ISBN 978-0-470-63165-2 Library of Congress Cataloging-in-Publication Data is available. Printed in the United States of America 10

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CONTENTS

Preface

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Contributors

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1. An Introduction to Subcellular Nanomedicine: Current Trends and Future Developments

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Gerard G. M. D’Souza and Volkmar Weissig

2. Delivery of Nanonsensors to Measure the Intracellular Environment

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Paul G. Coupland and Jonathan W. Aylott

3. Cytoplasmic Diffiusion of Dendrimers and Dendriplexes

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Alexander T. Florence and Pakatip Ruenraroengsak

4. Endocytosis and Intracellular Trafficking of Quantum Dot– Ligand Bioconjugates

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Tore-Geir Iversen, Nadine Frerker, and Kirsten Sandvig

5. Synthesis of Metal Nanoparticle-Based Intracellular Biosensors and Therapeutic Agents

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Neil Bricklebank

6. Subcellular Fate of Nanodelivery Systems

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Dusica Maysinger, Sebastien Boridy, and Eliza Hutter

7. Intracellular Fate of Plasmid DNA Polyplexes

123

Kevin Maier and Ernst Wagner

8. Intracellular Trafficking of Membrane Receptor-Mediated Uptake of Carbon Nanotubes

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Bin Kang and Yaodong Dai

9. Real-Time Particle Tracking for Studying Intracellular Transport of Nanotherapeutics

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Clive Chen and Junghae Suh

10. Tracking Intracellular Polymer Localization Via Fluorescence Microscopy

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Simon C. W. Richardson v

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11. Can QSAR Models Describing Small-Molecule Xenobiotics Give Useful Tips for Predicting Uptake and Localization of Nanoparticles in Living Cells? And If Not, Why Not?

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Richard W. Horobin

12. Self-Unpacking Gene Delivery Scaffolds

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Millicent O. Sullivan

13. Cellular Trafficking of Dendrimers

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Yunus Emre Kurtoglu and Rangaramanujam M. Kannan

14. Endolysosomolytically Active pH-Sensitive Polymeric Nanotechnology

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Han Chang Kang and You Han Bae

15. Uptake and Intracellular Dynamics of Proteins Internalized by Cell-Penetrating Peptides

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Arwyn T. Jones

16. Cargo Transport by Teams of Molecular Motors: Basic Mechanisms for Intracellular Drug Delivery

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Melanie J. I. Müller, Florian Berger, Stegan Klumpp, and Reinhard Lipowsky

17. The Potential of Photochemical Internalization (PCI) for the Cytosolic Delivery of Nanomedicines

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Kristian Berg, Anette Weyergang, Anders Høgset, and Pål Kristian Selbo

18. Peptide-Based Nanocarriers for Intracellular Delivery of Biologically Active Proteins

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Seong Loong Lo and Shu Wang

19. Organelle-Specific Pharmaceutical Nanotechnology: Active Cellular Transport of Submicro- and Nanoscale Particles

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Galya Orr

20. Subcellular Targeting of Virus-Envelope-Coated Nanoparticles

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Jia Wang, Mohammad F. Saeed, Andrey A. Kolokoltsov, and Robert A. Davey

21. Mitochondria-Targeted Pharmaceutical Nanocarriers

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Volkmar Weissig and Gerard G.M. D’Souza

22. Cell-Penetrating Peptides for Cytosolic Delivery of Biomacromolecules

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Camilla Foged, Xiaona Jing, and Hanne Moerck Nielsen

23. Therapeutic Nano-object Delivery to Subdomains of Cardiac Myocytes

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Valeriy Lukyanenko

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CONTENTS

24. Design Parameters Modulating Intracellular Drug Delivery: Anchoring to Specific Cellular Epitopes, Carrier Geometry, and Use of Auxiliary Pharmacological Agents

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Silvia Muro and Vladimir R. Muzykantov

25. Uptake Pathways Dependent Intracellular Trafficking of DNA Carrying Nanodelivery Systems

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Ikramy A. Khalil, Yuma Yamada, Hidetaka Akita, and Hideyoshi Harashima

26. Cellular Interactions of Plasmon-Resonant Gold Nanorods

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Qingshan Wei and Alexander Wei

27. Quantum Dot Labeling for Assessment of Intracellular Trafficking of Therapeutically Active Molecules

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Diane J. Burgess and Mamta Kapoor

Index

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PREFACE

Pharmaceutical nanotechnology as applied to the level of cell organelles, that is, to the subcellular level, is emerging as a new field in biomedical research. While efforts aiming at the targeted delivery of biologically active molecules date back to the early 1970s, the successful delivery of such agents to particular cell organelles has only in recent times gained broader recognition. Without exaggeration it can be stated that the subcellular organelle-specific targeting of therapeutic and diagnostic agents has become the new frontier for drug delivery. A variety of pharmaceutical nanocarrier platforms like liposomes, carbon nanotubes, quantum dots, micelles, and dendrimers have undergone testing for their ability to control the subcellular disposition of drugs. Potential improvements in therapy through the use of organelle-targeted nanocarriers have been demonstrated. Two major strategies are the basis for achieving organelle-specific targeting of pharmaceutical nano carriers. The first is based on the inherent predisposition of the nanocarrier for a particular cellular compartment and the second is based on an appropriate surface modification of the carrier with organelle-specific ligands. Despite encouraging preliminary findings, or maybe even because of them, many questions begin to emerge or still remain unanswered. Principally, our knowledge about the cellular internalization of nanocarriers and about the impact of intracellular morphology and dynamics on the fate and disposition of nanocarriers is very limited. Do nanocarriers remain intact upon cell entry and subsequent disposition? Naturally, significant differences between solid nanoparticles and liposomes can be expected. What is the impact of size and architecture of nanoparticles on their intracellular trafficking and distribution? Can the drug release from nanocarriers be controlled in a timely and spatial manner? Can or do cells actively transport nanocarriers in cell-membrane-derived vesicles? Can intracellular membrane trafficking be utilized for organelle-specific drug delivery? How do nanocarriers interact with different organelles? Do liposomal phospholipids become part of cellular vesicles? If so, what then is the fate of the lipophilic drugs originally incorporated into liposomal membranes? Can liposomes fuse with organellar membranes such as the mitochondrial outer or inner membrane? How do nonbiodegradable nanocarriers affect cellular transport and metabolism? Our ability to answer those and many more questions largely depends on technological advances in imaging technology. It can be hoped that improvements in real-time fluorescence confocal microscopy of ix

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live cells as well as the emergence of new imaging techniques together with the design of novel nanoprobes will help to more satisfactorily answer some of the questions raised above. This book is the result of our efforts to bring together the best of current knowledge in this new and emerging field of subcellular pharmaceutical nanotechnology. All chapters were written by leading experts in their particular fields, and we are extremely grateful to them for having spent part of their valuable time to contribute to this book. It is our hope that together we have succeeded in providing an essential source of state-of-the-art knowledge for any investigator, young and seasoned alike, whose research area involves the application of new nanotechnology to the subcellular level. VOLKMAR WEISSIG Glendale, Arizona GERARD G. M. D’SOUZA Boston, Massachusetts

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CONTRIBUTORS

Hidetaka Akita, Laboratory for Molecular Design of Pharmaceutics, Faculty of Pharmaceutical Sciences, Hokkaido University, Hokkaido, Japan and CREST, Japan Science and Technology Agency (JST), Japan Jonathan W. Aylott, School of Pharmacy, University of Nottingham, Nottingham, United Kingdom You Han Bae, Department of Pharmaceutics and Pharmaceutical Chemistry, University of Utah, Salt Lake City, Utah Kristian Berg, Department of Radiation Biology, Institute for Cancer Research, Norwegian Radium Hospital, Oslo University Hospital, Montebello, Oslo, Norway Florian Berger, Department of Theory & Bio-Systems, Max Planck Institute of Colloids and Interfaces, Potsdam, Germany Sebastian Boridy, Department of Pharmacology & Therapeutics, McGill University, Montreal, Quebec, Canada Neil Bricklebank, Faculty of Health and Wellbeing, Biomedical Research Center, Sheffield Hallam University, Sheffield, United Kingdom Diane Burgess, School of Pharmacy, University of Connecticut, Storrs, Connecticut Clive Chen, Department of Bioengineering, Rice University, Houston, Texas Paul G. Coupland, School of Pharmacy, University of Nottingham, Nottingham, United Kingdom Gerard G. M. D’Souza, Department of Pharmaceutical Sciences, Massachusetts College of Pharmacy and Health Sciences, Boston, Massachusetts Yaodong Dai, Nanjing University of Aeronautics and Astronautics, Nanjing, Peoples Republic of China Robert A. Davey, Department of Microbiology and Immunology, University of Texas Medical Branch, Galveston, Texas Alexander T. Florence, School of Pharmacy, University of London, London, United Kingdom xi

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Camilla Foged, Department of Pharmaceutics and Analytical Chemistry, Faculty of Pharmaceutical Sciences, University of Copenhagen, Copenhagen, Denmark Nadine Frerker, Centre for Cancer Biomedicine, Faculty Division, Norwegian Radium Hospital, University of Oslo and Department of Biochemistry, Institute for Cancer Research, Norwegian Radium Hospital, Oslo University Hospital, Oslo, Norway Hideyoshi Harashima, Laboratory for Molecular Design of Pharmaceutics, Faculty of Pharmaceutical Sciences, Hokkaido, University, Hokkaido, Japan and CREST, Japan Science and Technology Agency (JST), Japan Anders Hgset, PCI Biotech AS, Oslo, Norway Richard W. Horobin, Division of Integrated Biology, FBLS, The University of Glasgow, Glasgow, Scotland, United Kingdom Eliza Hutter, Department of Pharmacology and Therapeutics, McGill University, Montreal, Quebec, Canada Tore-Geir Iversen, Centre for Cancer Biomedicine, Faculty Division, Norwegian Radium Hospital, University of Oslo, and Department of Biochemistry, Institute for Cancer Research, Norwegian Radium Hospital, Oslo University Hospital, Oslo, Norway Xiaona Jing, Department of Pharmaceutics and Analytical Chemistry, Faculty of Pharmaceutical Sciences, University of Copenhagen, Copenhagen, Denmark Arwyn T. Jones, Welsh School of Pharmacy, Cardiff University, Cardiff, Wales, United Kingdom Bin Kang, Nanjing University of Aeronautics and Astronautics, Nanjing, Peoples Republic of China Han Chang Kang, Department of Pharmaceutics and Pharmaceutical Chemistry, University of Utah, Salt Lake City, Utah Rangaramanujam M. Kannan, Department of Chemical Engineering and Materials Science, NICHD Perinatology Research Branch, Wayne State University, Detroit, Michigan Mamta Kapoor, School of Pharmacy, University of Connecticut, Storrs, Connecticut Ikramy A. Khalil, Laboratory for Molecular Design of Pharmaceutics, Faculty of Pharmaceutical Sciences, Hokkaido, University, Hokkaido, Japan and CREST, Japan Science and Technology Agency (JST), Japan Stefan Klumpp, Department of Theory & Bio-Systems, Max Planck Institute of Colloids and Interfaces, Potsdam, Germany

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CONTRIBUTORS

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Andrey A. Kolokoltsov, Department of Microbiology and Immunology, University of Texas Medical Branch, Galveston, Texas Yunus Emre Kurtoglu, Department of Chemical Engineering and Materials Science, NICHD Perinatology Research Branch, Wayne State University, Detroit, Michigan Reinhard Lipowsky, Department of Theory & Bio-Systems, Max Planck Institute of Colloids and Interfaces, Potsdam, Germany Seong Loong Lo, Institute of Bioengineering and Nanotechnology, Singapore Valeriy Lukyanenko, Medical Biotechnology Center, University of Maryland Biotechnology Institute, Baltimore, Maryland Kevin Maier, Pharmaceutical Biotechnology, Munich Center for SystemBased Drug Research, and Center for NanoScience, Ludwig-MaximiliansUniversität, Munich, Germany Dusica Maysinger, Department of Pharmacology & Therapeutics, McGill University, Montreal, Quebec, Canada Melanie J. I. Müller, Department of Theory & Bio-Systems, Max Planck Institute of Colloids and Interfaces, Potsdam, Germany Silvia Muro, Center for Biosystems Research, University of Maryland Biotechnology Institute and Fischell Department of Bioengineering, University of Maryland, College Park, Maryland Vladimir R. Muzykantov, Department of Pharmacology and Targeted Therapeutics Program of Institute of Translational Medicine and Therapeutics, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania Hanne Moerck Nielson, Department of Pharmaceutics and Analytical Chemistry, Faculty of Pharmaceutical Sciences, University of Copenhagen, Copenhagen, Denmark Galya Orr, Chemical and Materials Sciences Division, Pacific Northwest National Laboratory, Richland, Washington Simon C. W. Richardson, University of Greenwich School of Science, Kent, England Pakatip Ruenraroengsak, Lung Cell Biology, Respiratory Medicine, National Heart and Lung Institute, Imperial College, London, United Kingdom Mohammad F. Saeed, Department of Microbiology and Immunology, University of Texas Medical Branch, Galveston, Texas Kirsten Sandvig, Centre for Cancer Biomedicine, Faculty Division, Norwegian Radium Hospital, University of Oslo, and Department of Biochemistry,

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Institute for Cancer Research, Norwegian Radium Hospital, Oslo University Hospital, Oslo, Norway Pål Kristian Selbo, Department of Radiation Biology, Institute for Cancer Research, Norwegian Radium Hospital, Oslo University Hospital, Montebello, Oslo, Norway Junghae Suh, Department of Bioengineering, Rice University, Houston, Texas Millicent O. Sullivan, Department of Chemical Engineering, University of Delaware, Newark, Delaware Ernst Wagner, Pharmaceutical Biotechnology, Munich Center for SystemBased Drug Research, and Center for NanoScience, Ludwig-MaximiliansUniversität, Munich, Germany Jia Wang, Department of Microbiology and Immunology, University of Texas Medical Branch, Galveston, Texas Shu Wang, Institute of Bioengineering and Nanotechnology, and Department of Biological Sciences, National University of Singapore, Singapore Alexander Wei, Department of Chemistry, Purdue University, West Lafayette, Indiana Qingshan Wei, Department of Chemistry, Purdue University, West Lafayette, Indiana Volkmar Weissig, Department of Pharmaceutical Sciences, Midwestern University College of Pharmacy, Glendale, Arizona Anette Weyergang, Department of Radiation Biology, Institute for Cancer Research, Norwegian Radium Hospital, Oslo University Hospital, Montebello, Oslo, Norway Yuma Yamada, Laboratory for Molecular Design of Pharmaceutics, Faculty of Pharmaceutical Sciences, Hokkaido University, Hokkaido, Japan and CREST, Japan Science and Technology Agency (JST), Japan

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CHAPTER 1

An Introduction to Subcellular Nanomedicine: Current Trends and Future Developments GERARD G. M. D’SOUZA Department of Pharmaceutical Sciences, Massachusetts College of Pharmacy and Health Sciences, Boston, Massachusetts

VOLKMAR WEISSIG Department of Pharmaceutical Sciences, Midwestern University College of Pharmacy Glendale, Glendale, Arizona

Drug therapy is based largely on the paradigm that an ideal drug will selectively exert a desired pharmacological activity free from negative side effects to modulate either the symptoms or the underlying biochemical cause of a disease to provide a benefit to the patient. In order to have such selective action, the drug molecule should ideally interact with only the disease-associated biochemical pathway but have no activity with respect to any normal biochemical pathway. This principle was famously explored by Paul Ehrlich in his search for agents with selective toxicity toward bacteria. Ehrlich’s work is widely accepted to have given rise to the concept of the ideal drug molecule as a “magic bullet,” a term that he used for the first time in his Harben Lectures [1]. Finding such selective molecules is relatively easy when there are significant differences between the disease-causing process and normal human biochemical pathways, as in the case of infectious diseases. Not surprisingly, in the decades since Ehrlich’s work infectious diseases have become much easier to treat but it does bear consideration that the lack of activity in nondisease cells is dose dependent and not absolute. Most drugs that are considered to be selectively toxic to invading pathogens are in fact toxic to human cells as well but just at higher doses. However, given that the new challenges in drug therapy lie in the treatment of diseases associated with malfunctions Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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of normal human biochemical pathways in certain tissues, the concept of the magic bullet perhaps needs to be redefined or at least clarified. In the infectious disease example it is essential to understand that the so called magic bullet did not necessarily have to home in on the disease agent but could in fact accumulate to the same level in both host and pathogen cells. Just as long as the agent was toxic only to the pathogen it was considered by many to be a magic bullet. In fact, Ehrlich’s Nobel Prize Lecture from December 11, 1908 speaks of “outlining the principles of selective toxicity” rather than selective accumulation. Therefore one could argue that for a molecule to be a magic bullet, it does not have to selectively accumulate at its intended site of action but just that it should not exert its action anywhere but at that intended site of action. This is different from what is now referred to as drug targeting. The term “targeting” is most often meant to imply that the molecule is in some way able to selectively accumulate at an intended site of action and that the selective accumulation is associated with its selective action as a magic bullet. Unfortunately (perhaps due to the widespread use of the noun target to describe a potential molecular site of action), there is often the misconception that a drug that is believed to act at a molecular target (noun) is by default also able to target (verb) or “home in” on that target (noun). It would therefore be more appropriate to define a true magic bullet as a drug molecule that is specific in its activity for a molecular target but that is also able to selectively accumulate at this molecular target and exert a selective therapeutic action by virtue of both its specific activity and its selective accumulation. This distinction is important when we consider the daunting challenge of developing magic bullets for diseases like cancer and neurodegenerative diseases like Alzheimer’s, as well as hormone imbalance diseases like diabetes that are becoming more widespread. Unless unique molecular targets found exclusively (or at sufficiently higher levels) in the diseased state and not in normal state are discovered, magic bullets by the traditional definition or the compromise of dose-dependent activity at the site of action may not be feasible. Strategies to effectively control the disposition of drug molecules thus represent an important tool for successful therapy. At a very basic level, selective accumulation is influenced by bioavailability and subsequent biodistribution. In the context of drug molecules, biodistribution is primarily related to physicochemical properties. Many potent drug candidates exhibit low bioavailability due to their limited water solubility. On the other hand, water-soluble compounds display a very limited ability to cross biological membranes, which essentially can exclude them from the cell interior. To overcome the limitations that a compound’s physicochemical nature can impose on its potential pharmaceutical application, the process of largescale screening of chemical libraries has been extended beyond just identifying desired bioactivity. Screening approaches routinely incorporate selection for desirable physicochemical properties that might confer high bioavailability as well. On the down side, this approach leads to many potent molecules being

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excluded from further development because they aren’t true magic bullets; that is, while they may have a potent pharmacological action at a desired molecular target they aren’t able to find their way exclusively to that target. There is most certainly a growing list of such molecules that are in essence potential drugs if only a delivery strategy can be devised to get them to their molecular target in the human body. As the search for the perfect magic bullet continues there is a significant effort to improve the action of currently available molecules by using targeted delivery approaches. It is not surprising that the field of drug targeting has grown significantly in the effort to develop better therapy. Based on the experience over the decades since Ehrlich first introduced the magic bullet concept, it seems more reasonable to separate the functions of pharmacological action and selective accumulation into properties desired in a drug molecule and in a delivery system, respectively, rather than the traditional expectation that the drug molecule alone possess both properties. Generally, drug disposition may be modulated via three broad approaches First, the drug molecule might be modified subtly to change its physicochemical properties without adversely affecting its inherent pharmacological action. This is essentially the intent of medicinal chemistry approaches and the concept of structure–activity relationship (SAR) studies that have now become standard practice and often aren’t even considered a means of achieving targeting. The second approach might be considered to be an extension of the first but is different in that it involves using chemistry to conjugate ligands that are often larger than simple organic functional groups to change the biodistribution of a molecule. Again this approach works as long as the conjugation does not adversely affect the desired pharmacological activity of the molecule. Conjugation using selectively cleavable linkers is an extension of this strategy. The third strategy involves the use of some sort of delivery system or a carrier system and does not involve chemical modifications to the pharmacologically active molecule. Pharmaceutical nanocarriers fall into this category and several such technologies are being developed that are fast becoming applicable to a variety of pharmacologically active molecules. Pharmaceutical nanocarriers offer what might be viewed as a nonchemical approach to modify the disposition of drug molecules. All chemistry can be performed on the components of the nanocarrier system that can then be loaded with the drug to afford targeted delivery [2–5]. Most pharmaceutical nanocarriers can be modified for some level of targeting to specific tissues if not specific cell types. Long circulating liposomes and nanoparticles are able to passively target areas of leaky vasculature by virtue of the EPR effect and can additionally be modified with antibodies or other targeting ligands to afford cell specific recognition [6–10]. However, despite such advances, the improvement in drug action is not always dramatic. This is likely because many drugs act at molecular targets inside the cells and these molecular targets are often in well-organized subcellular structures inside the mammalian cell.

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The interior of a cell is very different from an aqueous buffer solution, in which small drug molecules can freely diffuse and randomly interact with potential cosolutes. In addition to the presence of the cytoskeletal network and various dispersed organelles, the cytoplasm contains a large amount of dissolved macromolecules. The concentration of dissolved macromolecules in the nucleoplasm and cytoplasm of living cells has been determined to be between 50 and 400 g/L [11, 12]. Subsequently, transport or diffusion events in such a crowded solution cannot be expected to be the same as those in buffer solutions. Generally, intracellular diffusion has been characterized as hindered diffusion, reflecting among other factors the high level of molecular crowding [13, 14]. Additionally, the fluid-phase viscosity of the cytoplasm and binding to intracellular components are believed to influence the diffusion of solutes inside a cell [15, 16]. While efforts aimed at thoroughly understanding cellular material properties such as cytoplasmic viscosity are currently underway [17], it is generally accepted that the physicochemical properties of the drug also play a major role in determining the subcellular fate of the drug molecule. Consequently, the ability to predict the influence of various properties of the drug molecule on the likely site of accumulation within the cell could prove to be a powerful tool in drug design to either select molecules with a desired subcellular accumulation or identify molecules that would benefit from subcellular targeting strategies. There is apparent fractal symmetry between the case of drug delivery to a cell and drug delivery to a molecular target inside a subcellular compartment. The cell could be viewed as being a small, slightly simpler but nonetheless highly organized “body” with “organs” (organelles) and “cells” (defined structures and molecular arrangements) within these organs. It should therefore stand to reason that controlling drug disposition within the cell might also be necessary for optimal drug action [18–27]. Consequently, the next logical step in the development of targeted nanocarriers would be to extend our control over nanocarrier distribution to the subcellular level as well. At least two major schemes can be imagined to be useful in the design of nanocarriers with the potential for subcellular targeting: the first based on the inherent predisposition of the nanocarrier for a particular compartment and the second based on attaching subcellular targeting ligands to the surface of nanocarriers to redirect their accumulation to the desired compartment. Essential to the latter of these approaches is the use of a subcellular targeting ligand. Such ligands could be, as in the case of leader sequences, derived from normal cellular trafficking processes or, as in the case of triphenyl phosphonium, based on observations of a predisposition of an organic compound for subcellular compartments. The availability of a wide range of subcellular stains is proof enough that there are several molecules with an inherent ability to accumulate in a particular subcellular compartment. Based on the intracellular distribution of a large variety of fluorophores, a quantitative structure–activity relationship (QSAR) model for predicting cellular uptake and intracellular distribution of low molecular weight compounds

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has been proposed [28]. This QSAR approach was recently applied to identify potential common chemical features of molecules that are known to selectively accumulate at or inside mammalian mitochondria within living cells [29]. The QSAR approach has additionally proved useful for the modeling of cationic transfection lipids [30] and could therefore be applicable to predicting the subcellular disposition of a potential therapeutic molecule and even to design molecules with a desired subcellular affinity for the development of subcellular targeting approaches. Approaches to nanocarrier-mediated subcellular delivery are based on the principle that the subcellular destination of a drug is the same as that of the nanocarrier. Nanocarriers by virtue of their particulate nature are believed to be subject to endocytic cell entry mechanisms and subsequent endolysosomal processing. As such, nanocarriers could be considered to be ideally suited for delivering bioactive molecules to the endolysosomal system. Indeed, directing nanomedicine complexes to the endolysosomal system has increasingly gained attention, as pathological conditions associated with endosomes and lysosomes could potentially benefit from therapies targeting these pathways [31– 34]. Although endocytosis is a common mechanism that almost all cells possess for the internalization of macromolecules, a wide array of such vesicular internalization mechanisms exist [31]. For example, nanoscale drug carrier systems taken up by clathrin-dependent receptor-mediated endocytosis (RME) are most likely to undergo lysosomal degradation, while clathrin-independent RME may lead to endosomal accumulation [31]. Consequently, the type of targeting moiety displayed by the nanocarrier system will determine whether the carrier delivers its cargo to either endosomes or lysosomes. Well-characterized endocytic targeting moieties potentially useful for nanocarrier-mediated drug delivery are folic acid, low-density lipoprotein, cholera toxin B, mannose-6-phosphate, transferrin, riboflavin, the tripeptide RGD, ICAM-1 antibody, and nicotinic acid, as recently reviewed by Bareford and Swaan [31]. The cellular internalization mechanisms utilized by these ligands involve clathrin-dependent RME, caveolin-assisted endocytosis, lipid raft associated endocytosis, and cell adhesion molecule (CAM) directed cellular uptake [31]. In addition to several approaches to exploiting the inherent tendency of nanoparticles to accumulate in the endolysosomal compartment for possible therapeutic purposes, there is a growing body of work that suggests the feasibility of modifying nanocarriers to redirect delivery of their cargo to other subcellular compartments as well. Liposomes modified with mitochondriotropic ligands have been shown to improve the efficacy of an anticancer drug both in vitro and in vivo [35]. AuNPs have already been targeted to the nucleus using the adenoviral nuclear localization signal (NLS) and integrin binding domain [36]. Such an approach has been reported to be useful in the development of probes for cell tracking by surface-enhanced Raman scattering [37]. Modification with a leader sequence peptide has also been applied to creating delivery systems for mitochondria. A mitochondrial leader peptide

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(MLP), derived from the nucleocytosol expressed but mitochondria localized ornithine transcarbamylase, has been reported to render polyethylene imine (PEI) mitochondriotropic and represents a potential approach for mitochondrial DNA delivery [38]. It is interesting to note that the examples discussed above share a common assumption. Nanocarriers are assumed to have a predisposition for the endolysosomal pathway by virtue of their nanometer size and, without a subcellular targeting ligand, all nanocarriers would remain in the endolysosomal compartment. However, it is interesting to also consider the disposition of a nanocarrier made exclusively of a molecule with a predisposition for a subcellular compartment. A good example is the mitochondriotropic amphiphile dequalinium chloride. A serendipitous discovery while screening mitochondriotropic drugs potentially able to interfere with the mitochondrial DNA metabolism in Plasmodium falciparum [39] revealed this self-associating tendency of dequalinium chloride and its ability to form vesicles. At the time of their discovery, these unusual vesicles were termed DQAsomes (pronounced dequasomes), that is, dequalinium (DQA) based liposome-like vesicles [40]. Based on the fact that these carriers were composed exclusively of mitochondriotropic molecules and that they were able to bind and protect DNA, DQAsomes were explored as potential mitochondria-specific DNA delivery vehicles for direct mitochondrial gene therapy [41–44]. DQAsomes have also been explored as a mitochondria-targeted nanocarrier system for small drug molecules, in particular, for anticancer drugs known to trigger apoptosis via direct action on mitochondria [45, 46]. It would therefore appear that, for now, a basic proof of concept for an alternative strategy toward the design of subcellular targeting nanocarriers seems to have been established. It is also obvious that in order to design similar carriers for other subcellular compartments it would be necessary to first find self-assembling molecules with an affinity for the intended subcellular compartment. To this end, recent work on the subcellular distribution of micelle-forming agents offers some interesting insights [47–51]. Based on the examples discussed so far, it would seem that there is indeed hope that nanocarrier systems could be designed to achieve true molecular level targeting inside cells. However, to say that these systems will be available soon is perhaps premature given what little we know about the subcellular dynamics associated with nanoparticle trafficking. There are in our opinion several unanswered questions. For example, do all nanocarriers remain intact upon cell entry and subsequent disposition? Are there differences in the disposition of vesicles in comparison to particles? What is the true influence of size on the intracellular disposition of various nanocarriers? Most important, however, is the question of the mechanism by which the nanocarrier is able to achieve selective uptake and delivery into the subcellular compartment. All the strategies described so far report observations of altered or improved subcellular accumulation that appears to result in improved activity, but how exactly this happens is still unclear. Do the nanocarriers remain intact upon

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internalization and then get trafficked as intact structures? If so, how is the therapeutic cargo released to the correct subcellular compartment? Alternatively, it could be imagined that once taken up into the early endosomal vesicle, the nanocarrier components undergo a redistribution to become part of the endosomal vesicle. There is some evidence to suggest that in fact cells actively traffic nanocarriers in cell membrane-derived vesicles [52]. Assuming the targeting ligand was able to redistribute to the surface of the endosomal vesicle, it might be possible then that the vesicle would have an altered subcellular fate that could involve transport to and association with a target compartment other than the lysosome. While this may seem to be farfetched speculation, there has already been some work along similar lines toward the development of nanocarrier systems for delivery of molecules to the nucleus and even the mitochondria. A strategy that involved stepwise membrane fusion was devised based on the premise that, to efficiently deliver DNA to the nucleus, a delivery system must penetrate through the plasma membrane, nuclear envelope, prior to DNA release in the nucleus. Using a multilayered nanoparticle called a Tetralamellar Multifunctional Envelopetype Nano Device (T-MEND) and consisting of a DNA-polycation condensed core coated with two nuclear membrane-fusogenic inner envelopes and two endosome-fusogenic outer envelopes, which are shed in stepwise fashion, transgene expression in nondividing cells was reported to be dramatically increased [53]. A similar approach in designing a mitochondria-specific delivery sytem has been reported as well. Liposomal carriers called MITO-Porters, which carry octaarginine surface modifications to stimulate their entry into cells as intact vesicles (via macropinocytosis) were prepared with lipid compositions that were identified in various experiments to promote both fusion with the mitochondrial membrane and the release of liposomal cargo to the intramitochondrial compartment in living cells. Using GFP protein as a model cargo, it was shown that MITO-Porter liposomes are able to selectively deliver their cargo to mitochondria [54, 55]. It is also interesting to note that changes in nanoparticle architecture result in changes in subcellular disposition [56]. Fluorescein isothiocyanate labeled layered double hydroxide (LDH) nanoparticles were prepared from Mg2Al under conditions that yielded either hexagonal sheets (50–150 nm wide and 10–20 nm thick) or nanorods (30–60 nm wide and 100–200 nm long). A comparison of the subcellular distribution of these two types of preparations revealed that the nanorods trafficked to the nucleus but the hexagonal sheets remained in the cytoplasm [56]. Not surprisingly, an active microtubule mediated transport process is hypothesized to be responsible for the observed rapid nuclear accumulation of the nanorods [56]. As discussed so far, various nanocarrier platforms have already undergone preliminary investigation for their ability to control the subcellular disposition of drugs and a potential improvement in therapy through the use of nanocarriers for subcellular targeting of bioactive molecules. A common paradigm that currently seems to apply to most of these approaches is the use of

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subcellular targeting ligands to control subcellular distribution. Given the relative ease of modifying nanocarriers with various surface functionalities and the fact that such approaches are already in use to achieve targeting at the cellular and organ levels, the ligand-based approach does seem to be a logical extension of current technology. It helps that new tools are concurrently being investigated to understand some of the physicochemical aspects of how small molecules [28, 29] as well as proteins [57] are able to selectively accumulate in certain subcellular compartments to afford the rational design of a wide repertoire of subcellular targeting ligands. However, one must be cautious in the knowledge that the approaches described so far are based only on current understanding of subcellular trafficking. Current knowledge of subcellular trafficking processes is based largely on studies with solid nanoparticles and on quantum dots [58–64]. Whether these observations can be extended to vesicular carriers like liposomes and micelles remains in question and in our opinion is due in large part to current limitations in imaging technology. We are however hopeful that technological advances in real-time fluorescence confocal imaging of live cells [65–68], as well as the emergence of new imaging techniques like total internal reflection microscopy [69] and label-free approaches like Raman microscopy [70–72], will allow some of the questions raised to be more satisfactorily answered. This book is our attempt to bring the best of current knowledge together to provide a comprehensive resource for anyone interested in this emerging area of drug delivery. The subsequent chapters discuss in full detail the current state of the art in the various approaches to nanocarrier-mediated bioactives to cell organelles as well as emerging research methods for the identification of subcellular targeting ligands and the study of subcellular transport processes.

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CHAPTER 2

Delivery of Nanosensors to Measure the Intracellular Environment PAUL G. COUPLAND and JONATHAN W. AYLOTT School of Pharmacy, University of Nottingham, Nottingham, United Kingdom

2.1

INTRODUCTION

Nanosensors can be internalized in viable cells, providing an optical signal to report on the intracellular environment. This allows an understanding of intracellular function to be gained in real time. Nanosized sensors are especially desirable as they provide an output with minimal cellular disruption, which enhances the quality of data generated from each cell. Chemical perturbation is also minimized by the structural design of a nanosensor; it is these characteristics of nanosized sensing particles that give the potential for observation that is minimally invasive [1–5]. In recent years, new nanosized sensing particles have been demonstrated and reported widely in the literature. Borisov and Klimant [6] provide a good summary for the state of the art of optical nanosensor technology and its increasing use in scientific fields, including biology, biotechnology, and clinical medicine. In this chapter we will focus our discussion on nanosensors of a single polymeric matrix, for example, polyacrylamide or silica sol-gel, and the delivery methods used for intracellular translocation of these nanoparticles. The delivery principles discussed here are transferable to many types of nanoparticle systems currently being researched. The most common technique for intracellular research utilizes fluorescent dyes in combination with confocal microscopy [7]. The small size of the free sensing molecule is a key advantage, providing high spatial resolution and allowing information throughout the cell to be collected en masse. A problem faced when using free dye in direct contact with the intracellular environment of the cell is chemical perturbation due to any cytoxic action of the dye molecules [5]. Equally, the intracellular environment can impact on the efficacy Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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of the sensing molecules [8, 9], where random protein binding and intracellular sequestration are major complications [10] as well as simply retaining the dye within the cell during the lifetime of an experiment [11]. To overcome the analytical difficulties of using free sensing dyes, an alternative approach evolved—optochemical sensors. These are typically a pulled optical fiber with a modified polymeric tip, to entrap sensing dye molecules, inserted directly into the cell [12, 13]. Pulled fiber sensors provide a biocompatible platform, protecting the dye and the cell from one another, and the polymer matrix of the modified tip facilitates complex sensing schemes to be devised by closely packing elements that interact complementarily [14, 15]. However, the large size of the sensor relative to a single cell causes damaging physical perturbation to all but the largest cells. This is due to the physical pathway required to transmit the signal generated at the tip of the sensor to the detector. Additionally, only low spatial resolution is achievable as few optodes can be inserted into a cell before it is damaged irreversibly [10, 16]. Fiber-optic sensors and free-dye molecules have advantages over each other, yet both have the limitations described that prevents them from being ideally suited to single cell intracellular measurement. A new technique that retained the benefits of free dyes (small size, good spatial resolution) and optochemical sensors (biocompatible dye entrapment, complex sensing schemes) that prevented chemical and physical perturbation of the cell was required. The development of PEBBLE (probes encapsulated by biologically localized embedding) nanosensors by Kopelman [1] was the next step in miniaturization of pulled fiber optochemical sensors and removed the fiber-optic component used for signal transmission. Understanding that the tip of the fiber-optic sensor was all that was needed for sensing, and recognizing that fluorescence microscopy was ideally suited to record the fluorescence output, Kopelman undertook to fabricate the same polymer tip as a single unit. Typically sub 200 nm in diameter, these sensors occupy approximately 1 ppb of a typical mammalian cell and cause negligible physical perturbation [4]. Nanosensors resemble the tip of an optode, in that they consist of a polymer matrix surrounding chemical sensing elements; there is no longer a physical signal path—thus being more akin to free dyes—but the protective capacity of a biocompatible polymer matrix is retained. Chemical perturbation to the cell is minimized, as is the effect of the intracellular environment upon the efficiency of the reporter molecule; in essence, the polymer matrix protects the cell from the dye and the dye from the cell [1]. Fluorophores usually affected through random protein binding respond characteristically when entrapped within the polymer matrix of a nanosensor [17, 18]. The matrix also allows for enhanced longevity of the sensing components (by protecting the dye from protein binding) enabling fully reversible sensors to be produced [3]. The nanosensor matrix is a discrete domain in which components making up a more complex sensing regime can be located. This allows for highly selective sensors incorporating ratiometric sensing regimes that use more than one dye [4], or electron transfer regimes using optically silent ionophores, in conjunc-

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17

tion with fluorophore partners [15]; significantly, the use of a sensor matrix increases the scope of analysis. Nanosensors retain the spatial resolution advantage of free dyes rather than being limited to the immediate locality of a single optode tip, because many nanosensors can be loaded into a single cell. In brief, a nanosensor is a nanometer-size polymeric spherical matrix in which chemical sensing elements/fluorophores are entrapped. The polymer matrix is porous, allowing diffusion of analytes to interact with the entrapped fluorophores, giving a fluorescent response that is captured with an optical system [19, 20]. By minimizing the chemical effects of free dyes and the physical effects of bulky optodes, a more natural state of the intracellular environment can be investigated [21]. Nanosensors are most commonly fabricated from polyacrylamide [2], silicate sol-gels [5], or liquid polymer matrices [22], each being porous, photostable, optically transparent, and chemically inert. Functionalization of the nanosensor matrix is possible when required, to provide chemical functional groups on the nanosensor surface for subsequent modification and/or conjugation. The variety of matrices available for nanosensor fabrication and their chemical diversity allow for a flexible and modular approach to nanosensor design (Figure 2.1). O NH

2

O

O

O

N

N H

O N H

O

R functional matrix, chemical groups

lon+ polymer matrix, (silica sol-gel, polyacrylamide, liquid polymer)

CPP attachment

NH

O

O

O

N O NO H

O O

O N

N HO

S

NH 2

O

ionic matrix, comonomer additives +/–

SP* Lipid Multi-layer multifunction surface chemistry

O 2 NH

S

Fluorophore(s) (incl. FRET pairing), ionophores, enzymes hv

O

S

hv

O 2

S

O

N

Nanosensor function

Encapsulation in liposomes

PEG CPP *Signal Peptide

Figure 2.1. Nanosensor schematic. (See color insert.)

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The surface of the nanosensor is important due to its involvement in a number of biophysical interactions. Modification of the nanosensor surface with chemical moieties to provide a specific function allows for application to different cell types, delivery methods, and intracellular tasks. Modification of the nanosensor matrix can be achieved, for example, by the inclusion of an amine-modified acrylamide into a polyacrylamide matrix, providing primary amines throughout the nanosensor matrix [17]. Various molecules, from heterobifunctional linkers to phospholipids, can then be conjugated to the nanosensor, providing additional functionality and capability. As with all emerging technologies, continued development and improvements to the processes and protocols are ongoing. For nanosensors, one approach to improve their reporting of intracellular function and condition is by increasing the brightness of the signal emitted from each individual nanosensor. This is important in the realm of delivery and analytical performance, as brighter nanosensors will generate the same amount of signal from a cell with a lower nanosensor loading. This of course reduces the potential for physical disruption to the cell during and after delivery. Alternative strategies for attachment of fluorophores to the nanosensor surface, or entrapping them in the polymer matrix, to yield much brighter nanosensors are being developed. Historically, the most common approach is to physically entrap fluorophores into the polymer matrix. This method typically requires that fluorophores are attached to a large, inert molecule such as dextran, which will prevent subsequent leaching of the entrapped fluorophore from the polymer matrix. An alternative methodology exists for incorporating fluorophores in nanosensors, whereby the fluorophores are covalently attached directly to the backbone of the polymer matrix [23]. This eliminates the requirement of dextran altogether and allows for the potential incorporation of greater numbers of fluorophores by several orders of magnitude. However, indiscriminate attachment to the functional groups in the polymer matrix includes those on the nanosensor surface and as the nanosensor matrix provides a protective role, attachment of fluorophores on the nanosensor surface will contradict the idea of the protective capacity of a nanosensor. To prevent unwanted interaction between the fluorophore and its environment, additional steps are needed to remove or mask these surface-located fluorophores. Steps such as enzyme-mediated fluorophore removal would be required, where the fluorophores are attached via a cleavable linker allowing surface bound fluorophores to be easily removed [24]. Approaches to enhance the brightness of nanosensing particles have been investigated by other groups including the Wiesner group at Cornell University, who introduced superbright C-dots with brightness levels and enhanced photostability approaching those reported for semiconductor quantum dots [25]. C-dots are core-shell particles based on a dye-rich silica core surrounded by a pure silica shell. The shell acts as a protective layer similar to the polyacrylamide or sol-gel matrix of the nanosensors described here. Also, there are investigations into using quantum dots as intracellular probes and sensors [26–28]. The enhancement

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in brightness and functionality of nanosensors and other types of nanosensing particles is varied and promising but for each type of nano(sensor)particle the challenges of delivery remain.

2.2

INTRACELLULAR DELIVERY

The nanosensor is a tool that has been designed to observe and analyze intracellular domains with minimal cellular perturbation. However, one of the major challenges to the widespread uptake of nanosensor technology has been the lack of a generic delivery system to translocate the nanosensor across the cell membrane and to the intracellular domain of choice. There are several techniques by which nanoparticles can be delivered into cells in culture; historically, many have been developed for the delivery of plasmids and oligonucleotides in molecular biology, or for the uptake of molecules packaged within organic polymer structures developed for drug delivery. The various delivery techniques investigated can be designated into two main categories: mechanical and membranal. In the mechanical category each delivery technique uses a mechanical and physical force to insert the nanosensors into the cells, for example, using a needle to inject them directly or a pulse of electricity to disrupt the cellular membrane. Membranal techniques are based on an interaction at the biological level between the cellular membrane and a chemical group on the nanosensor surface that aids the nanosensor in its translocation to the intracellular environment. 2.2.1

Mechanical Methods

2.2.1.1 Gene Gun Bombardment One of the most commonly used nanosensor delivery methods to date is gene gun delivery. Originally developed in molecular biology research for the delivery of DNA into living cells, it was easily adapted for nanosensor delivery [2]. The gene gun method can be described as a shotgun method in which nanosensors are literally fired into cells. Nanosensors are layered onto a disk or the inner surface of a small length of tube (to create a bullet) depending on the model of gene gun, by applying a thin layer of suspended nanosensors in solution and allowing it to dry. Pressurized helium is then used to propel the nanosensors off the disk or from the prepared bullet into the cell culture. It is reported that by controlling the original concentration of nanosensors in the suspension when layered onto the carrier it is possible to deliver a single sensor or thousands of sensors into the cell culture sitting a short distance away. It is also argued [2, 5] that certain levels of spatial delivery are possible by altering the helium pressure and the distance from carrier disk to cell culture. As the nanosensors are intrinsically fired in a random pattern, this only serves to selectively choose the distance the nanosensors will travel through the cell—with a certain amount of control

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and practice (and luck!), this will allow one to deliver sensors to the nucleus or have them remain in the cytosol. Great care is needed to achieve the fine window of conditions that accomplishes cellular penetration without dislodging the cells from the culture plate. 2.2.1.2 Picoinjection Picoliter injection is a technique that has been applied to nanosensor internalization; physical damage and cellular distress are the main problems encountered during delivery. Needles are formed using a standard glass pipette puller and are used to inject a nanosensor solution directly into the cell. Generally, cell viability is good although normal cellular activity is likely affected and there is always the chance the cell becomes irreparably damaged. The deliverable range of nanosensor concentration is broad as this method dopes the cell with increasing numbers of nanosensorsolution aliquots until the desired level of concentration is reached. This method requires a high level of operator skill and is a batch process, meaning only one cell at a time can be loaded [9]. These limitations have led to this technique being used mostly in specialized applications where cell-by-cell analysis of a small number of cells is required, for example, embryology [21]. 2.2.1.3 Electroporation and Sonication These two methods have been used successfully for the delivery of various molecules into cells, such as plasmid DNA, for many years [29, 30]. One study suggests specifically that electroporation primarily transports molecules across the plasma membrane, because its mechanism is specific to lipid bilayer disruption, whereas sonication transports molecules across both the plasma membrane and cell walls, for example, in algae and plants, because it nonspecifically disrupts cell-surface barriers [31]. In electroporation an electrical pulse temporarily permeabilizes the phospholipid bilayer of the plasma membrane. As this technique bypasses the endocytic pathway, nanosensors can be delivered directly to the cytoplasm with no need for subsequent endosomal escape, which is a clear benefit when, for example, considering CPP mediated delivery, described later in this chapter. However, as the cells are both permeabilized and subjected to very strong electrical pulses, there is usually a high rate of cellular mortality, which is often not reported [28]. 2.2.1.4 Patch Clamp and Scanning Ion Conductance Microscopy (SICM) Both these methods require a pulled micro/nanopipette to be held in contact, or at least in very close proximity, to the cell membrane. The notion of using these techniques as methods for nanosensor delivery is simple; with slight modification to each technique, a nanosensor-loaded solution is contained within the pipette as it is brought to the cell membrane. During patch clamping transient cell membrane disruption caused by creating an electrical current allows a temporary route for nanosensors to internalize. Similarly, in SICM the scanning tip can be used to provide a disruptive electrical field, which again disrupts the phospholipid bilayer, allowing the surrounding

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nanosensors intracellular access. A major drawback to these techniques is the batch one-cell-at-a time nature of the technique. 2.2.1.5 Scrape Loading This procedure is used to transiently disrupt cell membranes and allow direct cell loading. This is achieved by drawing a tool (e.g., a glass rod with rubber end known as a rubber policeman) across an adherent population of cells, which scrapes them off the dish; the disrupted membranes temporarily allow various molecules [32], macromolecules [33], and nanoparticles [34] to cross this usually highly selective barrier. The method has not found great favor, however, due to the impracticality of severely reduced cell viability, approximately 50%, and because the method results in loaded cells that are in suspension. They must be replated and allowed to recover and spread, which is a serious disadvantage, as during this stage the labile probes/particles may be subjected to cytoplasmic redistribution and degradation. 2.2.2

Membranal Techniques

The dynamic nature of the endomembranal system is a consistent theme in different modes of intracellular delivery: specifically, sequestration through phagocytosis and pinocytosis in macrophages, liposomal delivery, and cell penetrating peptide mediated delivery. 2.2.2.1 Phagocytosis (Macrophages) and Pinocytosis Phagocytosis or “cell-eating” is the uptake of solid material by a cell. Only a few kinds of cells display this behavior, including protists (e.g., amebas and ciliates) and the phagocytic white blood cells of animals. The protists use it for feeding while the white blood cells use it as means of capturing bacteria and cleaning cell debris from infection sites. In phagocytosis the cellular membrane actively surrounds the solid object to be taken up, eventually pinching off internally to form a phagocytic vesicle, or phagosome. In general, this fuses with a lysosome whereupon the contents are digested or recycled. Macrophages have been shown to take up nanosensors from a surrounding solution, providing an easy delivery method for these types of cells [10]. The number of nanosensors internalized is a factor of the original concentration of nanosensors in the solution and how long the macrophages remain in solution. Cell viability is excellent. The disadvantages are that this technique is highly selective for macrophages, the sensors end up in certain cellular regions, as directed by the cell, and these specialized immune system cells can be difficult to culture. Pinocytosis or “cell-drinking” is the uptake of fluids from the outside of the cell. Here the plasma membrane tends to fold inwards, forming small vesicles containing the material. As in phagocytosis, the vesicle generally fuses with a lysosome and digestion of the contents takes place. Unlike receptor-mediated endocytosis, however, pinocytosis is nonspecific with regard to the materials taken up. Cell biologists first demonstrated pinocytosis by exposing amebas

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to concentrated protein solutions and observing protein uptake. Effective uptake of nanoparticles via pinocytosis has been demonstrated in mouse macrophages [35] and for nanosensors with mouse embryos [21]. 2.2.2.2 Liposomal Delivery Commercially available liposomes, such as Lipofectamine 2000, Fugene, and Escort, can be used to produce nanosensor– liposome complexes. The technique relies on the natural fusion of liposomes with certain cells, which occurs due to the structural similarities between liposome or lipid micelle and the phosholipid bilayer of the cell membrane. The nanosensor–liposome complex fuses with the cell membrane and nanosensors, internalized or bound to the wall of the liposomes, can become internalized within the cell. With cell-specific tailoring of cell to nanosensor to liposome concentration a wide range of nanosensor concentrations can be delivered in a similar fashion to the macrophage system, specifically the concentration of the original nanosensor solution and of nanosensor–liposome complexes incubated with the cells is important for achieving defined final concentrations [9]. Delivery to the cytosol is possible, which is a great benefit of this technique; this can be demonstrated with pH measurements of the cytoplasmic environment of approximately pH 7 (cytoplasmic pH is similar to cell culture media pH [36, 37]), indicating that sensors are not sequestered into acidified vesicles (endosomes/lysosomes). 2.2.2.3 Cell-Penetrating Peptides Cell-penetrating peptides (CPPs) are individual short peptides able to translocate across cell membranes. They are being studied widely because of their ability to aid translocation and drive the uptake of varied cargos into mammalian cells. There are several reviews on the potential and advantages of using CPPs as delivery vectors, in particular, therapeutic agents for human treatment [38–41]. One example illustrates how the functional efficacy of therapeutic agents would be greatly enhanced through the controlled import and intracellular targeting to specific organelles [41]. The value of using CPPs as delivery agents became more appealing as it emerged that CPPs could offer targeted delivery and that the short peptide structure of a CPP could be tailored for improved delivery strategies. CPP endomembranal delivery may prove to be an ideal method that is near silent— for both chemical and physical disruption. The effective cell translocation of the CPP nature was observed prior to the isolation of an individual peptide; this is because CPP sequences in nature form part of a larger protein. It was the observation that some proteins were able to translocate across cellular membranes by a process known as protein transduction that initiated the search for the element of each protein that was responsible. Several proteins are well known for this behavior, for example, human immunodeficiency virus (HIV-1) Tat protein [42], herpes simplex virus (HSV-1) VP22 protein [43], and Drosophila Antennapedia homeoprotein [44]. A portion of each protein associated with intracellular translocation was found through cleavage or site-directed mutagenesis, identifying the amino

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acids required for membrane translocation. This part of the protein was referred to as the “protein-transduction domain” (PTD) of each protein. The approach revealed PTDs that translocated the entire protein but also retained their activity in isolation, when either removed from the native protein or synthesized in the laboratory. In isolation, therefore, the PTD of a protein becomes a CPP [45]. Derossi et al. [46] working on the Drosophila Antennapedia homeoprotein revealed the PTD through site-directed mutagenesis, discovering the driving force behind the internalization of this protein; the third helix was essential for membrane translocation. Importantly, however, it was observed that the third helix was also sufficient for membrane translocation alone, and from this work it was possible to develop a 16 amino acid long CPP, referred to as penetratin (pAntp) [44, 47]. In a similar approach, Green and Loewenstein had identified an 86 amino acid section of the HIV-1 Tat protein that was internalized readily by viable cells [42]. Currently, CPPs are defined as peptides with a maximum of 30 amino acids, which are able to enter cells in a seemingly energy-independent manner, thus being able to translocate across membranes in a nonendocytotic fashion [48], although the exact biochemical and biophysical mechanisms involved in membrane translocation are yet to be established and debate is ongoing in this area of CPP research. It was thought that endocytosis did not play a part due to the evidence that peptide internalization occurred at 4 °C—because all active transport mechanisms involving endocytosis do not function at this low temperature. Also, CPPs being taken up by different cells and tissue types suggests a common internalization method, which suggests binding to conserved cell membrane determinants [49]. However, data from several groups argues for energydependent processes [50]. One regular feature of all CPPs and PTDs is the higher than average constitute of cationic amino acids [51]. This suggests that in some way at least the initiation of cellular uptake comes from ionic interactions between the cationic CPP and the anionic lipid membrane, although it is known that charge alone is not a sufficient driving force to induce translocation [40]. Recent literature describes a principal role for arginine in the CPP [52] and studies into the various permutations of Tat and poly-R-PTD (arginine-rich protein transduction domains) suggested an essential role of the guanidinium head-groups of the arginine residues for cellular uptake [52]. A good review of the delivery of quantam dots to cells, which describes the use of CPP in some detail, is provided by Delehanty et al. [28]. In our research we conjugated Tat peptide to the surface of aminefunctionalized polyacrylamide nanosensors for delivery to a variety of different cell lines, including CHO-K1, GH4 pituitary lactotrope, A172 human glioblastoma, and human embryonic stem cells [53] (Figure 2.2). It was demonstrated that the hydrodynamic radius of the cargo played a significant role in affecting the route of internalization. The function of the nanosensors was to determine pH and we showed that the nanosensors resided in vesicles with a measured pH of 5, postdelivery. These were presumed to be lysosomes, as they could be co-stained with LysoTracker Red® as shown in Figure 2.3. In

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Figure 2.2. Confocal fluorescence microscopy images of Tat-conjugated nanosensors delivered to (from left): CHO cells incubated with Ca2+-sensitive nanosensors (green) and DRAQ5 nuclear stain (blue); GH4 pituitary lactotrope cells loaded with Tat conjugated Ca2+-sensitive nanosensors; A172 human glioblastoma cell loaded with fluorescein nanosensors; human mesenchymal stem cells (hMSCs) loaded with rhodamine B nanosensors (red) and co-stained with CD105:FITC membrane stain (green) and DRAQ5 nuclear stain (blue). (See color insert.)

Figure 2.3. Confocal fluorescence microscopy images of Chinese hamster ovary K1 (CHO-K1) cells containing: (left) Tat-FITC conjugate (green) which do not show colocalized fluorescence with endosomes/lysosomes stained with LysoTracker Red® when loaded into CHO-K1 cells; (middle) Tat-conjugated nanosensors and LysoTracker Red® showing colocalized fluorescence (orange color) indicating that the nanosensors are residing in acidified endosomes: (right) Tat-FITC conjugate (green) and Tat-TRITC-nanosensors (red) loaded into a single cell. The overlay of these two channels shows no colocalization of fluorescence of the two different cargo sizes mediated by Tat peptide delivery. (See color insert.)

contrast to this, fluorescently labeled Tat, which had a much smaller hydrodynamic radius, localized to different vesicles in the cytoplasm and did not colocalize with the LysoTracker Red® dye. It was apparent that the difference in size of cargo that the Tat peptide was conjugated to seemed to directly affect the mechanism of translocation into the cell. The precise mechanisms involved following the first ionic interaction will continue to be debated; they may consist of currently unknown mechanisms or, simply, endocytosis. It is

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feasible, considering the physicochemical diversity of CPPs, that diverse mechanisms of uptake account for cellular uptake of different peptides. Even so, CPPs continue to be a useful research tool, having been demonstrated as delivery vectors for a variety of molecules and nanosized particles [54–57].

2.3 INTRACELLULAR TRAFFICKING, NANOSENSOR FATE, AND STRATEGIES FOR TARGETED DELIVERY After delivery, indicating the crossing of the cell membrane, the final location of nanosensors is often directed by cellular pathways that deal with the movement and trafficking of internalized particles and vesicles. As shown in Figure 2.3, CPP delivered nanosensors were found to locate to lysosomes within a few hours after incubation. This therefore means we have to carefully consider mechanisms by which the nanosensors can be directed to specific locations within the cell after internalization. Frequently, the required location of the delivered nanosensors is to reside freely in the cytoplasm without being sequestered into endosomal vesicles, for example, when wishing to monitor ion flux throughout the cell. Alternatively, the ultimate target may be a specific organelle. Either way, controlling the fate of the nanosensor is a detail that needs attention when developing a delivery system. 2.3.1

Cytochalasin-D

Cytochalasin-D is a cell-permeable fungal toxin from Zygosporium mansonii and an inhibitor of actin microfilament function. It binds to the barbed end of actin filaments, inhibiting the association and dissociation of subunits and causing disruption to the actin filaments and inhibition of actin polymerization. This action of cytochalasin-D is known to disrupt the fusion of the endosomes with lysosomes in late stage endocytosis [58], which is interesting with regard to nanoparticle delivery as internalization into lysosomes after translocating the membrane is limiting. Preincubation with cytochalasin-D has been shown to prevent nanoparticles being ultimately digested in lysosomes by blocking the fusion of endosomes with lysosomes [59]. For some delivery methods (e.g., CPP), this approach will only halt the endocytic pathway at the endosome stage; but this at least provides a point at which efforts can be focused on working to release the nanosensors from the endosome, before they are recycled out of the cell. 2.3.2

Targeted Delivery

For targeted delivery to specific intracellular organelles, like mitochondria, targeting molecules can be located on the nanosensor surface. These targeting molecules, often signaling peptides, are used ubiquitously in the cell to allow cell machinery to correctly transport proteins within the cellular environment

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[60]. Many examples have been found in cytology for protein translocation, including the sorting of proteins to peroxisomes, the endoplasmic reticulum, mitochondria, and chloroplasts. If these signal peptides were conjugated to the nanosensor surface, then there is potential for directing the nanosensor to an organelle of choice. Other surface-bound chemistries that may be useful in nanosensor surface chemistry are those that would enable endosomal escape. A route of investigation may be to accept the inevitability of endocytosis and concentrate on devising mechanisms for breaking the delivered payload (nanosensors) out of the endosome once inside the cell. Utilizing the fusion of nanosensor-bound lipids with the endosomal membrane, which can then be cleaved by cellular enzymes, may provide a means for endosomal escape and cytoplasmic localization. 2.3.3

Multilevel, Multifunctional Surface Chemistry

The nanosensor surface is a valuable locale that should be used to provide biophysical properties for enhanced delivery methodology. A theoretical nanosensor surface could carry out multiple functions in a specific sequence that chaperones the nanosensor through the various membranes and on to a specific intracellular microdomain. Consider a CPP-mediated delivery method that triggers endocytosis. In the outer level of the molecular coat the CPP is presented to the cell membrane; directly beneath this is a layer of PEG that protects other functional molecules closer to the nanosensor surface. The CPP triggers translocation across the outer cell membrane via receptor-mediated endocytosis that ultimately leads to the nanosensor being located in a lysosome. Importantly, the CPP and PEG layer are attached to the nanosensor by a linker that is cleaved by the lysosomal environment—be that a lower pH or lysosome-specific enzymes. Removal of the CPP and PEG layers then reveals a lipid layer on the nanosensor that fuses with, and breaks through, the lysosomal lipid membrane, gaining the nanosensor access to the cytoplasmic environment. Here the nanosensor may be available to take measurement or perhaps, when required, a photocleavable linker is illuminated that removes the lipids and reveals signal peptides that direct the nanosensor to a cellular organelle. Alternatively, the linker could simply be susceptible to cytoplasmic enzymes such as esterases.

2.4

CELL BEHAVIOR DIAGNOSTICS

What is vital in all delivery techniques is the minimal level of cellular disruption. It is worthy of reiteration here that developing a delivery method that is successful, in terms of the numbers of internalized sensors, is of little value if cellular perturbation (physical or chemical) is such that normal cellular processes are disrupted. Methods for assessing cellular normality/viability must

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therefore be carried out to appraise each delivery technique. This feature of nanosensor delivery can be referred to as “postdelivery cell behavior diagnostics,” which is the investigation that confirms and ensures a delivery method is as least disruptive as possible. A cell is considered viable if it has the ability to grow and develop [61, 62]; we extend our definition to include any morphogenic or phenotypic change that has occurred because of nanosensor delivery. For in vitro cultures of cells, assays to test viability can be based on the physical characteristics of the cells such as cell membrane integrity, cytoplasmic streaming in plant cells, deformability as monitored with optical tweezers [63], or granulation measured with flow cytometry. Several dyes are used for the assessment of cell membrane integrity, including trypan blue, methylene blue, and neutral red. These dyes are able to cross the disrupted membranes of dead cells and stain the intracellular contents. This is commonly referred to as a live : dead assay. The dead cells can be accounted for microscopically or with spectrometric estimation. Additionally, assays monitoring the metabolic activity, for example, the hydrolysis of fluorogenic compounds, are commonly used as indicators of cellular health. These assays often rely on the function of cellular enzymes (e.g., reductases or esterases) capable of altering the structure of an applied substrate to give an observable result. One such example is the administration of fluorescein diacetate, which is able to transfer across intact membranes. Inside the cell esterases cleave off the diacetate group yielding fluorescein; therefore the intracellular accumulation of this fluorescent compound correlates directly to the health of the cell. Again, fluorescent cells can be monitored with fluorescent microscopy, spectroscopy, or flow cytometry. The kinetic response to a chemoattractant could be monitored as another demonstration of normal cell function. Other methods based on the assessment of structural, morphological, and phenotypic characteristics can be used as additional or more convenient features of natural cell behavior including adherence and cell proliferation. For example, dielectrophoresis could be used to investigate changes in cell permittivity, and patterned electrode pair structures, used to measure impedance, can provide inferential measurement of proliferation. A method we employed to assay the well-being of a cell after delivery was to assess the degree of cellular identity. By this we refer to the unique identity of a cell, in terms of it being able to carry out a particular function or the type of receptors on the cell surface. Other proteinomic or genomic assessments may be used to identify the expressed biomolecular landscape of the cell and compare this with control populations. Human embryonic mesenchymal stem cells (hMSCs) were loaded with nanosensors prior to assessment with flow cytometry. A benefit of flow cytometry is the ability to investigate far greater numbers of cells than microscopy-based techniques; here each plot of data represents 50,000 cells. Flow cytometry can provide information on cellular size and granulation, as well as cell surface markers when the cells are previously incubated with relevant CD markers. In Figure 2.4 the dual use of flow cytometry data and confocal microscopy images show hMSC, containing

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Figure 2.4. Use of cell surface markers to confirm cellular identity. (Top) Simultaneous detection of CD29 expression on hMSC containing Tat-functionalized FITCnanosensors. (A) Isotype control showing only FITC fluorescence coming from internalized nanosensors. (B) CD29 second antibody control showing only FITC fluorescence coming from internalized nanosensors. (C) hMSC containing Tat-functionalized FITC nanosensors and stained with CD29-PE. Both signals are detected, as evidenced by the shift along the y-axis of the cell population. (Bottom) hMSC cells loaded with rhodamine B nanosensors (red) and counterstained with mouse anti-human CD105:FITC antibody (green). (See color insert.)

internalized Tat nanosensors, that retain the expression of CD29 and CD105, respectively. The hMSCs were positive for cell surface markers CD29 and CD105, and negative for CD34 and CD45 (control experiments; data not shown), demonstrating a population of undifferentiated hMSCs of nonhemopoietic lineage as defined by Pittenger et al. [64]. The combination of flow cytometry and fluorescence microscopy can be a powerful tool for cell behavior diagnostics.

2.5

CHALLENGES AND FUTURE PERSPECTIVES

The ultimate goal of nanosensor research is to have the ability to measure the analyte of choice at a cellular location of choice in real time. Further advantages accrue if the measurement system can be multiplexed to measure multiple analytes simultaneously. There is currently a rich stream of investigation to develop new measurement technologies at the nanoscale level to provide

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devices capable of probing the intracellular environment, concisely summarized by Borisov and Klimant [6]. New nanostructures, including “smart” sensors, and theranostics can be envisaged with increasing complexity and functionality being built into the devices. However, without improved delivery methods and, importantly, better understanding of how nanoparticles enter the cell and cross cell membranes the potential of these devices for intracellular analysis will not be realized. Each of the existing delivery methods described in this chapter has disadvantages when looked at in isolation. Most notable are the distress and physical perturbation to the cell from the “mechanical” loading techniques; gene gun delivery, picoliter injection, and scrape loading. Patch clamp and SICM methods have not yet found wide usage probably due to the cell-by-cell nature of loading, similar to picoinjection. Electroporation and sonication are thought to cause considerable disruption of the cellular membranes, which presents difficulties if trying to make measurements of “normal” cells. Of the membranal techniques phagocytosis and pinocytosis are limited to specific cell types, but liposomal delivery is used routinely for delivery of oligonucleotides and has been shown to work for nanoparticles too. It remains, however, that specific conditions need to be developed for each cell line and general applicability is notably limited. The attachment of cell-penetrating peptides has been investigated and shows promise as a technique for enabling nanosensor internalization without disruption to the cell wall or cellular biochemical activity of the cell. With nanosensors particularly, and most other nanosized particles destined for intracellular translocation, the surface of the particle provides the potential to be utilized for delivery strategies capable of traversing complicated routes through the cellular environment. Some of the exciting possibilities, in terms of intracellular targeting and fabricating multifunctional nanodevices were outlined. A final thought concerns the quality of fluorescence imaging as demand leads to the requirement of nanoscale resolution. A limitation at present is that the fluorescent output from nanosensors has to be assumed to arise from an ensemble of nanosensors as the standard optical microscopy resolution limit of λ/2 does not allow individual nanosensors to be imaged. The rapidly expanding field of superresolution microscopy, including the techniques of STED, PALM, STORM, and RESOLFT (see Hell [65] for a comprehensive review) will present opportunities to overcome the resolution limit; this may prove to be as transformative to intracellular imaging and analysis as the development and application of confocal microscopy has been to cellular analysis in the past 20 years.

REFERENCES 1. Buck, S. M., et al. Optochemical nanosensor PEBBLEs: photonic explorers for bioanalysis with biologically localized embedding. Curr. Opin. Chem. Biol. 8(5): 540–546 (2004).

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2. Clark, H. A., et al. Optical nanosensors for chemical analysis inside single living cells. 1. Fabrication, characterization, and methods for intracellular delivery of PEBBLE sensors. Anal. Chem. 71(21): 4831–4836 (1999). 3. Clark, H. A., et al. Optical nanosensors for chemical analysis inside single living cells. 2. Sensors for pH and calcium and the intracellular application of PEBBLE sensors. Anal. Chem. 71(21): 4837–4843 (1999). 4. Park, E. J., et al. Ratiometric optical PEBBLE nanosensors for real-time magnesium ion concentrations inside viable cells. Anal. Chem. 75(15): 3784–3791 (2003). 5. Xu, H., et al. A real-time ratiometric method for the determination of molecular oxygen inside living cells using sol-gel-based spherical optical nanosensors with applications to rat C6 glioma. Anal. Chem. 73(17): 4124–4133 (2001). 6. Borisov, S. M. and Klimant, I. Optical nanosensors—smart tools in bioanalytics. Analyst 133(10): 1302–1307 (2008). 7. Srivastava, A. and Krishnamoorthy, G. Time-resolved fluorescence microscopy could correct for probe binding while estimating intracellular pH. Anal. Biochem. 249(2): 140–146 (1997). 8. Clark, H. A., et al. Subcellular optochemical nanobiosensors: probes encapsulated by biologically localised embedding (PEBBLEs). Sensors and Actuators B—Chemical 51(1–3): 12–16 (1998). 9. Clark, H. A., et al. Optochemical nanosensors and subcellular applications in living cells. Mikrochim. Acta. 131(1–2): 121–128 (1999). 10. Buck, S. M., et al. Nanoscale probes encapsulated by biologically localized embedding (PEBBLEs) for ion sensing and imaging in live cells. Talanta 63(1): 41–59 (2004). 11. Sumner, J. P. and Kopelman, R. Alexa Fluor 488 as an iron sensing molecule and its application in PEBBLE nanosensors. Analyst 130(4): 528–533 (2005). 12. Tan, W. H., Shi, Z. Y., and Kopelman, R. Development of submicron chemical fiber optic sensors. Anal. Chem. 64(23): 2985–2990 (1992). 13. Vo-Dinh, T. and Kasili, P. Fiber-optic nanosensors for single-cell monitoring. Anal. Bioanal. Chem. 382(4): 918–925 (2005). 14. Vo-Dinh, T., et al. Antibody-based nanoprobe for measurement of a fluorescent analyte in a single cell. Nature Biotechnol. 18(7): 764–767 (2000). 15. Brasuel, M., et al. Production, characteristics and applications of fluorescent PEBBLE nanosensors: Potassium, oxygen, calcium and pH imaging inside live cells. Sensors and Materials 14(6): 309–338 (2002). 16. Tan, W. H., et al. Ultrasmall for cellular. Anal. Chem. 71(17): 606A–612A (1999). 17. Webster, A., Compton, S. J., and Aylott, J. W. Optical calcium sensors: development of a generic method for their introduction to the cell using conjugated cell penetrating peptides. Analyst 130(2): 163–170 (2005). 18. Sumner, J. P., et al. A fluorescent PEBBLE nanosensor for intracellular free zinc. Analyst 127(1): 11–16 (2002). 19. Koo, Y. E. L., et al. Photonic explorers based on multifunctional nanoplatforms for biosensing and photodynamic therapy. Appl. Optics 46(10): 1924–1930 (2007). 20. Peterson, J. I., et al. Fiber optic pH probe for physiological use. Anal. Chem. 52(6): 864–869 (1980).

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39. Vives, E., Schmidt, J., and Pelegrin, A. Cell-penetrating and cell-targeting peptides in drug delivery. Biochim. Biophys. Acta Rev. Cancer 1786(2): 126–138 (2008). 40. Futaki, S., et al. Arginine-rich peptides—an abundant source of membranepermeable peptides having potential as carriers for intracellular protein delivery. J. Biol. Chem. 276(8): 5836–5840 (2001). 41. Gariepy, J. and Kawamura, K. Vectorial delivery of macromolecules into cells using peptide-based vehicles. Trends Biotechnol. 19(1): 21–28 (2001). 42. Green, M. and Loewenstein, P. M. Autonomous functional domains of chemically synthesized human immunodeficiency virus tat trans-activator protein. Cell 55(6): 1179–1188 (1988). 43. Elliott, G. and Ohare, P. Intercellular trafficking and protein delivery by a herpesvirus structural protein. Cell 88(2): 223–233 (1997). 44. Derossi, D., Chassaing, G., and Prochiantz, A. Trojan peptides: the penetratin system for intracellular delivery. Trends Cell Biol. 8(2): 84–87 (1998). 45. Richard, J. P., et al. Cell-penetrating peptides—a reevaluation of the mechanism of cellular uptake. J. Biol. Chem. 278(1): 585–590 (2003). 46. Derossi, D., et al. The 3rd helix of the Antennapedia homeodomain translocates through biological-membranes. J. Biol. Chem. 269(14): 10444–10450 (1994). 47. Derossi, D., et al. Cell internalization of the third helix of the Antennapedia homeodomain is receptor-independent. J. Biol. Chem. 271(30): 18188–18193 (1996). 48. Hansen, M., Kilk, K., and Langel, U. Predicting cell-penetrating peptides. Adv. Drug Deliv. Rev. 60(4–5): 572–579 (2008). 49. Silhol, M., et al. Different mechanisms for cellular internalization of the HIV-1 Tat-derived cell penetrating peptide and recombinant proteins fused to Tat. Eur. J. Biochem. 269(2): 494–501 (2002). 50. Vives, E. Cellular uptake of the Tat peptide: an endocytosis mechanism following ionic interactions. J. Mol. Recognition 16(5): 265–271 (2003). 51. Futaki, S., Goto, S., and Sugiura, Y. Membrane permeability commonly shared among arginine-rich peptides. J. Mol. Recognition 16(5): 260–264 (2003). 52. Lundberg, P. and Langel, U. A brief introduction to cell-penetrating peptides. J. Mol. Recognition 16(5): 227–233 (2003). 53. Coupland, P. G., et al. Internalisation of polymeric nanosensors in mesenchymal stem cells: analysis by flow cytometry and confocal microscopy. J. Control. Release 130(2): 115–120 (2008). 54. Mae, M., et al. Design of a tumor homing cell-penetrating peptide for drug delivery. Int. J. Pept. Res. Ther. 15(1): 11–15 (2009). 55. Chen, B., et al. Transmembrane delivery of the cell-penetrating peptide conjugated semiconductor quantum dots. Langmuir 24(20): 11866–11871 (2008). 56. Torchilin, V. P. Cell penetrating peptide-modified pharmaceutical nanocarriers for intracellular drug and gene delivery. Biopolymers 90(5): 604–610 (2008). 57. Palm-Apergi, C. and Hallrink, M. A new rapid cell-penetrating peptide based strategy to produce bacterial ghosts for plasmid delivery. J. Control. Release 132(1): 49–54 (2008). 58. Qualmann, B., Kessels, M. M., and Kelly, R. B. Molecular links between endocytosis and the actin cytoskeleton. J. Cell Biol. 150(5): F111–F116 (2000).

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CHAPTER 3

Cytoplasmic Diffusion of Dendrimers and Dendriplexes ALEXANDER T. FLORENCE School of Pharmacy, University of London, London, United Kingdom

PAKATIP RUENRAROENGSAK Lung Cell Biology, Respiratory Medicine, National Heart and Lung Institute, Imperial College, London, United Kingdom

3.1

INTRODUCTION

Diffusion of nanoparticulate carriers within the cytoplasm is one of the controlling factors in the delivery of active therapeutic agents not only to the cell nucleus but also to a large extent to adjacent cells. Delivery to the nucleus is the goal of gene therapy but the overall flux of particles in tissues is determined cell by cell. The cytoplasm may be an aqueous environment but it is a complex one, crowded molecularly and far from isotropic. Hence the diffusion process of particles that have gained access to the individual cell in a free state, or escape from organelles in which they have been snared, and then diffuse, is complex. The flux of particles is determined by several factors: (1) by the properties of the medium in which the particles move; (2) by the properties of the particle, such as its radius, surface properties, shape, and flexibility; (3) by the concentration gradients that drive diffusion; and (4) by a range of other influences such as physical obstructions, which increase the effective path length that particles must traverse, and binding of nanoparticles to components within the cytoplasm. Convection currents may enhance or retard diffusion. The problem in both modeling and exact mathematical description is that the cytoplasm is a dynamic medium.

Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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In this chapter we describe some of the work conducted in our laboratories on the diffusion of dendrimers and dendriplexes, and discuss it in the light of findings in the literature. The bulk of the work discussed here centers on a novel synthetic 6.5-nm dendrimer based on branched lysine residues, which we found serendipitously to be autofluorescent. This feature of the dendrimer allowed us to investigate nanoparticle movement within the cytoplasm using confocal microscopy and other techniques. We also studied the diffusion of the dendrimer in aqueous media (water and glycerol solutions) and in aqueous gels for comparison with the much more complex cytoplasmic transport. One complication, which may not be unique to our dendrimer, is the finding that, at least in vitro, it has a biphasic effect on actin polymerization. Dendrimers are three-dimensional hyperbranched macromolecules [1]. They have been employed mainly in experimental applications encompassing diagnostic tools and as drug carriers; some have intrinsic biological activity. Despite this, elucidation of the uptake mechanisms and the cellular pathways of dendrimer carrier systems and dendrimers per se has been slow. Before diffusion can occur in the cytoplasm, the particles have to be taken up; the mode of uptake may influence subsequent events, depending on whether the particle emerges in the cytoplasm as a free entity or encapsulated within vesicles. The uptake of nanoparticles and dendrimers has perforce been studied by attachment of fluorescent probes either by conjugation [2] or by physical interactions [3]. Scission or loss of the fluorescent moiety will result in difficulty with interpretation of resulting data [4]. The use of an intrinsic fluorescent particle can overcome these problems as there are no chemical changes required and surface properties are the original. In our work, for example, with dendrimer–DNA complexes (dendriplexes), their diameter can increase by up to 20–25 nm on fluorescently labeling the DNA; this is accompanied by a twofold increase in the zeta potential of the complex. Artifacts are also created by fixing cells; thus experiments in live cells are preferable for estimating cytoplasmic transport [5, 6]. The synthetic self-fluorescent sixth generation amino-terminated polyamide polylysine dendrimer (Figure 3.1A), namely [(Gly)(Lys63)(NH2)64] with a molecular weight of 8149 Da, has been reported previously by us [7]. The use of this compound as a nanoscopic probe is discussed in this chapter, using confocal laser scanning microscopy (CLSM). The results relate both to the native dendrimer and to a dendrimer–DNA complex and are discussed in terms of diffusion, obstruction, and binding effects. We can only speculate on its effects on actin polymerization, which we found in vitro. From a naïve physicochemical point of view, if such effects occur in vivo, depolymerization would ease the passage of the dendrimer and polymerization would complicate it. However, actin polymerization actually provides the propulsive force for intracellular transport of endosomes and some microorganisms, so this is a complex subject [8].

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DIFFUSION WITHIN CELLS

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Figure 3.1. The structure (A) of the sixth generation polylysine dendrimer used in this study and a time-dependent study of the interaction between the dendrimer (green fluorescence) and Caco-2 cells (B–I): the cells were incubated with complete media containing 0.1% (w/v) of dendrimers for 15 min (C), 30 min (D), 1 h (E, F), 2 h (G, H), and 4 h (I). The cells were observed under confocal microscopy. In the control (A), the cells were exposed to the complete medium without dendrimer. The dendrimer is found to attach to the cell membrane periphery (red arrows) as a result of electrostatic attraction (C, D). At 1 h the dendrimer distributes toward the cell nuclei in all regions of the cells (E, F). At longer incubation times (G–I), the dendrimer appears to be concentrated on the surface of the cells. This technique cannot prove internalization. (See color insert.)

3.2

DIFFUSION WITHIN CELLS

Diffusion of spherical nanoparticles in a liquid medium obeys the Stokes– Einstein equation relating diffusion coefficient (D) to the radius of the particle (r) and the viscosity (η) of the medium in which the particle moves as

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D = kT 6πηr One can demonstrate in binary aqueous mixtures (e.g., glycerol–water) that the diffusion coefficient reduces with increase in bulk viscosity, as these are simple fluids composed of small molecules. In macromolecular systems, the viscosity parameter in the Stokes–Einstein equation is not the bulk viscosity but in fact the intrinsic or “microscopic” viscosity. Movement of particles through some gels is rapid because the “pore” sizes are large and the medium in which the particles move is akin to bulk water. We have calculated the intrinsic viscosity of the interior of HPMC gels to be very close to that of water, using a variety of nanoparticle probes. In nonionic viscous vehicles such as glycerol solutions and HPMC gels, there is no or little influence of particle charge [9]. As other chapters in this book discuss, the cell cytoplasm is concentrated and crowded, occupied as it is by a variety of macromolecules between 5% and 40% of the total cell volume or 400 mg/L [10]. The cellular architecture provides a complex non-Newtonian fluid comprising an aqueous phase filling the spaces between an entangled mesh of filamentous cytoskeleton and other macromolecular structures, resembling a gel-like structure [11–14]. This phenomenon is well known as “macromolecular crowding”, which plays a decisive role on several levels of cellular organization, particularly in the diffusion and binding of matter inside cells [13]. Translocation of nanosystems occurs in this three-dimensional forest. The Stokes–Einstein equation may not be entirely applicable to diffusion phenomena within the constantly changing interior of cells. Diffusion of matter in cells relies on fluid-phase viscosity (F1), solute binding to macromolecules (F2), and collisions between solutes and macromolecules (F3) [15, 16]. Diffusion theory of solutes in the cell cytoplasm has been described by Kao et al. [16], who assumed that the cytoplasm was composed of an aqueous fluid-phase compartment bathing a matrix of mobile and static macromolecules/particles much larger than water molecules and small solutes. They identify three main factors involved in the reduction in diffusion coefficient of a small solute in cytoplasm (Dcyto) relative to that in water (D0): Dcyto = F1 ( η) × F2 ( Du , {Db,i , fb,i }) × F3 ({ni , Vi }) D0

(3.1)

The function F1(η) represents the deceleration of net solute translational diffusion because of an increase in true fluid-phase cytoplasmic viscosity, an increase due to solute-induced perturbation in solvent structure, which can be written F1 ( η) =

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η0 ηcyto

(3.2)

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DIFFUSION WITHIN CELLS

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where η0 represents the viscosity of water and ηcyto is the true fluid-phase microviscosity of the cytoplasm. The function F2(Du,{Db,i, fb,i}) indicates the total reduction of solute translational diffusion due to the transient binding of solute molecules to cytoplasmic structures as addressed in Equation 3.3 below [16]. F2 represents the ratio of the weighted diffusion coefficient of bound and unbound solute to the diffusion coefficient of the unbound solute. In Equation 3.3 Du and Db,i are, respectively, the diffusion coefficients of unbound and ith bound solute, and fb,i is the fraction of net solute bound to component i. F2 ( Du , {Db,i , fb,i }) = fu + ∑ ( Db,i Du ) fb,i

(3.3)

fu + ∑ fb,i = 1

(3.4)

i

where i

The function F3({ni,Vi}) illustrates the deceleration of the total reduction of solute translational diffusion owing to collisional interaction with cytoplasmic structures assuming there are ni structures of type i, each of volume Vi. The “volume exclusion” by mobile obstacles has been used as a model (a stretchedexponential [SE] model) to explain this function [17–19]. This SE model is applicable to the diffusion of Brownian particles (or solutes), which are relatively large compared to the solvent molecules. v F3,SE ( ni , Vi ) = exp ⎡⎣ −α ( ni , Vi ) ⎤⎦

(3.5)

Here niVi represents the volume fraction occupied by the occluding molecule and the prefactor α and ν represent scaling parameters obtained by fitting Equation 3.5 to data, but can be calculated independently from theory [16]. Viscosities of the cytoplasm of various cell types have been calculated. Values ranged from 2 to 20 cP [16, 20–22]. By using time-resolved fluorescence anisotropy, Verkman and colleagues found that the cytoplasmic solvent measured in different types of cell lines may be significantly different from that of bulk water [23]. This has also been demonstrated [24] using a different approach in two mammalian tissue culture cell lines. Luby-Phelps et al. [25] characterized the nature of the diffusion barrier by comparing the diffusion of 3.2–25.8 nm Ficoll probe molecules. The results showed that relative to water the diffusion rate in the cytoplasm, which is already low with the smallest (3.2-nm) probe, reduces with an increase of the probe size. To describe this relationship, diffusion of various sizes of FITCFicoll was measured in model solutions [19]. The relative diffusion coefficient of FITC-Ficoll in concentrated solutions of Ficoll is low, but it is independent of the size of the FITC-Ficoll probe. In a tangle of F-actin filaments, on the

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other hand, relative diffusion of the 3.2-nm probe is not impeded, but its diffusion decreases with the larger FITC-Ficoll probes. Taken together, it was noticed that the effects of Ficoll (10%) and F-actin filaments (5 mg/mL) resemble the pattern seen in the cytoplasm. The solution conditions calculated to most closely mimic the size dependency of FITC-Ficoll diffusion in the cells are 120 mg/mL of bovine serum albumin dissolved in a fluid matrix containing 37 mg/mL F-actin fibers. Based on these results, a model of cytoplasm as a densely entangled filament network interpenetrated by a fluid phase crowded with globular macromolecules was developed. This is in good agreement with cell structures proposed by Medalia et al. [14]. Uncoated GFP-M6tail-associate vesicles within cells [26], found during the process of clathrin-dependent endocytosis, are trapped in an actin-rich region of the cytoplasm of ARPE-19 cells. These vesicles exhibited Brownian-like motion. They could exit from the actin-rich region by a slow diffusion-based mechanism. The diffusion coefficient of the vesicles (n = 250) was found to be 1.42 × 10−12 cm2/s (SD = 1.24). Small molecules traverse cell membranes by passive transport, whereas macromolecules cross membranes by endocytosis, the process of membranebound vesicles originated from the invagination and pinching off of the membrane [27]. Phagocytosis occurs in mammalian cells, while pinocytosis may occur in all cell types. It has been reported that some dendrimers may create holes in cell membranes ranging between 15 and 40 nm, allowing passive diffusion of dendriplexes [28–32]. In addition, polylysine polymer was found to initiate holes on the cell membrane with a diameter ranging from 1 to 10 nm [29]. This suggests that, as we believe, the polylysine dendrimers used in our work are taken up intact and are not sequestered. The use of cell-penetrating peptides to increase the bioavailability of drug/gene/siRNA delivery has been seen to overcome both extracellular and intracellular barriers [33]. Peptide carried MPG and Pep was found to form a stable nanoparticle complex with the active macromolecules. The particles were found to enter the cells independently of endosomal pathways. Both lipoplexes and dendriplexes possessing positive charges first adhere to the cell membrane as a result of electrostatic interactions. Endocytosis has been invoked depending on types of carrier systems and cell lines [34, 35]. The nature of the carrier surface, in fact, regulates specific or nonspecific binding of the complexes onto the cell membrane and triggers different endocytic pathways such as clathrin- and caveolin-dependent endocytosis, and clathrinand caveolin-independent endocytosis. Clathrin- and caveolin-dependent endocytosis has been proposed as the major pathway of internalization of lipoplexes and dendriplexes [36–38]. Interestingly, a difference in uptake pathway was also found between surface-engineered dendrimers with amine and carboxyl functional groups. The carboxyl-modified dendrimers were taken up by caveolin-dependent endocytosis, whereas the amine-modified dendrimers were taken up by the clathrin- and caveolin-independent pathway [2].

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41

3.3 METHODS USED TO ASSESS THE UPTAKE AND CELLULAR DIFFUSION OF NANOPARTICLES The self-fluorescent sixth generation amino-terminated polyamide polylysine dendrimer (Figure 3.1A), with a molecular weight of 8149 Da, was used in this study and the details of the synthesis method can be found elsewhere [7]. For the dendriplexes, the same dendrimer was complexed with plasmid DNA (pDNA), pDsRed2-N1 (4.689 kb, Clonetech, USA), at 10 : 1 molar charge ratio (+/−). As the fluorescence of the dendrimer decreases after the complexation, the pDNA was fluorescently labeled using Label IT® Tracker™ Fluorescein kit (Mirus Bio Coporation, USA) before forming the complex. Dynamic uptake of the dendrimer and dendriplexes in all types of cells was followed every 5 min before and after adding the dendrimer and dendriplexes using confocal microscopy and data analyzed using custom procedures written in IgorPro software (Wavemetric Inc). The z-stack image at time t = 5 min before the dendrimer or the dendriplexes was combined and used as the control image. The fluorescent intensity detected in the green channel in this stack represents the background fluorescent intensity. The regions of interest (ROIs) were drawn and saved as the reference file. The fluorescent intensity values of the dendrimer/dendriplexes inside each z-stack image were averaged and standardized by subtraction of the background fluorescent intensity. The diffusion coefficients (D) and relative diffusion coefficients (D/D0) of the dendrimer/dendriplexes were calculated assuming that the cytoplasm is homogeneous. The focus was on the transport of the dendrimer and dendriplexes from the plasma membrane to the nucleus [39]. The lag time (tL) for the dendrimer/dendriplexes to develop a uniform concentration gradient within cytoplasm allows calculation of a diffusion coefficient where the thickness (h) of the diffusion layer is known, as D = h2/6tL. The thickness is the mean distance between the plasma membrane and nuclear membrane, termed the “cytoplasmic radius.” The cytoplasmic radius was measured using TEM and confocal microscopy. The value of tL was obtained by measuring the fluorescence intensity of the dendrimer over time from uniformly sized regions of interest placed equidistantly across the cell using custom procedures written in IgorPro software. The diffusion coefficient (D) of the dendrimer in the cell cytoplasm was then calculated, using the experimental tL value. The set fluorescent intensity of ROIs in each time point (F) was then plotted against time (minutes).

3.4

DIFFUSION AND UPTAKE OF DENDRIMER AND DENDRIPLEXES

This topic is discussed here as uptake may be a rate-limiting step in subsequent events in the cytoplasm. There is of course a concentration dependency of both flux and diffusion coefficient [40]; hence rates and extents of uptake are important.

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3.4.1

CYTOPLASMIC DIFFUSION OF DENDRIMERS AND DENDRIPLEXES

Adhesion of Dendrimer to Different Cell Lines

The adhesion of the dendrimer to Caco-2 cells is time dependent as illustrated in Figure 3.1B–I. At 15 min, the dendrimer (seen by the green fluorescence) concentrates at the periphery of cell membrane (Figure 3.1C). Within 30 min, the dendrimer is found to accumulate in the cytoplasm (Figure 3.1D). The relatively rapid uptake into the Caco-2 cells and specifically the nucleus can be attributed first to the electrostatic attraction between the amino surface groups of the dendrimer and the negative charge of membrane proteins, allowing attachment prior to endocytosis. Within an hour, the dendrimer covers all parts of the cell (Figure 3.1E–F). Distribution of the dendrimer toward the cell nucleus is observed in dividing (D) and nondividing cells (E). The dendrimer was found to cover the cells and apparently concentrate in the cytoplasm region (Figure 3.1G–H). The cells after exposure to the dendrimer for 4 h are seen in Figure 3.1I; the higher number of areas of dendrimer coverage are found on the cells. Different results were found in SKMES-1 cells. The dendrimer appeared to randomly adhere to the cell membrane. At longer incubation periods a more typical trend was found: the dendrimer was found to cover the cell surface. The reasons for this variation remain unclear but may result from differences in uptake mechanisms. Xia et al. [41] found such diferences in uptake mechanisms for 60-nm diameter amine-modified polystyrene nanoparticles when studying macrophage (RAW 264.7) and endothelial cells (BEAS2B). The uptake of the particles in the latter was found to be caveolin-dependent endocytosis whereas that in the former was independent of caveolin. We have demonstrated differences in the rate of binding of our dendrimer to three different cell types, and also the reduction of cellular adhesion when the medium bathing the cells is flowing (results not shown here). The extent of reduction is dependent on the force of adhesion (unpublished data).

3.4.2 Diffusion and Uptake of Dendrimer and Dendriplexes in Living Cells At low concentrations of the dendrimer, the colocalization of the dendrimer may not be very obvious, but increased levels of the fluorescent signal of the dendrimer are found inside the cell nucleus after background subtraction. Interestingly, the same pattern was noticed in the nucleus after the dendrimer was added into the system (Figure 3.2A–C). Hoechst33342 was used for locating the nuclear compartment. The dynamic uptake of the dendrimer into the nucleus was examined by overlaying images of the dendrimers (in green channel (Ch2)) over images of the nucleus (in blue channel (Ch1)) as can be seen in Figure 3.2. The fluorescent signal in the green channel, representing the dendrimers in each ROI, was normalized by background fluorescence subtraction (white boxes no. 14

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43

Fluorescent Intensity

60 50 40 † Boxes in both channels are regions of interest (ROIs)

30

ROI 5 ROI 6 ROI 7 ROI 8 ROI 9 ROI 10

20 10 0 0

20

40 60 Time (min)

80

100

Figure 3.2. An overlay of the compressed z-stack images of the dendrimer uptake process in living Caco-2 cells. The cells were incubated with medium containing 0.1% dendrimer (represented in green) and images collected every 5 min before (A) and after addition of dendrimer (B–F). Images presented are at time t = 5 (B), 15 (C), 30 (D), 40 (E), and 90 (F) min of incubation. Data were analyzed over the time and plotted in (G); the inserts show the positions of regions of interest (ROIs, white boxes) for data analysis. The mean fluorescence signal in each ROI was measured from the projected z-stack images at different time points with background subtraction. The dendrimer was found in the cytoplasmic compartment 5–35 min after exposure to the dendrimer (ROI 9 and 10). The concentration of the dendrimer declined after 30 min, whereas the relative fluorescent intensity of the dendrimer inside the nucleus was detected (ROI 5–8). The cell nuclei were stained with Hoechst33342 represented in blue. The scale bar is 20 μm. (See color insert.)

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in the insert pictures of Figure 3.2G). The relative fluorescent signal of the dendrimer was then plotted (F) against time (t, min) as in Figure 3.2G. The results (n = 5) demonstrated that the dendrimer reached the nucleus of Caco-2 cells within 35–45 min (tL) of incubation (Figure 3.2G), compared to 25–30 min in SKMES-1 cells (data not shown). This might be due to the differences of the intracellular architecture within each cell line, but not cell size as the cell radius is taken into account in the calculation of D. Average cell radii were 12.09 (± 6.02 (SD)) and 8.01 (± 3.21 (SD)) μm for the Caco-2 cells and SKMES cells, respectively. Dcyto values of the dendrimer in Caco-2 cells and SKMES-1 cells were found to be 9.82 (± 0.98) × 10−11 cm2/s and 5.99 (± 0.16) × 10−11 cm2/s, respectively. Cellular components such as secretory granules have diffusion coefficients of 1.9 × 10−11 cm2/s, lower than that of the dendrimers [42] as might be expected from their larger dimensions. The diffusion coefficients relative to those in water (Dcyto/D0) are, respectively, 1.24 (± 0.12) × 10−4 and 0.76 (± 0.05) × 10−4 (D0 = 79.54 × 10−8cm2/s). These values imply that macromolecular crowding and obstacles in the cytoplasm provide a formidable barrier to dendrimer transport. They also emphasize that differences between cell lines can be significant. In work on transfection with dendriplexes, we have found that there can be a 1000-fold difference in efficiencies when using the same DNA construct with a range of cell lines [43]. The cytoplasm hinders the mobility of the dendrimer by some 1000-fold compared to that in water. This may be explained in loose terms by exclusion and obstruction effects, the former due to the loss of free water by hydration of macromolecules and the latter due to the tortuous pathway that the dendrimer must travel in the cytoplasm, avoiding organelles and other structural features, such as actin fibers. The diffusion of globular proteins in muscle cells approximates to zero when the hydrodynamic radius of the proteins increased from 1 nm (Dcyto = 16.3 ± 3 μm2/s) to 7 nm (Dcyto = 4.0 (± 0.7) × 10−3 μm2/s) [44]. This slower-than-predicted diffusion of the proteins or nanoparticles may be explained by an elasticity of F-actin/α-actinin networks in the cell cytoplasm [45]. 3.4.3

Dendrimer–Actin Interactions

We have evidence of an interaction due to the electrostatic attraction between the dendrimer and actin cytoskeleton in vitro, and we have determined the diffusion coefficient of the dendrimer in the presence of actin gel using the fluorescent recovery after photobleaching (FRAP) technique. The ratio Dactin/D0 of the dendrimer was found to be 0.26, at a concentration of actin gel of 1 mg/mL, much lower than the concentration inside cells (∼10 mg/mL). Although the medium in which particles diffuse may approximate to that of pure water [16, 46], the auxiliary reduction of the translation diffusion of the dendrimers in cytoplasm can be attributed to microscopic barriers, enhanced tortuosity, and binding. Further studies on this are needed.

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A

B

C

D

45

Relative fluorescent intensity (F)

E

90 80 70 60 50 40 30 20 10 0 0

20

40

60

Time (min)

80

100

ROI 2 ROI 3 ROI 4 ROI 5 ROI 6 ROI 7 ROI 8 ROI 9 ROI 10 ROI 11 ROI 12 ROI 13

White boxes are regions of interest (ROIs)

Figure 3.3. An overlay of the compressed z-stack images of the dendriplex uptake process in living Caco-2 cells. The cells were incubated with complete medium containing dendriplexes (represented in green) and images were collected every 5 min before (A) and after adding the dendrimer (B–D). Images presented here are at time t = 5 (B), 30 (C), and 60 (D) min of incubation period. Data were analyzed and plotted in (E) and the insert pictures show the positions of regions of interest (ROIs, white boxes) for data analysis in both channels. The mean fluorescent signal in each ROI was measured from projected z-stack images taken at different time points, with background subtraction. The dendriplexes were found to be taken up into the cytoplasmic compartment (ROI 2–3, 7, 9, 12–13) but there is no uptake of the dendriplexes into the nuclear compartment within 2 h of incubation (ROI 4, 8, 10). (See color insert.)

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3.4.4

Dendriplex Diffusion

The diffusion coefficients of the dendriplexes were expected to be lower than that of the parent dendrimer as they consist of many dendrimer molecules complexed with the DNA. In the dynamic uptake study (Figure 3.3) they were found after 2 h only in the cytoplasmic compartment throughout the stack images (A–D). Although dendriplexes were first found attached to the cell membrane after 5 min incubation, they remained in the cytosol until the end of the experiment (A–D). This was confirmed by the quantification of the fluorescent signal of the dendriplexes as plotted in Figure 3.3E (ROI 2–3, 7, 9, 12–13). There is no fluorescence in the nuclear compartment after 2 h. Changes in cell morphology were found in the cells after 2 h under the conditions of the experiment. Further work was carried out using living cells incubated with the dendrimer at longer time points; cells were removed to view at the end of each selected time point. The dendriplexes were found to be taken up into the nuclear compartment after 2.5 h of incubation and uptake was seen to be widespread in nearly every cell after 3 h. As this lag time is long, the calculated diffusion coefficient of the dendriplexes is lower than that of the parent dendrimer, as shown in Table 3.1. The lag time (tL) here was found to be between 2.5 and 3 h (n = 5), the diffusion coefficient of the dendriplexes in cytoplasm (Dcyto) was calculated to be 2.36 (± 0.34 (SD)) × 10−11 cm2/s (2.36 × 10−3 μm2/s) and the diffusion coefficients (Dcyto) relative to that in water (4.53 × 10−8 cm2/s (D0)) was 5.21 (± 0.75) × 10−4. In Caco-2 cells the cytoplasmic diffusion coefficient, Dcyto, of the dendriplexes relative to the dendrimer was found to be 0.24. The cytoplasm hinders the mobility of the dendriplexes up to 2000 times that in water. This might be explained by the interaction between the cationic dendriplexes and the cell organelles, which impede the mobility of the dendriplexes apart from the effect of particle size, sieving effects, and obstruction of the crowded condition of the cells. The larger unit should be more affected by the obstruction effect. Whether or not binding of the dendriplex, say, to actin filaments occurs is not known; it may be that dissociation of the complex occurs and the diffusion coefficients are mixed values. TABLE 3.1 Diffusion Coefficients and Relative Diffusion Coefficients of the Parent Dendrimers and the Dendriplexes in Cell Cytoplasm Compared to Other Media Diffusion Coefficients in Various Media (cm2/s)

Relative Diffusion Coefficients (cm2/s)

SKMES-1 Cells

Dactin D0

Dcyto D0

Ddendriplexes Ddendrimer

Dendrimer (6.5 nm)

79.54 × 10−8 23.75 × 10−8 9.82 × 10−11 5.99 × 10−11

0.26

—

Dendriplexes (107.3 nm)

4.53 × 10−8

1.24 × 10−4 (Caco-2) 0.76 × 10−4 (SKMES-1) 5.21 × 10−4

Nanoparticles

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Water

Actin Gel

—

Caco-2 Cells

2.36 × 10−11

—

—

0.24

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47

The differences are caused by two main factors: the physicochemical properties (size, charge, and mass) of the species and the uptake processes. The hydrodynamic diameter and zeta potential of the dendriplexes were, respectively, 107.3 nm and 39.80 mV compared to values for the parent dendrimer of 6.5 nm and 47.27 mV. The positive surface charge of both compounds facilitates electrostatic attraction with the cell membrane as a prelude to being taken up by endocytosis. Concomitantly, the dendrimer may be taken up by transient nanopores, which can be created after adherence to the cell membrane, allowing also passive diffusion and resulting in the shorter uptake process [28–32]. The larger size of the dendriplexes would attenuate their diffusion inside the cytoplasm. In comparison to the native dendrimer the diffusion coefficient of the dendriplex is reduced by a factor of 4. One might have expected a larger decrease simply because the radius has increased by ∼15-fold from 6.5 to 107 nm (D0 of dendrimer and dendriplexes are 79.54 × 10−8 and 4.53 × 10−8 cm2/s, respectively) in accord with the Stokes–Einstein equation. The diffusion coefficient in water of the dendriplexes is 20 times lower than that of the dendrimer. Although particle size precisely controls the diffusion of the particle in bulk or aqueous solution, it was found to also play a more complex role in the diffusion of the two particle types discussed here. Dcyto values of dendriplexes and dendrimer were reduced from 9.82 × 10−11 to 2.36 (± 0.34 (SD)) × 10−11 cm2/s (fourfold reduction). This may be related somehow to uptake pathways. In the cells, diffusion coefficients generally can be explained by the obstruction effect and sieving effects, which are a function of particle size (Figure 3.4A) [13, 25, 47–49]. Janson and Luby-Phelps proposed the pore slit model having an average cutoff radius between 15 and 50 nm. Following this model the diameter of the dendriplexes is beyond the maximum cutoff size and thus cannot pass through the cellular meshwork, which presents as the slow mobility of the dendriplexes at the rim of the plasma membrane, and their ability to traverse the cytoplasm. 3.5 MECHANISMS OF PARTICLE UPTAKE INTO THE CELL NUCLEUS The cell nucleus is the ultimate target to achieve efficient gene transfection. The exact process by which lipoplexes deliver their plasmid DNA to nuclei after escaping from endosomes has remained elusive. The nuclear pore complex (NPC) responsible for nucleocytoplasmic transport [51, 52] plays a fundamental role in human and other eukaryotic cells. The NPC possesses remarkable transport competencies with two distinct modes: passive and facilitated (or active) transport [53, 54]. Passive transport is a nonspecific process that involves ordinary diffusion of matter through the nuclear pore with a cutoff at about 10 nm in diameter [50] and/or molecular mass less than 25 kDa [53]. Facilitated transport is the highly specific process acting against concentration gradients [55] with cutoff diameters of approximately 50 nm [50] and a molecular mass (m) range of 25 < m < 75 kDa [53]. This limit easily includes

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the size of most protein transport substrates and macromolecules, for which the molecular weight of the cargo–receptor complex should be in the low hundreds of kilodaltons. In addition, Thachenko et al. [56] have suggested a cutoff size for particles for targeted nuclear delivery; particles should have diameters less than 100 nm to enter the cell membrane and less than 30 nm to be imported through the NPC, respectively. The passive diffusion of the DNA through nuclei in nonmitotic cells has been reported [57, 58]. Salman et al. [57] found that the uptake of DNA, 2 nm in linear dimensions, is independent of ATP or GTP hydrolysis, indicating linear diffusion without the need for conformation change or specific biochemical interaction with nuclear pore complexes (NPCs). The kinetic results, however, show diffusion to be much slower than would be estimated from purely hydrodynamic considerations. This perhaps suggested that the DNA may possible disassemble from its carrier before it passively diffuses through the NPC into the nucleus. Once through the NPC the journey of the DNA is not over. The nucleus is crowded. Lukacs et al. [59] have found that DNA diffusion in the nucleus is negligible—and, unlike diffusion in the cytoplasm, independent of DNA size, in agreement with our data shown in Figure 3.2. Active transport entails nuclear localization signal (NLS) peptides that can be linked to the plasmid DNA; this peptide triggers the transport of the DNA through the NPC by forming a complex with nuclear membrane importers, but such a strategy does not always produce higher transfection levels and can indeed reduce the effect, possibly because the added peptide changes the physical properties of the complex, not least its size or susceptibility to aggregation [60, 61]. The dendrimers and the dendriplexes traverse the cell cytoplasm to the nucleus as can be proposed in the model in Figure 3.4. The 1000- and 2000 fold reduction in diffusion coefficient of the dendrimer and the dendriplexes in the cell cytoplasm can be dissected into two main steps: diffusion within cell cytoplasm and passage through the nuclear pore complex (NPC). The factors involved in the first step are the obstruction and exclusion effects regarding the molecular crowding and the sieving effect mentioned earlier. The cell cytoplasm is depicted here acting as a meshwork, providing a sieving effect with maximum cutoff size at 50 nm in radius (Figure 3.4A). Likewise for a gel, the meshwork structure and crowded state appear as an insoluble compartment through which the dendrimer and the dendriplexes cannot diffuse (Figure 3.4B). Thus the dendrimer and the dendriplexes have to detour in the nonexcluded aqueous volume, until they reach their destination. After endocytosis or passive diffusion through transient nanopores ( – ), the dendrimer (red arrow) and the dendriplexes (green arrow) diffuse toward the nuclear compartment. Some of the dendrimer may interact with cell organelles ( ) and the interaction can be found after endosomal escape of the dendriplexes ( ). Particle size and type of carrier system seem to play a decisive role in the diffusion of the particles in this first step. After endosomal escape, the debate is still whether or not DNA disassembles from the dendrimer and how the DNA reaches the nucleus. Knowledge

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A

Cell radius (h) B 4 1

Dendriplexes Dendrimer

2 3

Cell membrane

Nuclear membrane

FG surface

C 1

Cytoplasmic filaments Threading area

2

Passive transport

Selectivity filter Nuclear filaments

3

Active transport

Figure 3.4. An uptake model for dendrimer and dendriplexes: (A) demonstrates the sieving effect of the cytoplasmic meshwork with a cutoff size of 100 nm in diameter. The crowded condition and the obstruction effect cause the slow diffusion of the dendrimer and dendriplexes after endocytosis ( – ) or passive diffusion ( ) through transient nanoholes, 1–10 nm in diameter, illustrated in (B). The dendrimer and dendriplexes have to detour in the aqueous compartment excluding crowded volumes. Some of the dendrimer and dendriplexes may interact with cell organelles or cytoplasmic proteins ( ). Passing through the NPC, the DNA released from the endosome ( ) may cross the NPC by passive diffusion ( ). As small as 6.5 nm, the dendrimer can diffuse through the central pore having a cutoff size of 8–10 nm ( ), whereas the larger of the dendriplexes, 107 nm, may employ active transport ( ). (Part C – is modified from Peters [50]; other parts are original.) (See color insert.)

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of transport mechanisms through the nuclear pore complexes (NPCs) might help to make this clear. A vertebrate somatic cell usually contains between 1000 and 10,000 NPCs [62]. Transport of the DNA through the NPC is assumed to be the rate-limiting step for gene therapy, a process divided into two strategies: passive and active pathways (Figure 3.4C). Particles with diameters between 8 and 10 nm can be transported by passive diffusion, whereas particles of larger size and particles with molecular masses between 25 and 75 kDa [53] utilize active transport processes. Recent models of the NPC have been proposed as a hydrogel [60, 63, 64], as seen in Figure 3.4C. The selectivity is supplied by the meshwork construction of the phenylalanine glycine (FG) motif and the nonselectivity or passive transport is acquired from the aqueous tube of the channel center, 8–10 nm in diameter. DNA and dendrimers may passively diffuse though this channel (Figure 3.4C – ), while the dendriplexes may have to interact as transport complexes in the perinuclear membrane. These transport complexes are searching for the FG motif in the “threading area,” seen in Figure 3.4C , and they will only selectively passage via FG active transport pathways through the NPC (Figure 3.4C ).

3.6

CONCLUSION

The intrinsically fluorescent lysine-based dendrimer used in our work has proved to be useful as a nanofluorescent probe for tracking uptake and translocation within the cells, as it possesses a fluorescent signal detected by both spectrofluorimetry and confocal microscopy. The uptake of the dendrimer and dendriplexes formed from it depend on the cell type and naturally incubation periods. The dendrimer is taken up into the nuclear compartment. Dendriplexes, because of their size, do so also at a slower rate. One uptake mechanism investigated here was found to be due to endocytic-mediated endocytosis. Other uptake mechanisms have also been proposed. A model of the uptake pathway of the dendrimer and the dendriplexes from the cytoplasm throughout the nuclear pore complexes (NPCs) has been proposed here. Macromolecular crowding in the cytoplasm presents a significant barrier to diffusion of dendrimer and dendriplexes and this has been discussed here in terms of exclusion and obstruction effects. Finally, the uptake of the dendrimer was found to be impeded by the influence of fluid medium flow. The diffusion of the dendrimer and the dendriplexes within the cell nucleus may be expected to be dependent on similar key factors obtained inside the cell cytoplasm, but the nucleus is even more crowded than the cytoplasm. Whether or not the dendriplexes release their DNA before or after passing through the NPC is not known. The transport of the parent dendrimer through the NPC into the cell nucleus is expected to be dependent on passive diffusion. Further investigation to clarify this issue is indispensable so that more successful gene delivery systems can be developed. The main challenge is to determine why there are differences in transfection and diffusion in different cell

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31. Mecke, A., Lee, D. K., Ramamoorthy, A., Orr, B. G., and Holl, M. M. Synthetic and natural polycationic polymer nanoparticles interact selectively with fluid-phase domains of DMPC lipid bilayers. Langmuir 21: 8588–8590 (2005). 32. Mecke, A., Majoros, I. J., Patri, A. K., Baker, J. R., Jr., Holl, M. M., and Orr, B. G. Lipid bilayer disruption by polycationic polymers: the roles of size and chemical functional group. Langmuir 21: 10348–10354 (2005). 33. Morris, M. C., Deshayes, S., Heitz, F., and Divita, G. Cell-penetrating peptides: from molecular mechanisms to therapeutics. Biol. Cell 100: 201–217 (2008). 34. Mislick, K. A. and Baldeschwieler, J. D. Evidence for the role of proteoglycans in cation-mediated gene transfer. Proc. Natl. Acad. Sci. U.S.A. 93: 12349–12354 (1996). 35. Manunta, M., Izzo, L., Duncan, R., and Jones, A. T. Establishment of subcellular fractionation techniques to monitor the intracellular fate of polymer therapeutics II. Identification of endosomal and lysosomal compartments in HepG2 cells combining single-step subcellular fractionation with fluorescent imaging. J. Drug Target. 15: 37–50 (2007). 36. Woodle, M. C. and Scaria, P. Cationic liposomes and nucleic acids. Curr. Opin. Colloid Interface Sci. 6: 78–84 (2001). 37. Manunta, M., Nichols, B. J., Tan, P. H., Sagoo, P., Harper, J., and George, A. J. Gene delivery by dendrimers operates via different pathways in different cells, but is enhanced by the presence of caveolin. J. Immunol. Methods 314: 134–146 (2006). 38. Elouahabi, A. and Ruysschaert, J. M. Formation and intracellular trafficking of lipoplexes and polyplexes. Mol. Ther. 11: 336–347 (2005). 39. Ruenraroengsak, P., Al-Jamal, K. T., Hartell, N., Braeckmans, K., De Smedt, S. C., and Florence, A. T. Cell uptake, cytoplasmic diffusion and nuclear access of a 6.5 nm diameter dendrimer. Int. J. Pharm. 331: 215–219 (2007). 40. Sagidullin, A. I., Muzafarov, A. M., Krykin, M., Ozerin, A. N., Skirda, V. D., and Ignat’eva, G. M. Generalized concentration dependence of self-diffusion coefficients in poly(allylcarbosilane) dendrimer solution. Macromolecules 35: 9472–9479 (2002). 41. Xia, T., Kovochich, M., Liong, M., Zink, J. I., and Nel, A. E. Cationic polystyrene nanosphere toxicity depends on cell-specific endocytic and mitochondrial injury pathways. ACS Nano 2: 85–96 (2008). 42. Kaether, C. and Gerdes, H. H. Monitoring of protein secretion with green fluorescent protein. Methods Enzymol. 302: 11–19 (1999). 43. Bayele, H. K., Sakthivel, T., O’Donell, M., Pasi, K. J., Wilderspin, A. F., Lee, C. A., Toth, I., and Florence, A. T. Versatile peptide dendrimers for nucleic acid delivery. J. Pharm. Sci. 94: 446–457 (2005). 44. Arrio-Dupont, M., Foucault, G., Vacher, M., Devaux, P. F., and Cribier, S. Translational diffusion of globular proteins in the cytoplasm of cultured muscle cells. Biophys. J. 78: 901–907 (2000). 45. Tseng, Y. and Wirtz, D. Mechanics and multiple-particle tracking microheterogeneity of alpha-actinin-cross-linked actin filament networks. Biophys. J. 81: 1643– 1656 (2001). 46. Fushimi, K. and Verkman, A. S. Low viscosity in the aqueous domain of cell cytoplasm measured by picosecond polarization microfluorimetry. J. Cell Biol. 112: 719–725 (1991).

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47. Janson, L. W., Ragsdale, K., and Luby-Phelps, K. Mechanism and size cutoff for steric exclusion from actin-rich cytoplasmic domains. Biophys. J. 71: 1228–1234 (1996). 48. Luby-Phelps, K. Effect of cytoarchitecture on the transport and localization of protein synthetic machinery. J. Cell Biochem. 52: 140–147 (1993). 49. Provance, D. W. Jr., McDowall, A., Marko, M., and Luby-Phelps, K. Cytoarchitecture of size-excluding compartments in living cells. J. Cell Sci. 106(2): 565–577 (1993). 50. Peters, R. Introduction to nucleocytoplasmic transport: molecules and mechanisms. Methods Mol. Biol. 322: 235–258 (2006). 51. Allen, T. D., Cronshaw, J. M., Bagley, S., Kiseleva, E., and Goldberg, M. W. The nuclear pore complex: mediator of translocation between nucleus and cytoplasm. J. Cell Sci. 113(10): 1651–1659 (2000). 52. Rout, M. P., Aitchison, J. D., Suprapto, A., Hjertaas, K., Zhao, Y., and Chait, B. T. The yeast nuclear pore complex: composition, architecture, and transport mechanism. J. Cell Biol. 148: 635–651 (2000). 53. Lusk, C. P., Blobel, G., and King, M. C. Highway to the inner nuclear membrane: rules for the road. Nat. Rev. Mol. Cell Biol. 8: 414–420 (2007). 54. Bickel, T. and Bruinsma, R. The nuclear pore complex mystery and anomalous diffusion in reversible gels. Biophys. J. 83: 3079–3087 (2002). 55. Dingwall, C., Sharnick, S. V., and Laskey, R. A. A polypeptide domain that specifies migration of nucleoplasmin into the nucleus. Cell 30: 449–458 (1982). 56. Tkachenko, A. G., Xie, H., Liu, Y., Coleman, D., Ryan, J., Glomm, W. R., Shipton, M. K., Franzen, S., and Feldheim, D. L. Cellular trajectories of peptide-modified gold particle complexes: comparison of nuclear localization signals and peptide transduction domains. Bioconjug. Chem. 15: 482–490 (2004). 57. Salman, H., Zbaida, D., Rabin, Y., Chatenay, D., and Elbaum, M. Kinetics and mechanism of DNA uptake into the cell nucleus. Proc. Natl. Acad. Sci. U.S.A. 98: 7247–7252 (2001). 58. de Gennes, P. G. Passive entry of a DNA molecule into a small pore. Proc. Natl. Acad. Sci., U.S.A. 96: 7262–7264 (1999). 59. Lukacs, G. L., Haggie, P., Seksek, O., Lechardeur, D., Freedman, N., and Verkman, A. S. Size-dependent DNA mobility in cytoplasm and nucleus. J. Biol. Chem. 275: 1625–1629 (2000). 60. Sakthivel, T., Toth, I., and Florence, A. T. Synthesis and physicochemical properties of lipophilic polyamide dendrimers. Pharm. Res. 15: 776–782 (1998). 61. Toth, I., Sakthivel, T., Wilderspin, A. F., Toth, I., Bayele, H. K., O’Donnell, M., Perry, D. J., Pasi, K. J., Lee, C. A., and Florence, A. T. Novel cationic lipidic peptide dendrimer vectors—in vitro gene delivery. STP Pharm. Sci. 9: 93–99 (1999). 62. Burke, B. Cell biology. Nuclear pore complex models gel. Science 314: 766–767 (2006). 63. Frey, S., Richter, R. P., and Gorlich, D. FG-rich repeats of nuclear pore proteins form a three-dimensional meshwork with hydrogel-like properties. Science 314: 815–817 (2006). 64. Elbaum, M. Materials science. Polymers in the pore. Science 314: 766–767 (2006).

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CHAPTER 4

Endocytosis and Intracellular Trafficking of Quantum Dot–Ligand Bioconjugates TORE-GEIR IVERSEN, NADINE FRERKER, and KIRSTEN SANDVIG Centre for Cancer Biomedicine, Faculty Division, Norwegian Radium Hospital, University of Oslo, and Department of Biochemistry, Institute for Cancer Research, Norwegian Radium Hospital, Oslo University Hospital, Oslo, Norway

4.1

INTRODUCTION

Nanoparticles (NPs) are gaining increasing relevance for use in biological and biomedical applications both within diagnostic imaging and therapeutic drug delivery systems. These applications require the targeting of specific tissues and cells, and important for several of the actions is the uptake of NPs into cells. In vivo and in vitro labeling with NPs allow detection and tracing of molecules into cells as well as cell fractionation. The entry of NPs into a targeted cell is determined and restricted by the endocytic pathways operating. Not all endocytic mechanisms have been identified and characterized so far. The following paragraphs give a short overview of the most studied pathways and introduce important factors and observations relevant in endocytosis and trafficking of NPs.

4.2

DIVERSITY OF ENDOCYTIC MECHANISMS

Endocytosis is a process in which molecules are internalized into the cell interior without passing through the membrane. There are different types of Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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pinocytosis

phagocytosis clathrin-mediated endocytosis

clathrin-independent endocytosis

dynamin-dependent endocytosis

dynamin-independent endocytosis

RhoA = dynamin = clathrin

caveolae

Cdc42

Arf6

Rac ruffling

Figure 4.1. Endocytic pathways. Phagocytosis (left: enclosure of large particle) and different forms of endocytosis—clathrin-mediated endocytosis and clathrinindependent forms such as macropinocytosis, caveolae, RhoA-, Cdc42-, and Arf6mediated endocytosis.

endocytosis—each involving the formation of intracellular vesicles by means of invagination of the plasma membrane and membrane fission. In general, a rough division can be made between phagocytosis (“cell eating”) and pinocytosis (“cell drinking”). Large particles such as bacteria are taken up via phagocytosis. The particle is engulfed by extrusions from the cell membrane (a “phagocytic cup”), which then mediates the formation of a phagosome. Phagocytosis occurs mainly in specialized mammalian cells such as macrophages, monocytes, and neutrophils but also to a lesser extent in nonprofessional phagocytes (i.e. fibroblasts, endothelial and epithelial cells). Phagocytosis constitutes the initial step for the degradation of particles larger than 0.5 μm. Pinocytosis is used to internalize fluid simultaneously with whatever substance is found within the area of invagination. There are multiple types of endocytotic pathways, which can be classified in part based on their requirement for the coat protein clathrin, the GTPase dynamin, or the formation of caveolae. Thus a general division into clathrindependent/independent and dynamin-dependent/independent endocytic pathways can be made (Figure 4.1). 4.2.1

Clathrin-Dependent Endocytosis

The clathrin-dependent endocytosis is one of the best characterized endocytic pathways and constitutes a major route for selective receptor internalization

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in eukaryotic cells [1]. The formation of a clathrin-coated pit starts with adaptor protein-mediated recruitment of clathrin to the plasma membrane, where the adaptor proteins bind transmembrane receptor molecules with cargo. Finally, the coated pit invaginates and, regulated by the GTPase dynamin and GTP, pinches off to form a clathrin-coated vesicle. In general, clathrin-mediated endocytosis (CME) is a rapid uptake process (<2 min). 4.2.2

Clathrin-Independent Endocytosis

A number of clathrin-independent pathways that can be classified upon involvement of dynamin and various other GTPases have been detected so far. A classification scheme based on dependence on the small Ras-superfamily GTPases RhoA, Cdc42, and Arf6 and the formation of caveolae has been suggested [2]. The GTP-binding protein dynamin is required in caveolar as well as in RhoA-regulated endocytosis. Caveolae are flask-shaped plasma membrane invaginations of 50–100 nm in diameter that are enriched in sphingolipids, cholesterol, and the transmembrane protein caveolin [3]. Although caveolin-1 has been shown to be important for the formation of caveolae, morphologically similar structures to caveolae have been observed in some cells that lack caveolin proteins [4, 5]. Interestingly, overexpression of flotillin1 and flotillin2 in HeLa cells was recently reported to generate caveolae-like structures [6]. Caveolae are mostly static structures but they can be induced to pinch off by certain ligands such as SV40 [7, 8]. Moreover, less characterized endocytic pathways exist sharing some of the above mentioned characteristics or none of them [9, 10]. In this context, the existence of polymorphous structures with an elongated and tubular shape that endocytose glycosphingolipids and other plasma membrane components have been described [5]. Similar structures can be induced by the Shiga toxin B subunit [11]. An endocytic mechanism involving tubular structures (CLICs, clathrin-independent carriers) has been proposed [12, 13], and recently GRAF1 was identified as specific marker for CLICs [14]. However, the picture is not clear since it has been reported that CLICs can form without dynamin whereas GRAF1 is able to bind dynamin. Macropinocytosis, a special form of pinocytosis, is one of the best characterized types of clathrin-independent endocytosis, and it is often preceded by the actin-driven process of membrane ruffling, which can be stimulated by growth factors (for recent reviews see Mercer and Helenius [15] and Kerr and Teasdale [16]). In macropinocytosis, the cell engulfs mass quantities of fluid after formation of protrusions that can be lamellipodia-like, bleb-shaped, or appear in the form of circular ruffles. These protrusions may collapse onto and fuse with the plasma membrane. After membrane fission, large noncoated vesicles of heterogeneous sizes (0.5–10 μm in diameter) and irregular in

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shape—the macropinosomes—are formed. Macropinocytosis depends on several kinases such as p21-activated kinase 1 or protein kinase C [17], but also Rho family GTPases such as Rac1 [18], which plays an important role in remodeling of the actin cytoskeleton, can be involved.

4.3 UPTAKE OF NANOPARTICLES OF VARIOUS SIZE AND SURFACE CHARGE Endocytosis mechanisms involved in uptake of nanoparticles of varying sizes and compositions have been investigated in a number of studies relying mainly on inhibitors that are supposed to perturb a specific endocytic pathway. It has been shown that anionic polymeric particles of 24 nm were endocytosed in HeLa and primary HUVEC cells via a cholesterol-independent, non-clathrinand non-caveolae-dependent pathway, which did not route them to the degradative pathway, whereas particles with a size of 43 nm were mainly internalized via clathrin-dependent endocytosis and further directed to the endolysosomal pathway, suggesting that cells can precisely distinguish and accordingly sort these particles via different uptake mechanisms [19]. In another study, Hoekstra and co-workers examined the uptake mechanisms of latex beads of various sizes (50, 100, 200, 500, and 1000 nm) in the nonphagocytic mouse cell line B16 by using inhibitors of either clathrin-dependent or caveolae-dependent endocytosis to discriminate between these two pathways. They found that beads with a diameter below 200 nm were internalized via clathrin-mediated endocytosis (CME), although at slower rates for the larger particles. Surprisingly, considering the restricted size of caveolae, they reported that a caveolae-dependent mechanism was the predominant endocytic mechanism for particles of 500 nm, whereas no uptake could be observed with particles of 1000 nm size [20]. The glycosphingolipid (GSL) lactocyl-ceramide (LacCer) was used as a fluorescent marker for caveolae and was found to colocalize with the 500-nm nanoparticles. However, one should be cautious when using Bodipy-LacCer as a marker for caveolae since it has been shown that exogenously added GSL can cause formation of GSL-enriched lipid domains at the plasma membrane and induce internalization of integrins via caveolae-like structures [21]. DNA lipoplexes containing the cationic lipid DOTAP were exclusively internalized by CME, while polyplexes prepared from the cationic polymer polyethylene imine (PEI) were internalized equally by both clathrin-dependent and clathrin-independent pathways. However, luciferase transfection by the PEI/DNA-polyplexes was blocked by inhibitors such as genistein and filipin, and the result was interpreted as involvement of caveolae. However, it should be noted that filipin can be expected to inhibit any cholesterol-dependent internalization, and genistein is an inhibitor of tyrosine phosphorylations that will affect uptake via other pathways as well, for instance, uptake through clathrin-coated pits when

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receptor phosphorylations on tyrosine are involved. Interestingly, the fraction internalized via the caveolae-mediated pathway was not directed into lysosomes [22, 23]. In general, particles with positive surface charge will bind to the negatively charged membrane proteins and lipids at the cell surface and are therefore likely to be easily captured and taken up. Thus the surface charge of the NPs could be decisive for endocytosis and trafficking. A study of endocytosis of cationic and anionic polymeric NPs with a similar size (80 nm) in HeLa cells showed that the cationic NPs displayed a twofold higher endocytosis rate compared to the anionic NPs [24]. In addition, cationic beads were internalized via CME, whereas internalization of anionic beads was shown to be independent of clathrin and caveolin. Suprisingly, upon inhibition of dynamin-dependent endocytosis by expression of dominant negative mutant dynamin, there was an increase in endocytosis especially of the cationic NPs, which was attributed to the induction of macropinocytosis previously reported in these cells [25]. In another study, Harush-Frenkel et al. [26] found that, in polarized MDCK cells, anionic as well as cationic polymeric nanoparticles mainly entered the apical pole of these cells via CME. Furthermore, a significant fraction of the NPs was also transcytosed to the basolateral pole. Recently, it was suggested that uptake of charged polystyrene NPs (120 nm) in HeLa cells occurred via a dynamin-dependent and lipid-raft-dependent pathway independently of their surface charge [27]. Macropinocytosis seemed to be involved in uptake of the cationic NPs due to the inhibiting effect of 5-(N-ethyl-N-isopropyl)amiloride (EIPA). Surprisingly, cholesterol depletion and filipin treatment did not inhibit uptake of the NPs, which contradicts previous studies demonstrating that cholesterol is involved both in macropinocytosis and lipid-raft-dependent uptake. Chlorpromazine was used as an inhibitor of CME and resulted in a minor inhibition in the uptake of positively charged NPs, but had no effect in uptake of the negatively charged NPs. The effect of surface charge on the uptake of mesoporous silica NPs (MSNs) has been studied in 3T3-L1 cells and mesenchymal stem cells (hMSCs) [28]: In both cell types, uptake of the anionic NP was inhibited by phenyl arsene oxide and cytochalasin-D, suggesting involvement of clathrinand actin-dependent endocytosis. With the strongly cationic MSNs, the inhibitory effects on uptake were only observed in the 3T3-L1 cells but not in the hMSCs, implying that the effect of NP surface charge is dependent on cell type. The endocytic pathway followed by a nanoparticle without bound ligands seems to depend on its size and surface charge. Macropinocytosis seems to be a preferred pathway for positively charged nanoparticles. However, endocytosis mechanisms followed by a certain nanoparticle apparently vary between cell lines and types depending on the endocytic pathways operating (Table 4.1). Thus most cell lines seem to have different endocytic mechanisms available for internalizing nanoparticles (<200 nm).

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60 TABLE 4.1 NP Type

Endocytosis Studies on Different Nanoparticles Including QDs Charge (+/−)

Size (Diameter, nm)

Ligand

Cell Line

Reported Endocytosis Mechanisms or Intracellular Localization

mPeg-PLA

+32 mV −26 mV

Polystyrene

+ −

120 120

HeLa HeLa

Silica

+ −

110 110

Polystyrene

− − − −

24 45 and 200 11 2

3T3-L1 hMSC 3T3-L1, hMSC HeLa, HUVEC HeLa, HUVEC HeLa Macrophages

CME: inhibited by dynamin and clathrin mutants, then macropinocytosis; clathrin/ dynamin-independent Dynamin/actin-dependent, clathrinindependent, macropinocytosis Dyn/actin-dependent, clathrin-independent Clathrin/actin-dependent Clathrin/actin-independent Clathrin/actin-dependent in both cell types Clathrin/caveolae-independent pathway Clathrin-dependent Significant lysosomal localization Transport into the nucleus

−

6 25–30 45

A431 HeLa, HEp-2, SW480

Accumulation in the cytoplasm CME CME, inhibited by dynamin, no recycling but endosomal/lysosomal accumulation

CdSe QDs CdTe QDs

CdSe QDs CdSe QDs

90 96

Stearyl-amine

–COO− COO− DHLA Thioglycolic acid (TGA) TGA EGF Tf

HeLa dyn K44A

Reference 24

27

28

19 Here 29

30 31, 32

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61

4.4 USE OF PHARMACOLOGICAL INHIBITORS AND MICROSCOPY IN STUDIES OF ENDOCYTOSIS Most studies on the various endocytosis mechanisms responsible for membrane and ligand internalization have so far involved the use of pharmacological inhibitors. It is important to stress that many of the inhibitors used in these studies are not specific for one endocytic mechanism. Genistein is an inhibitor of several tyrosine kinases and thus has been shown to inhibit internalization of ligands via clathrin-dependent endocytosis, in addition to affecting changes required for pinching off of caveolae. Furthermore, perturbing the function of cholesterol either by using filipin or by cholesterol depletion with methylβ-cyclodextrin has been shown to affect other endocytic pathways beside the caveolae-mediated ones. Interestingly, it has been demonstrated that cholesterol is required not only for raft-dependent endocytosis, giving rise to small vesicles, but also for macropinocytosis [33]. More robust conclusions should be obtained by using a collection of inhibitors, mutant proteins, and siRNAs. Internalization of fluorescently labeled NPs into cells has mostly been studied by confocal microscopy, which has a limited resolution. Nanoparticles might be forming clusters at the cell’s surface, giving rise to a spotted and vesicular pattern, and upon imaging of flat adherent cells it might be difficult to judge whether a spot represents a true vesicle even from a z-stack of confocal images. Therefore counterstaining with various endosomal markers and measuring the degree of colocalization are generally helpful in resolving whether the NPs have been internalized. Moreover, an apparent colocalization can be obtained from vesicles in close proximity, especially in the thicker perinuclear area of the cell, and should not necessarily be interpreted as localizing to the same organelle.

4.4.1

Endocytosis of Fluorescent QDs Studied by Microscopy

The production of compact functionalized QDs that are stable in serum/ blood and display little unspecific binding to cells is a prerequisite for their use as probes to target specific cells in vivo. Using the intrinsic lipophilic QDs in biological applications requires further modification of their surface to make them soluble in aqueous solution. This can be achieved by absorption of various amphiphilic polymers to the QDs or by exchange of the native capping agent with thiolated molecules such as peptides [34], cysteine [35], or dihydrolipoic acid (DHLA) [36]. The capping molecules might contain functional groups, carboxyls, or amines, for further covalent attachment to the QD surface of biological molecules such as proteins, peptides, or nucleic acids required to achieve the desired targeting.

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4.4.2

ENDOCYTOSIS AND INTRACELLULAR TRAFFICKING

Endocytosis of QDs Without Ligands that Bind to Receptors

Once the QDs are made hydrophilic and stable in aqueous media, they can bind to cells through electrostatic interactions with molecules at the cell surface. The cell surface displays an anionic charge, and therefore cationic NPs in general bind more strongly to the cells. Despite an overall electrostatic repulsion to the cell surface, anionic dihydrolipoic acid (DHLA)-capped CdSe/ZnS QDs at a high concentration (300 nM) have been found to bind unspecifically to cells and be endocytosed [37]. Recent studies have shown that unspecific binding to cells could be greatly reduced by attaching a PEG-spacer to DHLA while keeping a compact size of the QDs (<15 nm) [38]. Recently, we also monitored significant endocytosis of DHLA-capped QDs (hydrodynamic diameter, diamh, 11 nm) into HeLa cells after 1 h at a ∼tenfold lower concentration (30 nM) (Figure 4.2). After incubation for 1 h at 37 °C the particles displayed partial colocalization with the early endosomal marker EEA1 (Figure 4.2, upper panel), and after 4 h incubation they displayed a

DHLA-QDs 1h:

EEA1

CD63 merged

EEA1

CD63 merged

10 μm

DHLA-QDs

10 μm

10 μm

4h:

10 μm

10 μm

10 μm

Figure 4.2. Intracellular location of dihydrolipoic acid (DHLA)-capped quantum dots (QDs, Em. 585 nm) in HeLa cells. The DHLA-QDs (30 nM) in complete growth medium were added to the HeLa cells and incubated at 37 °C for 1 and 4 h, respectively. Yellow and magenta colors in the merged images indicate colocalization with the early endosomal marker EEA1 and with the lysosomal marker CD63, respectively. Bars, 10 μm. (See color insert.)

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significant increase in colocalization with the lysosomal marker CD63 (Figure 4.2, lower panel). Interestingly, it has recently been demonstrated that differences in cancer cell phenotypes can lead to significant differences in intracellular sorting, trafficking, and localization of nanoparticles. Unconjugated anionic QDs displayed dramatically different intracellular profiles in three closely related human prostate cancer cells [39]. Moreover, hypoxia conditions in tumor cells have been show to change intracellular trafficking and sorting of receptor tyrosine kinase (RTK) receptors, delaying their transport down the endolysosomal pathway and thus their deactivation [40]. Furthermore, in a study of cellular uptake of tiny thioglycolic acid-capped CdTe QDs, Nabiev et al. [29] observed endocytosis in macrophages only, and not in epithelial or endothelial cells. Moreover, they reported that the intracellular distribution of the QDs varied with particle size, in that the smallest QDs (2 nm) seemed to target histones in the nucleus, whereas the larger QDs (6 nm) remained in the cytoplasm [29].

4.4.3

Endocytosis of Ligand-Bound QDs

In order to achieve the desired targeting to specific cells, various peptides, proteins, or nucleic acids can be coupled to the QDs. Another key issue to address is whether the functionalized QDs can act as relevant intracellular probes to investigate the routing of ligands in live cells. Except for a few recent studies, little information has been published about the mechanisms of intracellular trafficking of different Qdot-bound ligands [41–43]. QDs bound to receptors such as glycine and epidermal growth factor (EGF) receptors have been used to measure and resolve the dynamics of single receptors at the cell surface. Biotinylated epidermal growth factor has been conjugated to streptavidin-coupled QDs for the binding to and activation of EGF receptors at the cell surface [30]. Importantly, a retrograde transport of EGF-QDs from filopodia to the cell body was observed. Furthermore, the EGF-QDs were rapidly endocytosed and the internalized receptor dynamics were followed over time. Dahan and co-workers used QDs to track individual glycine receptors (GlyRs) and analyzed their lateral dynamics in neuronal membrane cells [44]. Very recently, the rate of lateral diffusion of individual glycosyl-phosphatidyl-inositide anchored proteins coupled to avidin (Av-GPI) has been studied by labeling with QD and using dual-color total internal reflection fluorescence (TIRF) microscopy [45]. It was shown that the Av-GPI probe exhibited a fast and a slow diffusion regime in different membrane regions, and that cholesterol-/ sphingolipid-rich microdomains can compartmentalize the diffusion of GPIanchored proteins. Internalization of the transferrin (Tf) bound iron by the transferrin receptor is the major route of iron uptake. The complexes are constitutively

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endocytosed by clathrin-dependent endocytosis, and after release of iron in an acidic endosome, the Tf bound to its receptor very efficiently recycles from the sorting endosomes to the plasma membrane. Tf has also been investigated as a ligand for targeted delivery of therapeutic agents that suffer from nonoptimal pharmacokinetic properties [46]. Tf has been used both by us and by others to facilitate uptake of QDs and other NPs into various cells [47, 48]. In a recent study [31], we coupled biotinylated Tf to commercially available streptavidin QDs resulting in bioconjugates with a hydrodynamic diameter (diamh) of 40 nm. The Tf : QD bioconjugates specifically bound to the transferrin receptors and were internalized via dynamin-dependent endocytosis. The Tf : QDs displayed good colocalization with the early endosomal marker (EEA1) already after 15 min of endocytosis. After chasing of their endocytosis for 3–4 h they accumulated in more perinuclear endosomes, displaying only partial colocalization with lysosomal markers such as LAMP-1 and CD63. Endocytosis of Tf : QDs was also studied in two more cancer cell lines, HEp-2 cells and SW480, where the Tf : QDs displayed a more significant (40%) colocalization with the lysosomal marker after 3 h of endocytosis [32]. Thus, contrary to transferrin itself, they did not recycle to the plasma membrane. This endosomal retention outside lysosomes can be considered advantageous in relation to the issue of drug delivery and avoiding lysosomal degradation.

4.4.3.1 Toxin : QD Conjugates Several protein toxins have cytosolic targets, and studies of their uptake and intracellular transport have provided knowledge about mechanisms used to gain access to the cytosol as well as details about endocytosis mechanisms. Some toxins such as diphtheria toxin escape from endosomes in response to low endosomal pH, whereas Shiga toxin and ricin are transported all the way back to the Golgi apparatus and the ER before they are translocated into the cytosol. In fact, retrograde transport to the Golgi and ER is necessary for the toxins to reach the cytosol, where they exert their toxic effect [49]. If toxin : QD bioconjugates can be transported to the Golgi apparatus and the ER, this would facilitate studies of retrograde transport. So far, one has not been able to visualize ricin in the ER. Thus if the toxin : QDs could mediate transport to such destinations and be monitored there by live-cell microscopy, it would be a step forward in clarifying further details about endocytic and retrograde transport. Shiga toxin B subunit binds to the globotriaosyl ceramide Gb3 receptor, which is expressed and binds toxin on the cell surface of the kidney endothelial cells of children [50]. Gb3 is also overexpressed at the cell surface of various metastatic cancer cells, making the Shiga B subunit an interesting ligand for specific targeting of cytostatika to these cells and for imaging [51, 52]. Ricin B subunit binds to glycoproteins and glycolipids containing a terminal galactose moiety present at the cell surface of nearly all mammalian cells. Thus it

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has been used to facilitate intracellular delivery of other molecules, such as immunotoxins. In our previous studies, we reported that the ricinB : QDs (diamh, 30 nm) were internalized and partially routed into perinuclear vesicles, displaying a significant overlap with lysosomal structures in HeLa, HEp-2, and SW480 cells after 3 h of endocytosis [31, 32]. However, another more dispersed fraction of these ricinB : QD-positive structure did not stain positive for lysosomal markers. Furthermore, whereas a fraction of ricin can be visualized in the Golgi apparatus after 3 h of endocytosis, we did not observe any colocalization of the ricinB : QDs with the Golgi markers used. The ricinB : QDs remained accumulated in these endosomes for 24 h. Cholera toxin subunit B-QD conjugates (diamh, 25 nm) have been found to be internalized into dispersed vesicles of various cells, where they also were retained for days [53]. Here, we show internalization of ricinB : QDs (molar ratio, 5 : 1) in HeLa cells and HEp-2 cells, respectively (Figure 4.3). A fraction of the ricinB : QDs was routed into LAMP-1 positive endosomes after 90 min of endocytosis in both cell types, and the degree of colocalization between ricinB : QDs and LAMP-1 was similar after 180 min. However, after internalization for 24 h a decrease in overall ricinB : QD fluorescence was observed in both cell lines. Moreover, the ricinB : QDs displayed less colocalization with LAMP-1. This suggests that the lysosomal content with accumulated ricinB : QDs was slowly secreted [54, 55], or that there was a certain extent of lysosomal and oxidative degradation of the QD coats resulting in aggregation of the particles and thereby to a decrease in their photoluminescence [56]. 4.4.3.2 Antibodies QDs coupled to antibodies or a humanized antibody fragment against cell surface proteins that are endocytosed is another way to deliver the QDs into the cell. This approach has been especially appealing for detection of cancer cells with a cancer-associated overexpressed membrane marker [57, 58]. 4.4.3.3 Peptide-Mediated Endocytosis Instead of using entire proteins as ligands, peptides can be bound to the QDs, for example, constituting the receptor-binding motif of the ligand. The TAT peptide motif derived from the HIV-1 virus TAT protein has proved to be a popular targeting molecule to deliver cargo into cells. It is a highly positively charged ligand that interacts with negatively charged receptors at the cell surface. The TAT peptide has been self-assembled onto the surface of DHLA-QDs [59]. The TAT-peptide-coupled DHLA-QD conjugates were endocytosed efficiently within a 1-h period and accumulated within endosomes for 24 h without causing any cell death. Cellular uptake of TAT peptide coupled to the end groups of PEG-capped QDs has been used to label mesenchymal stem cells that subsequently were injected into nude mice and tracked to the liver, lung, and spleen [60].

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Figure 4.3. Intracellular location of ricinB : QD655 bioconjugates in HeLa cells (left panel) and HEp-2 cells (right panel). The ricinB : QDs were constituted and bound to the cell surface at 4 °C, and then allowed to be internalized into the cells for various times at 37 °C. The cells were then fixed with 4% paraformaldehyde and prepared for IF microscopy imaging by labeling them with a rabbit antibody against LAMP-1 and with a corresponding secondary anti-rabbit-Cy2 conjugate. Images show ricinB : QD conjugates in red, LAMP-1 in green, and the merged z-stack. Yellow in the merged images indicates colocalization. As seen in the z-stack, the main fraction of QD conjugates is still located at the cell surface in HeLa as well as in HEp-2 cells after 30 min. RicinB : QDs show increasing uptake and partial colocalization with LAMP-1 after longer incubation (90–180 min). RicinB : QD staining is significantly reduced after 24 h of incubation in HeLa and even more in HEp-2 cells. (See color insert.)

4.5 DISTURBANCES IN CELLULAR PROCESSES INDUCED BY LIGAND–NANOPARTICLE BIOCONJUGATES Most of the ligand–nanoparticle bioconjugates studied contain a multivalent binding of ligands to the nanoparticle. In addition to the resulting increase in binding avidity, this can definitely also trigger adverse cellular effects. Multivalent ligand–QD conjugates might cause crosslinking of cell surface receptors and thereby trigger cell signaling pathways. In a recent study by Howarth et al. [61], they prepared compact QDs with monovalent binding of streptavidin. They observed that ephrin receptors EphA3 were evenly distributed at the cell surface upon binding of the monovalent-QDs without triggering of EphA3 kinase activation, whereas the multivalent-QDs lead to EphA3 receptor clustering and tyrosine kinase signaling resulting in their endocytosis [61]. In our study of Tf : QDs and ricinB : QDs we prepared a substoichiometric derivatization of the bioconjugates by varying the ligand : QD molar ratio from 1 : 1 up to 20 : 1 [31]. The ligand : QDs with a 1 : 1 ratio displayed weaker fluorescent intensities when bound to the cells compared with the ligand : QDs of a 5 : 1 ratio. However, we could not observe any differences in their degree of endocytosis or intracellular routing. It has been found that multivalent binding of ricin to HRP and gold nanoparticles directed ricin to lysosomes, whereas a monovalent binding also directed ricin transport into the Golgi apparatus [62]. Furthermore, endocytosis of polyvalent IgG to the Fc receptors of macrophages directed the IgG-Fc receptor complexes to lysosomes for degradation, whereas a monovalent antireceptor antibody caused the endocytosed Fc receptors to recycle [63, 64]. Importantly, we demonstrated that accumulation of the ricinB : QDs perturbed the intracellular transport of the unconjugated ligands ricin and Shiga toxin to the Golgi apparatus. Transport to the Golgi apparatus of ricin was severely inhibited, whereas transport of Shiga was increased. Thus in addition to measuring the typical cytotoxic effects (effects on mitochondria, protein

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synthesis, apoptosis, etc.), it seems important to investigate their potential disturbances of cellular activities related to cell signaling and intracellular trafficking, for instance downregulation of growth factors.

4.6

PERSPECTIVES

Many questions remain open about the mechanisms of endocytosis and their involvement in QD/nanoparticle entry into cells. Precise answers are difficult to give as long as there are still a number of endocytic pathways that are incompletely characterized. Furthermore, the endocytic pathways involved in cellular trafficking of NPs will probably also depend on the cell types studied. Moreover, the possibility of tracking single QD–ligand molecules makes them valuable tools for identifying and resolving the specific endocytic pathways involved. A main objective is to develop QD-labeled ligands that can serve as relevant fluorescent probes for assessing the detailed itinerary of the ligands. However, the large sizes (>20 nm) of many QDs encapsulated within polymers may limit the access into the relevant intracellular sorting stations, and thereby alter the normal trafficking and activity of the ligand. Thus it is important to continue the development of stable QD–ligand conjugates with a small size.

4.7

CONCLUSION

The study of membrane protein dynamics at the molecular level in living cells has become a major issue in understanding important cellular processes such as endocytosis, intracellular transport, and signal transduction. The semiconductor quantum dots (QDs) with their remarkable fluorescent and colloidal properties have potential as probes for membrane proteins at the single molecule level. Furthermore, the rational development of drug delivery and contrast agents based on nanoparticles requires a deeper understanding of particle internalization mechanisms to target cellular pathways capable of providing the required response. Herein, we provided a brief overview of the various endocytosis mechanisms (clathrin-mediated, caveolar, dynamin-independent: noncaveolar, macropinocytosis, phagocytosis) operating in the cell, and discussed studies on endocytosis of different biologically active nanoparticles and QDs varying with respect to size, surface charge, and the ligands conjugated to them.

ACKNOWLEDGMENT This work was supported by Grant 172663/S10 to TGI from the FUGE and NANOMAT programmes of the Research Council of Norway.

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CHAPTER 5

Synthesis of Metal Nanoparticle-Based Intracellular Biosensors and Therapeutic Agents NEIL BRICKLEBANK Faculty of Health and Wellbeing, Biomedical Research Centre, Sheffield Hallam University, Sheffield, United Kingdom

5.1

INTRODUCTION

The emergence of nanotechnology as an independent field of research in the late 1980s and early 1990s has led to a surge in interest in the synthesis of nanoscale (1–100 nm) particles of metals, metal oxides, and semiconductors (also known as quantum dots), due to their technological potential as electronic, magnetic, and photonic materials, as catalysts and as substrates for a host of biological applications including diagnostics and transport vectors. However, metal nanoparticles or colloids are not a new phenomenon; they have been known for many thousands of years, albeit unrecognized as such, and utilized primarily as pigments for decorative purposes. During the Middle Ages aqueous solutions of gold colloids gained a reputation as a remedy for a variety of illnesses, a use which, somewhat paradoxically, anticipated the aims of much current research in the area of nanobiotechnology [1]. Many biomolecules have size dimensions in the range 5–200 nm, which means that materials science and biotechnology converge at the nanoscale. The properties of nanoparticles that make them attractive for biotechnological applications result from a number of attributes including size, shape, and surface morphology. The physical properties of nanoparticles are very different from those of the parent bulk materials and are more complex than those of the individual constituent atoms. These differences arise through the small size and large surface area of the particles; for a 10-nm diameter particle about Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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15% of the atoms occupy “surface” positions, compared with 0.0015% for a 1-μm (1000-nm) particle. Such is the volume of published work that a review such as this cannot comprehensively cover the field of metal nanoparticle synthesis. In this chapter we will concentrate on providing an overview of recent studies, with the first part describing the primary routes for the synthesis of metal nanoparticles, functionalized with organic molecules. The latter sections of this chapter will overview the applications of functional metal nanoparticles as agents for labeling and detection and as therapeutics. The final section addresses the issue of toxicology of nanoparticles, an area of particular relevance to pharmacology.

5.2

SYNTHESIS OF METAL NANOPARTICLES

A wide variety of routes have been developed for the synthesis of metal nanoparticles. The main methods can generally be categorized as wet chemical reduction of metal salts, thermal degradation of metal complexes, or electrochemical synthesis. Less common routes include photochemical decomposition, laser ablation, and biological routes [2]. The latter are of interest because they rely on the fact that certain species of plants and microorganisms are able to uptake large quantities of metallic ions, which they reduce and precipitate as nanoparticles [3]. The shape, size, and particle size distribution of nanoparticles are dependent on many variables, including the reaction temperature and time, solvent, concentration, and nature of the precursors and reagents, particularly the capping ligand. Alteration of any of these factors can lead to a change in the size or shape of the resulting particles [4]. 5.2.1

Role of the Capping Ligand

The capping ligand is a crucial component in both the synthesis and subsequent use of metal nanoparticles. During the synthesis of nanoparticles the capping ligand helps to stabilize the particle, preventing uncontrolled growth and agglomeration. The choice of capping ligand influences the size and shape of the resulting particles. Many different species have been employed including organic bases, surfactants, and polymers. Other approaches for controlling the growth of nanoparticles and templating their morphology include the use of gels, micelles, zeolites, or layered minerals. However, among the most frequently used species are organic thiolates (RS−, where R = alkyl or aryl), usually derived from the corresponding thiols or disulfides. Other common ligands include tertiary phosphines or alkyl amines. Mixed ligand systems offer a further opportunity (and level of complexity) to control or template the formation of particles with a specific size or morphology. The resulting species are often referred to as monolayer protected clusters (MPCs) or, when two

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or more ligands are attached to the surface, mixed monolayer protected clusters (MMPCs). Polymers have also been used for the synthesis of biocompatible nanoparticles and inorganic silicon-based coatings have also been utilized, although molecular organic species predominate. In addition to their stabilizing role, the capping ligands can be used to functionalize the particles, enabling them to be used as sensors, for biomolecular recognition, or as transport vectors. Functionalization can be achieved by a number of ways. The organic groups that are attached to the particle during the initial synthesis can be modified by subjecting the particle to further chemical protocols. Alternatively, the particles can be used to form conjugates with biomolecules. The capping ligands that are utilized in the synthesis stage can be displaced by other species that exhibit the required functionality or biocompatibility through ligand exchange procedures. The latter is attractive for biological species that would not survive the frequently harsh conditions employed in traditional “wet chemistry” procedures. Organic thiolates have been widely used as capping ligands, particularly for gold nanoparticles, because of the high affinity of the sulfur for gold and the high stability of the resulting S–Au bonds. The nature of the bond formed between the donor atom of capping ligand and the surface of nanoparticle has been the subject of much investigation and speculation! However, a significant breakthrough came in 2007 with the reporting of the single crystal X-ray diffraction study of a sample of monodisperse gold particles functionalized with mercaptobenzoic acid (MBA) [5]. The structure revealed a core containing 102 gold atoms that is surrounded by 44 MBA ligands. The gold core comprises 89 Au atoms, 49 of which are arranged in a Marks decahedron with two 20-atom caps. The remaining 13 atoms form a band around the equator of the core. A significant finding was that the MBA ligands not only bind to the gold through Au–S bonds but also interact with each other through a series of face-to-face and face-to-edge interactions between adjacent phenyl rings and also phenyl–sulfur interactions. The interligand interactions observed in this system may provide an indication of how capping ligands are ordered on the particle surface. The significance of this discovery lies in the fact that it enables us to consider metal nanoparticles as “molecular” species, with a clearly defined chemical formula, rather than the more abstract images that have been hitherto accepted. Furthermore, it will aid our understanding of the formation and reactivity of functionalized metal nanoparticles and how these can be manipulated for the benefit of biotechnology. 5.2.2 Synthesis by Direct Chemical Reduction of Metal Salts Arguably the simplest route for the preparation of metal nanoparticles is the direct reduction of metal salts with a suitable reducing agent and/or a capping ligand; in some cases the capping ligand also acts as the reducing agent (Figure 5.1). The best example of this route is the citrate reduction of gold(III) chloride salts in water, the citrate forming a loosely bound shell around the

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AuCI4

(C)

(A)

(i) p-HOC6H4SH,MeOH (ii) NaBH4,H2O

Sodium citrate, H2O, Δ

(B) (i) HAuCl4·H2O (ii) Oct4N+Br–,PhMe (iii) CH3(CH2)11SH, (iv) NaBH4,H2O

–

CO2

– –

OH

O2C

OH Au

–

O2C

O2C – O C 2 –

–

CO2

O2C

CO2

HO

S

S S S

S Au

S

OH

–

OH

CO2

S

S S

CO2 CO2

OH

HO –

HO

OH

HO

–

CO2

S

OH

HO

–

–

HO

OH

S SSS S S S Au S S S S SS S S

Figure 5.1. Common routes for the synthesis of gold nanoparticles: (A) citrate reduction, (B) Brust–Schiffrin two-phase system, and (C) Brust–Schiffrin one-phase system.

gold core. This method was first reported by Turkevitch in 1951 and leads to gold nanoparticles of approximately 20 nm in diameter [6]. The method was refined by Frens, who found that the size of the gold nanoparticles could be controlled by varying the ratio of gold salt and citrate [7]. Similar strategies can be used to prepare nanoparticles of other metals; for example, silver nitrate is readily reduced by sodium borohydride or citrate in water producing relatively monodisperse silver nanoparticles, about 27 nm in diameter [8, 9]. The silver particles produced using this route display excellent stability and their advantageous optical properties, which result from their surface plasmon resonance, have been used in analytical science in SERRS (surface enhanced resonance Raman spectroscopy) [9]. One point worth noting is that the nanoparticles produced using these routes generally display a comparatively broad particle size distribution. Production of particles with a single size and shape remains a challenge.

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A significant breakthrough came through the seminal work of Brust and Schiffrin who devised a method for producing gold nanoparticles with good polydispersity, which are very easily handled and can be isolated and redissolved in common organic solvents without aggregation or decomposition [10]. The initial method utilized an aqueous-toluene two-phase system with dodecanethiol as the capping ligand, sodium borohydride as the reducing agent, and tetraoctylammonium bromide as a phase transfer agent. The colorful reaction mixture changes from orange to dark brown upon addition of the reducing agent. The size of the particles can be varied by modifying the relative amounts of capping ligand and gold salt; the greater the amount of capping ligand the smaller the average particle size. Varying the temperature of the reaction also affects the particle size. Brust and co-workers refined the initial method by developing a single-phase method that obviated the need for the phase transfer agent, which can be difficult to remove from the metallic particles produced using the two-phase method [11]. Subsequently, many workers have adopted the Brust–Schiffrin method and have prepared gold nanoparticles functionalized with a huge variety of thiolate-derived capping ligands, and the methodology has been extended for the production of metallic particles of other metals [1]. The use of surfactants is another common route for the production of nanoparticles. Jana and Peng have reported a surfactant-mediated protocol for the gram-scale synthesis of monodisperse metal nanoparticles [12]. In their procedure, the metal salt (gold(III)chloride, silver acetate, copper acetate, or platinum chloride) was dissolved in toluene together with didodecyldimethylammonium bromide. Fatty acids, aliphatic amines, or thiols were added as stabilizing ligands and play a role in dictating the particle size. The reduction was effected by tetrabutylammonium bromide either alone or with hydrazine. Metal nanoparticles produced using this route cannot be used directly in biomedical applications. However, the particles can be precipitated from toluene solution and the ligands exchanged for water-soluble, biocompatible species. 5.2.3

Reduction of Organometallic Complexes

The reduction of low valent organometallic complexes has been widely used for the preparation of nanoparticles of semiconductor materials such as zinc sulfide and cadmium chalcogenides [13]. Reduction is achieved by chemical, thermal, or photochemical decomposition of the organometallic complex. The nanoparticles produced are often of high purity with very closely defined, and reproducible, sizes and shapes. However, limitations include the requirement for specialist equipment and facilities and the safety hazards associated with the precursors and reactants, many of which are highly toxic. Quantum dots have found significant use as biological labels and this has led a number of research groups to seek improvements in their synthesis that obviate the need for hazardous reagents. For example, Green and co-workers have devised an improved protocol for the production of water-soluble cadmium telluride

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nanoparticles which uses ammonium telluride, (NH4)2Te, in place of H2Te. The CdTe nanoparticles, which were capped with cysteine or mercaptoundecanoic acid, are taken up by human breast cancer cells [14]. 5.2.4

Electrochemical Synthesis

Here the metal of interest is used as the sacrificial anode of an electrochemical cell [15]. As current is passed through the electrochemical cell, metal ions dissolve in the solvent, which contains a tetraalkylammonium salt, this having a dual role of stabilizing ligand and supporting electrolyte. The metal ions migrate to the cathode where nucleation and growth of the nanoparticles occurs as the metal ions are reduced to the zero-valent state. Electrochemical synthesis has been used to prepare particles of a host of transition metals including Fe, Co, Ni, and Pd [2]. A key advantage of this process is that the particles it yields tend to be purer than those obtained from chemical reduction methods, avoiding contamination with reducing agents and chemical by-products. 5.2.5

Other Routes for the Synthesis of Metal Nanoparticles

In addition to the more general approaches described above, a number of other protocols have been developed for the synthesis of functionalized nanoparticles. These have often been developed to produce materials for particular applications or with specific properties. For example, precipitation or seeding protocols are widely used to produce nanoparticles such as CdS quantum dots [13] and gold nanoparticles [1]. Size control is affected by varying the ratio of reagents. Laser ablation is a method that does not necessarily require the use of stabilizing ligands because the species produced using this method are charged and the Coulombic repulsions between particles prevents agglomeration [16, 17]. The reaction can be carried out in either aqueous or organic solvents with reasonable control of concentration, aggregation, and particle size. Irradiation of a gold plate suspended in water with a Nd:YAG laser yields gold nanoparticles with a diameter of about 18 nm [17]. These particles were coated with a thermoresponsive thiol-terminated poly-N-isopropylacrylamide-co-acrylamide co-polymer; above a transition temperature of 37 °C the polymer facilitates the reversible aggregation of the AuNPs. These polymer-functionalized particles are taken up by human breast adenocarcinoma MCF7 cells, with cells treated above 40 °C showing an 80-fold greater uptake compared to cells treated at 34 °C. This difference can be attributed to the aggregated particles having increased hydrophobicity [17]. Biological and biomimetic routes have used bacteria, plants, and plant extracts to synthesize noble metal nanoparticles [2]. Proteins such as ferritin, devoid of iron, have been utilized as templates and cages that can be filled with other nanomaterials, including cadmium sulfide, creating a quantum

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dot–ferritin nanocomposite [18]. Subsequently, this biomimetic approach has been applied to produce nanocomposites of other metals and proteins [19, 20]. An illustration of the level of complexity in the procedures that have been devised is provided by a recent report by Lin and co-workers, who have prepared fluorescent gold nanoparticles [21]. Their protocol started with the production of 5.5-nm surfactant-stabilized gold nanoparticles obtained using chemical reduction of AuCl3 in toluene. The particles were isolated and their size reduced by adding further quantities of HAuCl4 precursor, which etches the gold nanoparticles, producing smaller particles with a mean diameter of 3.7 nm. The particles were then transferred from toluene to aqueous media by exchanging the surfactant molecules with dihydrolipoic acid. Further purification by centrifugation and filtration yields stable AuNPs that are readily conjugated with biological molecules and taken up by cells.

5.3 SYNTHESIS OF FUNCTIONALIZED PARTICLES THROUGH LIGAND EXCHANGE The concept of ligand exchange was first described by Murray and co-workers [22]. The process is deceptively simple; mixing nanoparticles capped with a particular ligand with a second free ligand results in the replacement of a proportion of the original ligands (Figure 5.2). The method is particularly useful for preparing MMPCs and for introducing ligands onto the surface of the particle that may not be able to withstand the conditions employed during the original synthetic protocol. Biological molecules that possess a functional group that can bind to the surface of the particle, especially thiols, are readily attached to the surface of nanoparticles in this way. Examples include oligonucleotides, peptides, and biotin. An alternative strategy is to introduce thiol ligands bearing free carboxylate groups onto the surface of the nanoparticle. These can then be chemically coupled to biological molecules containing amino groups.

R R S S S

S S S Au

S SS

S S S

+R −

R S S S S

SH

S S

SH

Au S S S

R

S S S R

R

Figure 5.2. Schematic representation of ligand exchange.

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The apparent simplicity of ligand exchange belies the fact that exchange reactions involve a series of complex equilibria and kinetics that are difficult to study using conventional techniques. Moreover, most samples are not polydisperse and contain a range of particle sizes and hence surface coverage of ligands, with different ligand-binding sites on the nanoparticle surface displaying different levels of reactivity. A number of mechanisms are possible including associative or dissociative pathways and the formation of intermediate species. In addition to exchange between the functionalized particles and free ligand, interparticle ligand exchange may also occur. The latter possibility has been elegantly demonstrated by Murray, who investigated interparticle ligand exchange between two different functionalized nanoparticles that were dissolved in different, immiscible solvents [23]. For example, when an aqueous solution of tiopronin (N-(2-mercaptopropionyl)glycine)-coated gold particles was mixed with a toluene solution of alkanethiolate-coated gold particles, tiopronin-coated gold nanoparticles could be detected in the toluene phase and alkanethiolate-coated gold nanoparticles in the aqueous layer. Similar observations were made when aqueous solutions of tiopronin-capped silver nanoparticles were mixed with toluene solutions of alkanethiolate-capped gold nanoparticles. Interestingly, the ligand and metal exchange is inhibited when reactions are completed under N2, suggesting that oxidation reactions play a role in the transfer. This latter point illustrates the complexity of exchange reactions beautifully. At present, there is no standard procedure for determining the surface coverage of nanoparticles or the absolute number of ligands in MMPCs and a key conclusion of a recent review of mechanistic studies of ligand exchange is that for thiol-protected gold particles, at least, it is never possible to displace or exchange all of the ligands attached to the nanoparticle surface [24].

5.4

NANOPARTICLE–DNA INTERACTIONS

Conjugates of metal nanoparticles and DNA are of much current interest because of their potential biomedical applications and have been the subject of several recent reviews [25, 26]. DNA is usually attached to the surface of nanoparticles covalently. This can be done directly through the formation of bonds between functional groups, such as cysteine residues or thiols, present in the nucleotide. Alternatively, the DNA can be bound to functional groups that are appended to the capping ligands on the nanoparticle surface using conventional chemical coupling strategies. Mirkin and co-workers pioneered the synthesis of metal nanoparticle– DNA assemblies. In their initial work two batches of 13-nm gold nanoparticles were each functionalized with a noncomplementary thiol-derivatized oligonucleotide. When a third oligonucleotide that contains end sequences that are complementary to those that are grafted onto the surface of the nanoparticles is added, aggregation occurs forming the nanoparticle–DNA assembly. The

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assembly process is reversed by thermal denaturation [27]. Simultaneously, Alivisatos and co-workers reported a slightly different approach, using DNA to template two or three much smaller 1.4-nm gold nanoparticles, functionalized with oligonucleotides, into larger assemblies [28]. These seminal papers have catalyzed a phenomenal amount of research into nanoparticle–DNA assemblies with current emphasis focusing on their functionality [25, 26]. For example, it has recently been shown that DNA can be used to direct the assembly of gold nanoparticles into different micrometer-sized crystals, with either face-centered-cubic or body-centered-cubic lattices, dependent on the choice of DNA sequences attached to the surface of the gold nanoparticle and the DNA linkers [29]. An alternative approach to covalent linkage is the formation of bioconjugates through noncovalent interactions (Figure 5.3) [26, 30–32]; it is well known that noncovalent interactions play crucial structure-directing roles in biological molecules. The attachment of capping ligands bearing charged

–

O O P O O

–

Au

S

O –

O P O O

O

–

O

NH2

N

H2N

Base

Base

O

+

+

SH

N

SH 11

(B)

(A)

O +

–

S SO3

P

(C)

+

S

P

CH3 Br

–

(D)

Figure 5.3. Schematic representation of the interaction of cationically functionalized gold nanoparticles with oligonucleotides and the molecular structures of some cationic capping ligand precursors: (A) ethidium thiol, (B) trimethylammonium alkylthiol, (C) phosphonioalkylthiosulfate zwitterions, and (D) ω-thioacetylalkylphosphonium salts.

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groups to the surface of nanoparticles is an attractive proposition for improving the aqueous solubility of the resulting MPCs. Noncovalent interactions between DNA and functionalized nanoparticles can be facilitated through intercalation, groove binding, or simple electrostatic attractions. Ethidium bromide is a well-known DNA intercalator and this species has been introduced onto the surface of cationic and anionic gold nanoparticles as a means of inducing binding to DNA. The ethidium groups are grafted onto the gold nanoparticles as thiolate ligands through a ligand exchange reaction; each particle contains only a few ethidium thiolate groups. The remaining capping ligands are TMA (trimethyl(mercaptoundecyl)ammonium) for the cationic species, or tiopronin (N-(2-mercaptopropionyl)glycine) for the anionic particles. In the absence of DNA, the normal fluorescence of the ethidium groups is quenched by the gold core. Introduction of DNA results in a partial restoration of the fluorescence. As one might anticipate, the cationic TMAfunctionalized gold nanoparticles bind rapidly and efficiently with the DNA, whereas the binding of the anionic tiopronin-functionalized is slow compared to ethidium that is not bound to the gold [30]. Rotello and co-workers have advanced the use of nanoparticles functionalized with alkylthiolate ligands bearing cationic ammonium head groups, which provide a complementary surface for binding DNA through electrostatic interactions with the negatively charged phosphate groups in the DNA backbone [26, 31, 32]. For example, the nanoparticles have been used to recognize a 37-mer DNA duplex inhibiting transcription by T7 RNA polymerase. This result illustrated the high affinity of the DNA for the functionalized nanoparticle and highlights their potential use in gene regulation [32]. Our own contribution to this area has been the synthesis of a series of phosphonium-capped gold nanoparticles (Figure 5.4) [33, 34]. The most widely

Ph

Ph

Ph Ph Ph + P Ph

Ph P+

Ph Ph Ph P+

Ph + Ph P

Ph + Ph P Ph

–

3S

SO3 – 4

or

+ AuCI O

Ph + Ph P Ph

–

3S

Ph

CH3 Br

NaBH4 H2O/CH2CI2

Ph

P

Ph Ph + P Ph

s

s s s s

s s

Ph + Ph P Ph

+ Ph

s

Ph Ph

Ph Ph

s

s

P

+

P

P +P Ph Ph Ph

Ph Ph

+ Ph

s

+

Ph

P

s

s

Ph + P

+ Ph

s

Au

+

P Ph Ph Ph

Ph

Ph Ph

Ph Ph

Figure 5.4. Schematic representation of protocol for the synthesis of phosphoniumcapped gold nanoparticles.

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studied cationic capping ligands feature ammonium head groups as the chargecarrying species [26, 31]. Nonetheless, phosphonium groups offer a number of advantages including the availability of a wide range of organic substituents, which allows the possibility of creating a range of additional functionality, such as fluorescence or chirality and, most importantly, biocompatability; the ability of the triphenylphosphonium group to travel across cell membranes is well established. The latter property has led to the use of phosphonium compounds as transport vectors for targeting mitochondria [35, 36], anticancer agents [37], and as diagnostic agents [38]. Although there are many studies of the use of organophosphorus ligands, notably phosphines and phosphine oxides [39–41], for passivating the surface of nanoparticles, there have been far fewer studies of the use of phosphorus ligands to impart functionality to nanoparticles. Tris(hydroxymethyl) phosphine (THP)-capped gold nanoparticles, which have an average diameter of 1.5 nm, have been formed through the reduction of HAuCl4 with tetrakis (hydroxymethyl)phosphonium chloride. The resulting THP-gold nanoparticles, although negatively charged, readily form conjugates with DNA. It is believed that the interaction with DNA is facilitated by hydrogen bonds between the DNA bases and the hydroxy groups of the THP [41]. Biocompatible gold nanoparticles, stabilized by phosphorylcholine thiolate ligands, have been prepared and show enhanced stability compared to gold nanocrystals stabilized with polyethylene glycol [42]. Phosphonium-functionalized gold nanoparticles are synthesized using a variation of the Brust method. The phosphonium ligands are derived from either phosphonioalkylthiosulfate zwitterions or ωthioacetylalkylphosphonium salts, which behave as masked thiolate ligands [33, 34]. The term “masked thiolate” is used to describe alkylthiolate species that are “protected” as another group, in this case either thiosulfate or thioacetate, which, under reductive conditions or contact with metal surfaces, undergo cleavage of the S–SO3 or S–C(O)CH3 bonds, generating thiolate anions that attach to the surface of the growing nanoparticle. Masked thiolates are easier to handle and less prone to oxidation than conventional thiol ligands. The ability of the phosphonium gold nanoparticles to interact with RNA and DNA and has been investigated using a Biacore instrument that relies on changes in the surface plasmon resonance of sensor chips when exposed to analytes to study biomolecular interactions [43]. The phosphoniumfunctionalized gold nanoparticles interact strongly with negatively charged carboxylate groups and streptavidin, which are attached to the Biacore sensor chip. Neutral, citrate-stabilized, gold nanoparticles do not interact in the same way and are easily released by the sensor chip. Once bound to the sensor chip, the phosphonium-capped gold nanoparticles will bind other biomolecules such as DNA and RNA and this is currently being investigated for biomolecular recognition.The cellular uptake of phosphonioalkylthiosulfate zwitterions, ω-thioacetylalkylphosphonium salts, and phosphonium-functionalized gold

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nanopartciles has also been investigated. The results indicate that the ligands and the functionalized gold nanoparticles are accumulated by the cells. The presence of the capping ligands and the gold within the cells was confirmed using MALDI-MS, which provides a relatively straightforward means of detecting these species in cells. The results confirm that phosphonium ligands offer a convenient way of transporting gold nanoparticles into cells.

5.5 FUNCTIONALIZED NANOPARTICLES FOR BIOLOGICAL DETECTION The use of nanoparticles in biological imaging and detection is one of the most widely explored applications of metal nanoparticles [14, 21, 26, 44–46]. Traditional methods for the labeling and detection of biological molecules utilize fluorescent chemical dyes or radiolabels. Both of these approaches have their limitations, including broad emission spectra of dyes, which makes use of multiple dyes difficult. Moreover, many organic dyes suffer from photobleaching, resulting in the deterioration of the emission signals after prolonged exposure to the excitation light. The challenges associated with radiolabels include the requirement for specialist facilities for handling and disposal of radioactive species. Many of these disadvantages are overcome by quantum dots, which have been widely explored as agents for biomedical imaging [43, 44]. Semiconductor quantum dots display tunable emission spectra, dependent on the size, shape, and functionalization of the particles. Furthermore, the emission spectra are narrow, which enables nanoparticles to be utilized in multiplexed assays. Quantum dots also show greater stability to photobleaching and a number of materials are now available commercially as reagents for biological imaging. One of the challenges associated with the synthesis of quantum dots for imaging applications is transfer from organic solvents to aqueous systems, although this can be overcome by growing a silica shell around the semiconductor nanoparticle core to which biocompatible ligands can be attached, or by swapping the capping ligands used in the synthetic procedure with water-soluble species such as mercaptoacetic acid by ligand exchange [44, 45]. A potential downside of nanoparticulate imaging agents is their production of cytotoxic side products upon degradation, an issue that needs to be considered when conducting biological imaging experiments [14]. The surface plasmon resonance of gold nanoparticles, which occurs when the particles absorb and emit light, has been exploited in biological contrast agents [46]. The nanoparticles are functionalized with antibodies that bind with antigens in the cells or regions of interest. In addition to visualization by light, gold nanoparticles can also be imaged by X-ray methods or by microscopic techniques such as TEM and AFM. Gold particles smaller than 2 nm do not display plasmon resonance but do exhibit size-dependent fluorescence.

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Recent studies have also shown that this fluorescence can be used for biological labeling, offering an alternative to quantum dots [21].

5.6

METAL NANOPARTICLES AS THERAPEUTIC AGENTS

Funtionalized metal nanoparticles are readily taken up by cells, which makes them attractive carrier systems for transporting drugs, biomolecules, or other therapeutic agents into cells [26, 46–51]. Furthermore, the nanoparticles themselves can be used as the therapeutic agents, as, for example, in the use of gold nanoparticles as an intercellular heat source [46]. Trapping the drug molecule between capping ligands attached to the nanoparticle surface is a method that has recently been reported [48]. The drug molecules are held in place through noncovalent intermolecular interactions and are released by diffusion. One of the primary aims of nanoparticle-based drug delivery systems is the targeted delivery of the therapeutic agent at the site of the disease. This can be achieved by synthesizing nanoparticles bearing ligands that are only recognized by specific receptors that are present on the cell surface. A significant amount of attention has focused on the use of nanomaterials in the treatment of cancer and gold nanoparticles have been synthesized that carry folic acid attached to their surface; folate receptors are overexpressed on the surfaces of many cancer cells [49]. Similarly, protein tumor necrosis factor TNFα has been grafted onto the surface of PEG-stabilized gold nanoparticles, which have been shown to be preferentially accumulated by tumors [50]. Methotrexate-functionalized magnetic Fe3O4 nanoparticles have been synthesized (Figure 5.5), the methotrexate interacting with the cellular folate receptors. These materials offer the dual function of diagnostics, acting as a contrast agent for MRI, and drug delivery. The methotrexate was covalently bound to (3-aminopropyl)-trimethoxysilane assembled on the particle surface through amidation between the carboxylic acid end groups on methotrexate and the pendant amine groups on the capping ligand. The methotrexate was shown to be cleaved from the nanoparticles inside cells [51]. An alternative approach to targeting nanoparticles toward specific cellular sites is to use so-called gene guns, where nanoparticle–DNA conjugates are forced inside cells using ballistic acceleration [46]. The use of metal nanoparticles themselves as therapeutic agents stems from their ability to absorb energy, usually light or radiofrequency fields, which causes them to become excited. The excitation energy is eventually dissipated into the surrounding tissues as heat. Since cells are very sensitive to even small variations in temperatures, this can lead to cell death. This approach has been used to destroy cancerous tissues that have been infused with gold nanoparticles. The tissue is illuminated with light that is absorbed by the gold nanoparticles, resulting in cell death. A limitation of this use of gold nanoparticles is that visible and IR light can only penetrate surface tissue. Therefore magnetic

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Fe2O3

MeO OH + MeO Si MeO

NH2

PhMe,sonication

OMe Fe2O3

O Si

NH2

OMe

(i) Methotrexate,DMSO (ii) 1-ethy1-3- (3-(dimethylamino) propylcarbodiimide (EDC), N-hydroxysuccinimide (NHS). O OMe Fe2O3

O Si OMe

N H

NH2 O

HO

N H

N

N

N

O

N

N

NH2

Figure 5.5. Drug molecules can be attached to the surface of nanoparticles using established methodologies, as exemplified by the conjugation of methotrexate with magnetic ferric oxide nanoparticles.

nanoparticles have been developed that are excited by radiofrequency radiation, which are able to reach deep inside the body [46].

5.7

TOXICOLOGY OF METAL NANOPARTICLES

As we have seen, metal nanoparticles offer huge potential benefits to medicine through their applications as diagnostic and therapeutic agents. However, a much greater understanding of the possible risks to human health is also required before nanoparticles are used routinely, and yet this remains a comparatively underexplored aspect of nanomaterials science. A number of recent papers have reviewed aspects of nanoparticle toxicology and cytotoxicity [52– 55]. The paucity of toxicological studies stems, in part, from the fact that nanomaterials are not a uniform group but show great diversity, not only in chemical composition but also in morphology and functionalization. Indeed, the dissociation and subsequent metabolism of capping ligands and other functional molecules appended to the surface of the nanoparticles adds a further level of complexity.

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The health risks of nanoparticles stem from their small size and high surface area and their ability to penetrate tissues and pass across cellular membranes. Studies show that adverse affects are related to the dosage and concentration and the length of exposure. Much attention has focused on quantum dots due to the risk or releasing of heavy metal ions with known toxicity as the particles are metabolized [55, 56]. A pharmacokinetic study of commercially available QD705 quantum dots, which contain a CdSeTe core surrounded by a ZnS shell and a methoxy-PEG coating, in male ICR mice has been reported. The study reveals that the quantum dots are accumulated in the kidneys, liver, and spleen for at least 28 days but are gradually and partially eliminated by 6 months [56]. Much of the interest in gold nanoparticles results from their stability and perceived biocompatibility. Of particular interest is a report by Connor and co-workers who studied the effects of particle size and capping ligand on the cytotoxicity of human leukemia cells [57]. The gold particle size ranged from 4 to 18 nm. The capping ligands included citrate, glucose, cysteine, CTAB, and biotin. The results indicated that the 18-nm particles functionalized with citrate or biotin were not toxic at concentrations up to 250 μM. In contrast, similar concentration solutions of gold(III) chloride salt are very cytotoxic. Gold particles capped with glucose and cysteine displayed greater cytotoxicty. A comparative study of the toxicology of 2-nm gold nanoparticles functionalized with either cationic (ammonium) or anionic (carboxylate) thiolate ligands toward mammalian and bacterial cell lines has been undertaken [58]. The research revealed that both types of cells show similar levels of uptake and the cationic particles are moderately toxic, whereas the anionic particles are nontoxic, demonstrating the relationship between cytotoxicity and surface functionalization. The studies undertaken to date provide a base for further, more detailed, studies of the pharmacological and toxicological effects of metal nanoparticles, which are almost certainly required before these materials are routinely used in human medical applications.

5.8

CONCLUSIONS AND FUTURE PROSPECTS

Metal nanoparticles offer huge potential in a wide range of biomedical applications. As the field of research has emerged and matured over the past 20 years, the emphasis of much of the research into the synthesis of metal nanoparticles has moved from developing new methodologies to refining and improving existing procedures to make materials for custom applications or with more closely defined morphological parameters. Despite the numerous advances that have been made in recent times, challenges and opportunities remain. The routine production of monodisperse particles with precisely known surface morphology and ligand coverage has still to be achieved. Although we know that metal nanoparticles can be taken up by cells, the

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general mechanistic aspects remain poorly understood. Furthermore, there is a lack of detailed knowledge of the fate of the metal particles once inside cells, which links with the comparatively limited amount of detailed pharmacokinetic and toxicological data on metal nanoparticles. However, it is almost certain that these challenges will be overcome in the near future and that metal nanoparticles will be routinely used for the detection and treatment of disease.

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35. Weissig, V., Boddapati, S. V., Jabr, L., and D’Souza, G. G. Mitochondria-specific nanotechnology. Nanomedicine 2(3): 275–285 (2007). 36. Murphy, M. P. and Smith, R. J. A. Targeting antioxidants to mitochondria by conjugation to lipophilic cations. Annu. Rev. Pharmacol. Toxicol. 47: 629–656 (2007). 37. Rideout, D. C. et al. Phosphonium salts exhibiting selective anti-carcinoma activity in vitro. Anti-Cancer Drug Design 4(4): 265–280 (1989). 38. Yang, C.-T., et al. Synthesis and structural characterization of complexes of a DO3A-conjugated triphenylphosphonium cation with diagnostically important metal ions. Inorg. Chem. 46(21): 8988–8997 (2007). 39. Binder, W. H., Sachsenhofer, R., Straif, C. J., and Zirbs, R. Surface-modified nanoparticles via thermal and Cu(I)-mediated “click” chemistry: generation of luminescent CdSe nanoparticles with polar ligands guiding supramolecular recognition. J. Materials Chem. 17: 2125–2132 (2007). 40. Duff, D. G., Baiker, A., and Edwards, P. P. A new hydrosol of gold clusters. 1. Formation and particle size variation. Langmuir 9(9): 2301–2309 (1993). 41. Harnack, O., Ford, W. E., Yasuda, A., and Wessels, J. M. Tris(hydroxymethyl) phosphine-capped gold nanoparticles templated by DNA as nanowire precursors. Nano Lett. 2(9): 919–923 (2002). 42. Qiao, J., Xu, J.-P., Ji, J., and Shen, J.-C. Zwitterionic phosphorylcholine as a better ligand for stabilizing large biocompatible gold nanoparticles. Chem. Commun. 3058–3060 (2008). 43. Ju-Nam, Y., Functionalised Metal Nanoparticles as Novel Reagents for Biomedical Analysis, Ph.D. thesis, Sheffield Hallam University, UK, 2007. 44. Green, M. Semiconductor quantum dots as biological imaging agents. Angew. Chem. Int. Ed. 43: 4129–4131 (2004). 45. Smith, A. M., Duan, H., Mohs, A. M., and Nie, S. Bioconjugated quantum dots for in vivo molecular and cellular imaging. Adv. Drug Deliv. Rev. 60: 1226–1240 (2008). 46. Sperling, R. A., et al. Biological applications of gold nanoparticles. Chem. Soc. Rev. 37: 1896–1908 (2008). 47. Han, G., Ghosh, P., and Rotello, V. M. Functionalized gold nanoparticles for drug delivery. Nanomedicine 2(1): 113–123 (2007). 48. Kim, C. K., et al. Entrapment of hydrophobic drugs in nanoparticle monolayers with efficient release into cancer cells. J. Am. Chem. Soc. 131(4): 1360–1361 (2009). 49. Dixit, V., et al. Synthesis and grafting of thioctic acid-PEG-folate conjugates onto Au nanoparticles for selective targeting of folate receptor-positive tumor cells. Bioconjug. Chem. 17: 603–609 (2006). 50. Kholer, N., Sun, C., Wang, J., and Zhang, M. Q. Methotrexate-modified superparamagnetic nanoparticles and their intracellular uptake into human cancer cells. Langmuir 21: 8858–8864 (2005). 51. Paciotti, G. F., et al. Colloidal gold: a novel nanoparticle vector for tumor directed drug delivery. Drug Deliv. 11: 169–183 (2004). 52. Oberdorster, G., Stone, V., and Donaldson, K. Toxicology of nanoparticles: a historical perspective. Nanotoxicology 1(1): 2–25 (2007). 53. Yang, W., Peters, J. I., and Williams, R. O. III. Inhaled nanoparticles—a current review. Int. J. Pharmaceutics 356: 239–247 (2008).

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54. Murphy, C. J., et al. Gold nanoparticles in biology: beyond toxicity to cellular imaging. Acc. Chem. Res. 41(12): 1721–1730 (2008). 55. Lewinski, N., Colvin, V., and Drezek, R. Cytotoxicity of nanoparticles. Small 4(1): 26–49 (2008). 56. Lin, P., et al. Computational and ultrastructural toxicology of a nanoparticle, quantum dot 705, in mice. Environ. Sci. Technol. 42(16): 6264–6270 (2008). 57. Connor, E. E., et al. Gold nanoparticles are taken up by human cells but do not cause acute cytotoxicity. Small 1: 325–327 (2005). 58. Goodman, C. M., McCusker, C. D., Yilmaz, T., and Rotello, V. M. Toxicity of gold nanoparticles functionalized with cationic and anionic side chains. Bioconjug. Chem. 15: 897–900 (2004).

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CHAPTER 6

Subcellular Fate of Nanodelivery Systems DUSICA MAYSINGER and SEBASTIEN BORIDY Department of Pharmacology & Therapeutics, McGill University, Montreal, Quebec, Canada

ELIZA HUTTER Department of Pharmacology and Therapeutics, McGill University, Montreal, Quebec, Canada

6.1 INTRODUCTION Today, nanosciences are experiencing massive investment worldwide. Possible consequences of nanoparticle (NP) entry into biological systems, animals, plants, and humans are not known and a quest to produce safe NPs useful for the electronic industry, food production, artificial paints, and biomedical applications is actively ongoing. The production of manufactured NPs underscores the question: Does long-term cumulative exposure to NPs have deleterious effects on our health? Small size NPs in low nanomolar or picomolar concentrations could reach different tissues including the brain but they cannot be easily detected. Since the induction of intracellular oxidative stress appears to be an essential component of the biological side effects of many metallic and carbon NP types [1–3], a lifetime’s accumulation of NPs in minute quantities could seriously accelerate the onset of inflammatory and degenerative processes in the brain. Therefore, studies to provide information on NP entry into cells and how to reduce their untoward effects at cellular and system levels have been actively pursued in different laboratories. On the other hand, polymeric NPs, which could stimulate repair mechanisms by releasing trophic factors, antioxidants, or chelating agents incorporated into, for example, block copolymer micelles [4–10], are promising candidate NPs for therapeutic interventions. Indeed, several of them are already at different stages of clinical trials. Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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In this chapter we will discuss (1) how the properties of three selected classes of NPs determine their uptake and distribution at the single cell level and in specific organelles, (2) the mechanisms involved in internalization of these NPs, and finally, (3) the possible untoward effects that some (metallic) NPs produce and how to provide the means of preventing and/or protecting cells/tissues from possible adverse effects. Our research is primarily focused on mimicking and activating intrinsic cellular protective mechanisms (e.g., by mimicking and enhancing the capacity of endogenous chaperones, induction of antioxidant systems, and stimulation of endogenous protectins and resolvins) [11–13]. We will focus here on three groups of NPs that are of particular interest in nanomedicine. These NPs are (1) quantum dots, (2) gold NPs, and (3) polymeric artificial chaperones.

6.2

QUANTUM DOTS

6.2.1. Properties of Quantum Dots Quantum dots (QDs) are luminescent NPs with unique optical properties that have been exploited for single cell and whole animal imaging. Excellent reviews summarizing the current status of QDs are available [14–16]. Some of the physicochemical properties of QDs, particularly those used for assessment of intracellular trafficking of therapeutically active molecules, are presented in Chapter 27 of this book. When coated with proteins or biocompatible polymers, QDs are generally not deleterious to cells and organisms. However, when QDs are retained in cells or if they accumulate in the body over a long period of time, their coatings may be degraded, yielding “naked” QDs. We were particularly interested in the effects of “naked” QDs. Our studies showed that they can induce damage to the plasma membrane, mitochondrion, and nucleus, ultimately leading to cell death [17]. Reactive oxygen species (ROSs) are important players in mediating QD-induced cellular damage, for example, with cadmium telluride QDs. QD-induced cytotoxicity can be reduced or even eliminated without covalent binding of protective agents to the QD surface. Among the surfaces that are the most effective are multilayer ZnS capping and polymers such as polyethylene glycol (PEG). Of course, the size and charge of the PEG chain is critical for QD internalization, whole body distribution, and subcellular distribution. The following sections are meant to illustrate these points. 6.2.2 Biodistribution in the Whole Body and Subcellular Distribution of QDs Quantum dots are attractive fluorescent probes extensively investigated in the emerging field of nanomedicine [18]. Recent studies by Choi et al. [19] show

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how size and charge of QDs affect their renal clearance. Selected types of QDs were injected intravenously in rodents and the key results showed that zwitterionic or neutral organic coatings prevented the adsorption of serum proteins and did not markedly impair renal excretion. These studies also show that a final hydrodynamic diameter <5.5 nm provides the appropriate size for urinary excretion and elimination of QDs from the body. Studies related to QDs in whole animals are sparse. One of the reasons for this is that imaging of the whole animal using fluorescent NPs, in general, remains difficult, because of strong autofluorescence of tissues due to endogenous chromophores (e.g., collagens, porphyrins, and flavins), in addition to the stability of the core and corona ligands. In most studies using CdSe/ZnS, the issue of core stability within the time period tested did not seem to pose a problem. However, noncovalently attached surface molecules might have been partly lost en route toward the predicted destination. It is also unknown to which degree QDs are quenched due to interactions with extracellular and cellular proteins. A newer class of QDs with superior photophysical properties, which can at least in part overcome these limitations by emitting in nearinfrared (NIR) regions, were prepared and tested in rodents [20]. Using NIR-emitting QDs avoids interfering noise from the animal autofluorescence. An example of such QDs is QD705 (20–25 nm), which were injected intravenously in mice. Results from these studies show that after 24 hours, they primarily accumulate in the liver and spleen. Analyses of urine and feces did not reveal significant levels, suggesting that most of the QD705 had remained in the body. Concordant with such a notion are results obtained by Fischer et al. [21]. It seems that surface-modified QDs with PEG are biocompatible with both living cells in culture [22] and whole animals. The physical, chemical, and biological properties, in addition to the availability, make PEG an attractive corona-forming candidate for micelles and QDs [23–26]. Reports on distribution and their pharmacokinetic properties [27–29] show that CdSe QDs are sequestered in several organs following intravenous administration, including lymph nodes [30] and solid tumors [23]. Studies by Inoue et al. [31] showed that QD injection allowed for long-term and repeated observation of the reticuloendothelial system (RES) in mice despite gradual decline of signal intensity. Combined fluorescence and luminescence imaging in luciferase-expressing tumor cells provided a way of concomitant detection of tumor cells and QDs. 6.2.3

Nanoparticle Distribution in the Brain

QD distribution in the brain in real time was not extensively explored. We studied glial cells in the nervous system because astrocytes and microglia are most likely the major players in delivering NPs to the central nervous system (CNS) [32, 33]. Importantly, they regulate neuronal signaling and viability; without their biochemical, morphological, and physiological communication, higher mental or emotional functions at the level of the whole

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body would not be achievable. An example of imaging fluorescent CdSe/ZnS and InGaP/ZnS NPs in cells in culture and in vivo in transgenic mice showed functional responses of glia to NPs [34]. Such studies are particularly useful for the assessment of nonfluorescent and degenerate NPs, such as cerium oxide and degenerate “naked” CdTe NPs, which could not be easily detected by common techniques but can still interfere with CNS functions. Similar investigations could provide information on biocompatibility of QDs in live animals, in different glial cell types, especially after repeated imaging sessions. Studies by Maysinger et al. [35] focused on astrocyte response to different types of QDs. Astrocytes are the principal macroglial cell type in the brain and their activation is one of the key components of the cellular responses to stress and brain injuries. The passage from the quiescent to reactive astrocytes is associated with strong upregulation of the intermediate filament, glial fibrillary acidic protein (GFAP) [36, 37]. GFAP upregulation is considered a surrogate marker of neuronal stress and brain inflammatory response. Current methods of astrocyte and microglia detection are mainly based on immunocytochemistry. Using a transgenic mouse that carries the luciferase gene under the transcriptional control of murine GFAP promoter [38], upregulation of GFAP or luciferase expression in response to QDs was analyzed noninvasively in living animals using biophotonic imaging and a high-resolution CCD camera [39]. The principle of generating luciferase-expressing mice and imaging assays in real time are illustrated in Figure 6.1. 6.2.4

NP Distribution and Entry into Cells

Imaging of living cells provided a wealth of knowledge leading seminal findings regarding several types of QDs. These studies answered in part some of the key biological questions related to QD uptake [40], subcellular distribution, and functional consequences. Studies focused primarily on identifying what factors were primarily involved in determining the extent of uptake. Was it the size, charge, or other surface properties that determine the extent of the uptake? Are QDs taken up and retained or eliminated from the cells or do they accumulate intracellularly and ultimately get degraded? Despite the convincing data, these results are limited to particular cell types and particular QDs; much more studies are required to have information for other types of cells and QDs. Nonfunctionalized nanocrystals can exploit cells’ active transport machinery and be delivered into the cells and even the nucleus [41]. These studies show that living human macrophages are able to rapidly internalize and accumulate QDs in distinct cellular compartments, the extent of which depends on the QD’s size and charge. The most remarkable finding is that the smallest CdTe QDs specifically target histones in cell nuclei by a multistep process. The nuclear entry seems to be mediated via the nuclear pore complexes. These studies concur with previous findings by Lovric et al. [42] showing, by confocal

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TLR2 PROMOTER

Luciferase Reporter

GFAP PROMOTER

Luciferase Reporter

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cDNA

Inject cDNA into fertilised oocyte

Plant oocyte into a pseudomother and breed transgenic pups

+ Stress + Luciferin

= Bioluminescent signal

Figure 6.1. Transgenic reporter mice with microglial and astroglial activation markers. Luciferase promoter constructs for the glial fibrillary acidic protein (GFAP; astroglial activation marker) and toll-like receptor (TLR2; microglial activation marker) are injected into donor oocytes, which are transplanted into pseudomothers, used to breed transgenic reporter mice. Prior to imaging, mice receive intraperitoneal injections of d-luciferin, the substrate for luciferase. Given that luciferase expression is regulated by stressful stimuli specific to these glial cell types, the resultant conversion of the substrate to oxyluciferin, the biolumiscent product, is proportional to the degree of stress.

microscopy, that this could be the case. More recent studies [43] provide further evidence that minute amounts of small (<5 nm in diameter) QDs can interfere with histones and induce epigenetic changes. Studies in this direction are warranted to uncover possible histone modifications with or without subsequent gene activations. This is an exciting and emerging new branch in nanotoxicology, which we propose to call “nanoepigenetics” [43]. An overview of modes of NP entry is provided in Figure 6.2. The concept of combining bioluminescence and fluorescence in living cells was first illustrated by So et al. [28]. These studies showed self-illuminating QDs tested in living cells and whole animals. The results from these studies provide evidence for the usefulness of QDs covalently bound with luciferase. When the conjugates were exposed to luciferase substrate, coelentrazine, the

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ClathrinMediated

MacroPinocytosis

Amiloride Wortmannin Ly294002

Chlorpromazine Ikarugamycin Phagocytosis EEA-1

Nucleus

EEA-1

Bafilomycin A1 Chloroquine

CaveoleaMediated Clathrin Dynamin

GNPs QDs

Caveolin

CHPs

Methyl-βCyclodextrin Lovastatin

Figure 6.2. Mechanisms of nanoparticle uptake by cells. The four primary modes of NP uptake are illustrated. Clathrin-mediated endocytosis, caveolae-mediated endocytosis, macropinocytosis and phagocytosis. Early endosomes are indicated by the presence of the early endosome antigen 1 (EEA-1), and acidification to the late endosome is indicated by red-yellow gradient, representing a drop in pH. Specifically, it has been shown that via receptor-mediated endocytosis, gold nanoparticles (GNPs), quantum dots (QDs), and nanogels (CHPs) are internalized within clathrin-coated vesicles. Evidence supporting caveolae-dependent uptake has also been demonstrated for GNPs and QDs. Secondary, nonspecific mechanisms are also illustrated, namely, macropinocytosis and phagocytosis. In addition, drugs that selectively inhibit these mechanisms are incorporated in the schematic, some of which have been used to investigate the uptake of NPs discussed in this chapter.

energy released by substrate catabolism was transferred to QDs by bioluminescence energy transfer leading to QD fluorescence. The protocol for making these self-illuminating QDs, provided by So, is widely applicable to other conjugation products, which can be utilized for different biosensing purposes. 6.2.5

Current Status and Future Perspective

Stabilized QDs with high fluorescent yield and specific ligands on their surfaces may eventually become common nanomaterials for the development of highly accurate and sensitive tests for clinical diagnosis. Such tests could

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overcome the limitations of organic fluorophores (e.g., bleaching and wide spectral emission bands). In the meantime, QDs will continue to be very useful tools to explore the mechanisms of cell–NP interactions in experimental animals and living cells in cultures.

6.3 6.3.1

GOLD NANOPARTICLES General

Gold nanoparticles (GNPs) have attracted special attention among the known nanomaterials in the field. The localized surface plasmons (charge density oscillations) provide GNPs with unique optical properties, of which various biological applications (i.e., imaging, sensing, drug and gene delivery, and photothermal therapy) can take advantage. The excitation of localized surface plasmons that occurs for a specific frequency of light results in strong light scattering (an 80-nm diameter metallic NP is estimated to have a scattering flux that corresponds to those of 106 fluorescein molecules [44]), intense absorption, and enhancement of electromagnetic fields. In biological applications, these effects make GNPs particularly useful as labeling agents, signal amplifiers, sensors, or therapeutic agents. Indeed, metallic NPs were shown to be able to replace (or complement) established radioactive, fluorescent, chemiluminescent, or colorimetric labels, used in biochemistry, cell biology, and medical diagnostic applications [45–47]. The ultrabright, nonbleaching NPs can be observed individually and can be prepared with a scattering peak at any color of the visible or near-IR spectrum (by changing their size, shape, or surface), thus enabling the detection of single molecules and/or multiple targets. Rod-shaped GNPs of different lengths, for example, were successfully employed for multiplex identification of cell surface markers [48, 49]. Another advantage of the tunability of the surface plasmon excitation is that it opens the way to work in a desirable spectral range. Gold nanorods, nanoshells, and nanourchins are particularly attractive tools for in vivo applications, since their optical resonance lies in the near-infrared spectral window, away from the region of biomolecular excitation transitions, thus precluding photochemical damage and ensuring deeper penetration of the light into live systems [50–55]. A number of publications demonstrated the feasibility of in vivo imaging using GNPs. For example, gold nanoshells were imaged by photoacoustic tomography in rat brain [56], and PEG-coated spherical GNPs were used to show a clear delineation of cardiac ventricles and great vessels by computer tomography [57]. A twofold contrast was found between hepatoma and normal liver tissue by the same method [57]. In surface-enhanced Raman spectroscopy, large optical enhancement was achieved for the detection of tumors under in vivo conditions using small GNPs [58]. Gold nanorods have been imaged in vivo by two-photon luminescence (TPL) microscopy in epithelial cancer cells [59, 60] and in mouse ear blood vessels [61]. Our group

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explored the fate of gold nanorods in living primary cortical neurons by TPL. We found that gold nanorods with CTAB, PEG, and insulin on their surfaces were internalized by living cortical neurons and could be imaged with excellent signal-to-noise ratio by two-photon luminescence microscopy, as shown in Figure 6.3. These examples of in vivo applications suggest great potential for accurate GNP-based assessment of acute ischemic stroke, lesion development, thermal injury, neovascularization, tumor angiogenesis, tumor necrosis, hepatic function, and regional hemodynamic activities. In addition to imaging, targeted GNPs with near-IR absorbance are excellent candidates for photothermal therapy (the plasmon resonant light-induced heat release is lethal to cancer cells in the proximity of GNPs). Cancer cells, labeled by tumor-specific antibody conjugated gold nanorods or nanoshells, can be selectively destroyed by near-IR laser irradiation [62–65]. Intravenously administered small GNPs were also shown to enhance the effect of radiation therapy in tumor-bearing mice [66, 67]. 6.3.2

Biocompatibility and Biodistribution of GNPs

While the favorable optical and photophysical parameters strongly promote GNPs for in vivo applications, relatively little is known about the interaction of GNPs with living cells and organisms. GNPs were considered inert for many years. However, new findings indicate that even these “noble” NPs are not so noble in dealing with living cells and they can also induce some of undesirable effects. Fortunately, GNP sizes and shapes can be stringently controlled and untoward effects can be reduced by precisely tailoring the size, charge, and ligand on the GNP surface. The longstanding use of colloidal gold in the treatment of rheumatoid arthritis and the use of radioactive GNPs to treat liver cancer [68, 69] suggests that colloidal GNPs are relatively inert and biologically compatible carriers. However, with recent developments in GNP synthesis and application systems, extensive biocompatibility studies are an absolute necessity. Besides the increasing variety of surface modification, a great number of novel shapes have been prepared during the last decade. Gold nanorods [53, 70], nanourchins [50, 54, 55], nanoprisms [71], nanowires [72], nanodisks [73], and nanostars [74, 75] were synthesized with high yield and reproducibility, awaiting their testing in biological systems. The internalization and biocompatibility of GNPs have been assessed in a number of studies employing mostly spherical GNPs and, occasionally, gold nanorods. Most studies revealed that exposure of cell lines or living organisms to spherical GNPs causes no acute cytotoxicity [76–78]. Colloidal GNPs need to be stabilized against aggregation by various ligands, which may be inherently toxic or induce GNP toxicity. For example, Rotello and co-workers reported that small cationic GNPs (2 nm) are more toxic toward red blood cells, Cos-1 cells, and Escherichia coli bacteria than anionic particles [79].

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Figure 6.3. Two-photon luminescence (TPL) imaging of primary cortical neurons. TPL imaging of primary cortical neurons treated 21 days in vitro (DiV 21) with gold nanorods (rod GNP) for 4 hours prior to imaging (CTAB—cetyl-trimethylammonium bromide). Neurons were also stained with mitotracker green FM. Scale bar in the control panel is representative for both panels. (See color insert.)

Chan’s team work indicates that the size of the GNPs can also modulate their toxicity, small spherical GNPs being more toxic than their larger counterparts [80]. In addition, cell-specific toxicity has also been reported: the same 33-nm citrate-capped gold nanospheres that were found to be cytotoxic, at certain concentrations, to a human carcinoma lung cell line, proved to be innocuous toward baby hamster kidney and human hepatocellular liver carcinoma cells [81]. We tested different morphologies (spherical, rod-shaped, and urchin-like) for GNPs in primary neurons (Figure 6.3) in neuroblastoma cells (SH-SY5Y), and in microglial cultures, considered to be the macrophages of the nervous system (Figure 6.4). Results from these studies show that within the concentration range examined (up to 57.9 μM) GNPs do not harm neurons or glia, regardless of their shape and surface chemistry [81a]. Upon administration in animals, GNPs were found to accumulate in the liver and the spleen, but without causing apparent cytotoxic effects [66]. In a study evaluating the influence of particle size on the biodistribution of GNPs, rats were intravenously injected in the tail vein with GNPs with a diameter of 10, 50, 100, and 250 nm, respectively [82]. After 24 h, the rats were sacrificed and blood and various organs were collected for gold determination. The presence of gold was measured quantitatively with inductively coupled plasma mass spectrometry (ICP-MS) [82]. For all GNP sizes the majority of the gold was found to be present in liver and spleen. A clear difference was observed between the distribution of the 10-nm particles and the larger particles. The 10-nm particles were present in various organ systems including blood, liver, spleen, kidney, testis, thymus, heart, lung, and brain, whereas the larger particles were only detected in blood, liver, and spleen. The results demonstrate that tissue distribution of GNPs is size dependent with the smallest 10-nm NPs

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Figure 6.4. Cellular internalization of GNPs. Microglial cells (N9) were treated for 4 hours with two different functionalized rod GNPs (rod GNP-CTAB, rod GNP-INS) and urchin-like GNPs (GNP-URCHIN). Cells were fixed with paraformaldehyde and subsequently analyzed for the internalization of GNPs. (*Nanoscale hyperspectral images, where red signals denote a positive match with the spectral profile of the respective GNP, were performed using Cytoviva Image Analysis Software.) (See color insert.)

showing the most widespread organ distribution [82]. Similar findings were reported by Sonavane et al. [83], with 15-, 50-, 100-, and 200-nm GNPs. Interestingly, 15- and 50-nm GNPs were able to pass the blood–brain barrier [83]. After oral administration of metallic colloidal GNPs of decreasing size (58, 28, 10, and 4 nm) to mice, an increased distribution to other organs was observed [84]. The smallest particle (4 nm) administered orally resulted in an increased presence of gold particles in kidney, liver, spleen, lungs, and even the brain. The biggest particle (58 nm) tested was detected almost solely inside the gastrointestinal tract [84]. For 13-nm sized colloidal gold beads the highest amount of gold was observed in liver and spleen after intraperitoneal administration [67]. In another study, intravenously injected gold nanorods revealed that within 30 minutes these particles accumulated predominantly in the liver [85]. The PEGylation (coating with polyethylene glycol) of these gold nanorods resulted in a prolonged circulation [85]. Naturally, the surface molecules attached to the GNPs play a determining role in the final localization of the

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particles. By choosing a suitable ligand for surface modification, GNPs may be concentrated at certain locations in the living organism. Targeting tumorspecific surface receptors, for example, will bring the majority of the GNPs to the tumor cell membrane. There is not enough data for a complete overview of the effect of size, shape, and surface properties of GNPs on the cellular uptake, biodistribution, and cytotoxicity, but from the modest number of studies available to date one can conclude that these parameters are of great importance for biocompatibility and that different cell types have to be tested because of their different sensitivities toward GNPs. 6.3.3 Mechanism of Cellular Entry and Subcellular Distribution of Gold Nanoparticles Since the interaction between cells and GNPs depends strongly on the cell type and size and surface of the GNPs, there are cases when GNPs do not enter into the cells at all. PEG-modified GNPs, even after prolonged incubation for 24 h or after tenfold increase in concentration, were not found inside HeLa cells by electron microscopy, and no gold could be detected by absorption emission spectroscopy [86]. Under the right conditions, however, GNPs enter cells by a number of different routes. Not all of these ways are well understood, and it is not yet clear which factors favor any particular uptake mechanism (Figure 6.2). Receptor/clathrin-mediated endocytosis, followed by confinement to the endosomes, is the most commonly observed route of internalization [80, 86–88]. This process was studied for transferrin-coated GNPs and was found to be highly dependent on size, with the most efficient uptake occurring around the 50-nm size range [80, 88]. Shape dependency was also described, with spherical GNPs being internalized more readily than rodshaped GNPs [88]. Many factors, such as ratio of adhesion and membrane stretching and the membrane’s bending energy, may be involved in why size affects the uptake of the NPs. These parameters determine the so-called wrapping time, which describes how a membrane enclosed a particle [89]. Gao et al. [89] suggested that NPs that were ∼55 nm would have the fastest wrapping time and the receptor–ligand interaction can produce enough free energy to drive the NPs into the cell. This minimum wrapping time leads to a higher accumulation of the 55-nm NPs into the cell [89]. For the smaller NPs to go in, they must be clustered together. While a 50-nm GNP can enter the cell as a single NP, at least 6 of the 14-nm GNPs are required to cluster together before uptake [80]. For bigger NPs (>50 nm), the wrapping time is slower because of the slower receptor diffusion kinetics (more receptors were taken up during the receptor–ligand binding process and are not available for binding). Different proteins coated on the GNPs may lead to different results. The surface composition of the GNP can influence both the extent of internalization and the mechanism of uptake. For example, GNPs protected with

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tiopronin induce either endocytosis or adhesion to the cell membrane depending on the chemical composition of the NP surface. The tiopronin GNPs derivatized with ethylenediamine induce endocytosis, and the same NPs functionalized with polyethylene glycol derivative promote particle-cell adhesion [90]. The endosomal pathway can be avoided if the NPs are delivered to the cells via liposomes or if they are chemically modified with membrane-penetrating peptide sequences, such as the oligopeptide TAT from HIV or Pntn protein from Drosophila Antennapedia [86]. There are a number of reports on successful nuclear targeting and transfection [86, 91–93] in which case the particles appear to enter the cytosol either directly through the cell membrane or by endosomal escape [86]. In these cases, particles can be observed in the cytosol and/or in the nucleus [86]. For gene delivery, in particular, but also to label designated sites, the ability to direct NPs toward targets inside the cell is of great interest.

6.3.4

Concluding Remarks

More studies are necessary for a complete overview of the effect of size, shape, and surface properties of GNPs on the cellular uptake, biodistribution, and cytotoxicity. To date, the modest number of studies available indicate that these parameters play a very important role in the GNP–cell interactions. Also, it is likely that the different cell types have to be tested individually for their sensitivity toward GNPs. The long-term effect of GNPs on living organisms remains obscure.

6.4

NANOGELS AND ARTIFICIAL CHAPERONES

Polymeric delivery systems currently dominate the nanomedicinal field [94, 95]. Polymeric NPs are matrix-like, characterized by the ability to provide targeted and predictable controlled release of a “drug” of interest [96, 97]. The drug of interest either can be encapsulated within the polymer matrix or can be adsorbed or conjugated onto the surface of the NP. Intrinsic to this model of delivery, cargo release is mediated by two processes that take place over time: diffusion of the drug out of the polymeric matrix and/or degradation and digestion of the polymer. A number of different polymers, both natural (gelatin, alginate, chitosan, etc.) and synthetic (polylactide–polyglycolide copolymers, polyacrylates, polycaprolactones, etc.), have been utilized for the formulation of polymeric NPs. By changing the formulation composition or method of formulation, the NP size, charge, hydrophobicity/hydrophilicity, and drug release can easily be controlled. A diverse array of therapeutic and diagnostic agents, such as lipophilic or hydrophilic drugs, oligonucle-

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otides, DNA, peptides, proteins, and antigens or even imaging agents, such as quantum dots, can be encapsulated in polymeric NPs [98]. Nanogels, which constitute polymeric NPs, are strictly defined as crosslinked polymer networks that form stable nanosized hydrogel particles upon self-association, commonly observed in an aqueous milieu. Nanogels, or hydrogel NPs, are another example of nanomaterials with applications in drug delivery, specifically to the CNS because of their ability to cross the blood– brain barrier (BBB) [6, 99]. The BBB is known to be the most restrictive barrier in the body, preventing the entry of many small molecules and most macromolecular drugs into the CNS [100]. Given the prevalence and debilitating nature of neurodegenerative diseases such as Alzheimer disease (AD), Parkinson disease (PD), and amyotrophic lateral sclerosis (ALS), systems and strategies that improve drug delivery to specific intracellular sites within the CNS are fervently sought after. In many respects, nanogels have shown significant promise toward this avenue. The following addresses the most prominent of these advancements, with a particular emphasis on nanogels with the capacity to act as artificial chaperones and nanodelivery devices with potential therapeutic application in the diagnosis and treatment of neurodegenerative diseases, such as AD. 6.4.1 Nanodelivery and Artificial Chaperoning Properties of Nanogels Several different nanogel formulations have been developed for varying purposes, including the intracellular delivery of DNA [101] and proteins [102, 103] in a controlled and targeted manner, as well as for mediating chaperonelike activity [104]. In light of the fact that neurodegenerative diseases are commonly characterized by the accumulation of toxic protein aggregates [105], nanogels with the capacity to act both as artificial chaperones and drug delivery systems (DDS) hold a lot of promise as effective therapeutic strategies. In the early 1990s, long pullulan chains hydrophobized by the addition of cholesterol moieties at a frequency of 1–2 residues per 100 glucose units, termed cholesterol-bearing pullulans (CHPs), were shown to self-associate in water [106]. The self-aggregation of these amphiphilic polymers later led to the development of one of the first hydrogel NPs [107], which spontaneously forms due to noncovalent crosslinking between hydrophobic cholesterol moieties. In an in vitro model of protein refolding, carbonic anhydrase and citrate synthase were chemically denatured using guanidium chloride [108]. Interestingly, these nanogels were capable of preventing protein aggregation, while simultaneously stimulating the refolding of enzymes upon β-cyclodextrin binding in vitro, as evidenced by the recovery of activity. The mechanism of protein refolding was stipulated to be similar to that of an endogenous chaperone; that is, by catch and release, allowing the protein several new occasions to refold properly. Akiyoshi’s group was ultimately capable of demonstrating

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similar refolding capacities of GroEL, the main bacterial chaperone, and CHP using acid-denatured green fluorescence protein (GFP) as an in vitro model [109]. The direct therapeutic application of artificial chaperones to neurodegenerative diseases was first evidenced by the ability of CHP nanogels to bind and prevent the fibrillation of β-amyloid in vitro [110], a property later shown to be common among other polymeric NPs in vitro [111, 112]. Nanogels with artificial chaperoning capacity also have the capacity to encapsulate proteins for targeted release as nanodelivery devices, already shown for bovine serum albumin (BSA) protein and β-galactosidase using cationic amino-group (NH2) derivatives of these nanogels [102], in addition to interleukin-12 (IL-12) with implications in tumor immunotherapy [103]. Our group proposes the use of nanogel DDS for the encapsulation of not only small charged or hydrophobic proteins, but for hydrophobic drugs as well. The ability to specifically target toxic hydrophobic aggregates extra- and intracellularly, while synergistically resulting in the controlled release of drug at the site of insult or injury, is especially advantageous in diseases that require effective targeting of several processes, simultaneously for successful intervention, such as cancer and neurodegeneration. 6.4.2

Intracellular Delivery and Fate of Nanogels

The first study conducted in cells with CHP nanogels employed the cationic derivatives (CHPNH2) encapsulating protein conjugated QD655, a hybrid NP with implications in bioimaging [113]. It was first shown that nanogels of CHPNH2 with 15 amino groups per 100 glucose residues encapsulating QD655 conjugated with protein A (CHPNH2(15)-QD) were the most efficiently uptaken by HeLa cells, a cervical cancer cell line, after 3 hours of incubation when compared with CHP-QD and CHPNH2(9)-QD derivatives. The fate of CHPNH2(15)-QDs were further investigated using confocal laser scanning fluorescence microscopy (CLSFM), and Hasegawa et al. [113] clearly show that they are more efficiently uptaken within 3 hours compared with QDs on their own or liposomes encapsulating QDs. This same group has recently shown that CHPNH2-QDs remained detectable in rabbit mesenchymal stem cells (MSCs) for at least 2 weeks following internalization and had minimal effects on the chondrogenic activity of MSCs [114], in addition to labeling periodontal ligament cells for at least 3 weeks [115]. Using flow cytometry, Ayame et al. [102] demonstrated that CHPNH2 consisting of 17 amino groups per 100 glucose units resulted in the most effective delivery of fluorescein isothiocyanate (FITC) labeled BSA (FITC-BSA) to HeLa cells, supporting the use of nanogels for intracellular delivery of proteins. They further illustrated that CHPNH2-FITC-BSA internalized the most efficiently compared with FITC-BSA in the absence of carrier, or in the presence of a protein transduction domain (PTD)–based carrier or even using cationic liposomes as an alternate NP-based carrier system. This was shown to be the case in three other cell lines, NIH-3T3, cos-7, and CHO-K1, as well

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as for FITC-β-gal, in the place of FITC-BSA. Given that the mode of entry into cells will dictate the subcellular fate of nanogels (e.g., endosomes, cytosol, and nucleus), Ayame et al. [102] investigated potential mechanisms of entry for cationic nanogel protein carriers. Cellular uptake was shown to drop by over 60% upon placing cells at 4 °C, a method known to inhibit general endocytic mechanisms. Further investigation with inhibitors of clathrin-mediated endocytosis (i.e., ikarugamycin) and macropinocytosis (i.e., amiloride) suggested the primary implication of macropinocytic mechanisms in the uptake of CHPNH2. We studied the uptake and intracellular fate of nanogels (i.e., CHP and CHPNH2) in N9 cells, a microglial cell line. Microglia can act as carriers for drug delivery systems and can result in specific targeting to the site of injury where microglia tend to accumulate, as is observed in AD at the site of senile plaques [116, 117]. Using plate-based fluorometric studies, fluorescence microscopy (Figure 6.5), and fluorescence activated cell sorter (FACS) analysis, we have shown that tetramethyl rhodamine isothiocyanate (TRITC)– labeled cationic nanogels (CHPNH2) readily accumulate intracellularly within 60 minutes in microglial cells, whereas CHP nanogels slowly accumulate over 3 hours [118]. However, CHPNH2 in the absence of negatively charged interacting species (i.e., oligonucleotide cargo) causes toxicity to microglial cells, whereas CHP nanogels are nontoxic, which is in agreement with other studies regarding cationic nanogel delivery systems [99]. These observations tend to suggest that neutral nanogels are useful for the incorporation of hydrophobic, uncharged molecules, whereas cationic nanogels require the incorporation of charged species. Our present studies indicate that CHPNH2, although readily taken up by primary murine neural cells, does not colocalize with lysosomes, labeled with lysotracker green DND-22, within this time period. Further studies are being conducted in microglial cells to demonstrate whether or not, and at what time point, cationic and neutral nanogels colocalize with lysosomes. Importantly, our group has clearly shown that nanogels favorably interact with Aβ, in the monomeric and oligomeric forms, the latter of which has recently been shown to be the most toxic species [119]. Using Förster resonance energy transfer (FRET) studies, nanogels, both cationic and neutral, bind Aβ monomers and oligomers in vitro. Moreover, cationic and neutral nanogels prevent Aβ-mediated toxicity in mouse microglia (N9) and primary murine cells, respectively, as evidenced using the 3-(4,5-dimethylthiazol-2-yl)2,5-diphenyl tetrazolium bromide (MTT) assay. 6.4.3 Current Status in Clinical Trials and Future Perspectives of Nanogels Nanogels have already been employed as DDS in vivo and in clinical trials, primarily for cancer therapy. In mice with subcutaneous fibrosarcoma, subcutaneous injections of recombinant murine interleukin-12 (IL-12) encapsulated

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CHP

CHPNH2

0

1h

2h

3h

Figure 6.5. Cellular uptake of TRITC-labelled CHP-nanoparticles. N9 microglial cells were treated with tetramethyl rhodamine isothiocyanate (TRITC)-labeled CHP nanogels (12 nM) for 3 hours. Fluorescent images were taken at each time point: before treatment (T = 0 min) and at 3 time points therafter (T = 1, 2, 3 hours post-treatment). Scale bar in top left panel (20 μm) is representative for all images. (See color insert.)

in CHP nanogels, via incubation at room temperature, led to a prolonged elevation of IL-12 in the sera and resulted in significant growth retardation of the tumor [103]. Clinical trials for a vaccine incorporating the HER2 tumor antigen into CHP nanogels as an effective delivery system to antigen presenting cells (APCs), such as denditic cells and macrophages, has also been developed. It was shown that with biweekly subcutaneous administration of CHP-HER2, patients not only tolerated the vaccine, but induced CD8+ and CD4+ T-cell responses [120]. They further illustrated that HER2specific IgG antibodies were elevated in the sera of patients vaccinated

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with CHP-HER2 relative to controls and demonstrated that granulocytemacrophage colony-stimulating factor accelerated this humoral response to HER2 [121]. Given the minimal amount of information available presently with regard to the effects of nanogels on cellular functioning and more importantly on the subcellular targeting of nanogels for greater specificity, more studies need to be conducted at the single cell level. Importantly, nanogels appear to be excellent candidates for brain delivery, thus indicating that the majority of studies should focus on cell lines pertaining to brain cells, and more importantly on primary cultures, both human and rodent. The cellular uptake of specific types of nanogels will no doubt depend on the size, shape, and most importantly the composition. An investigation into the mechanisms of uptake not only at the blood–brain barrier, but also at the level of neurons and/or glial cells within the central nervous, will demonstrate which nanogels favor a cytosolic destination over an endosomal or nuclear, for example. Such studies are necessary if nanogels are ever to be proposed as specific drug delivery systems for targeting at the subcellular level.

6.5 BIOLOGICAL RESPONSES TO INTERNALIZED METALLIC NANOPARTICLES When metallic NPs enter cells they can induce inflammation in different cells [22] and, conceivably, there are several ways that NPs could exert their effects leading to or accelerating neurodegeneration: (1) by generating reactive oxygen species (ROS) and reactive nitrogen species (RNS) [17, 122–128], thereby altering the cellular redox status; (2) by induction or translocation of redox-sensitive transcription factors to the nucleus (e.g., NF-κB), which regulate proinflammatory genes, such as TNF-α and inducible nitric oxide synthase (iNOS) [129]; (3) by modification of proteins, lipids, and nucleic acids, which further stimulate the antioxidant defense system or even lead to cell death; (4) by excessively activating microglia and astrocytes; and (5) by promoting abnormalities in communications between organelles and post-translational modifications of proteins. Many diseases have been linked to oxidative stress and increased formation of ROS/RNS. Among these are lung, cardiovascular, and autoimmune diseases, as well as brain inflammation and aging [35]. As an underlying principle, cells exposed to inflammatory stimuli have a decreased capacity to deal with ROS/RNS, thus magnifying the extent of NP-induced injury. We are only now beginning to explore and understand the specific mechanisms underlying how low-dose NP exposure exerts these actions and consequently contributes to an enhanced rate and extent of neuroinflammation and neurodegeneration in the long term [34]. Oxidative stress is a state of redox disequilibrium that develops when ROS/ RNS production overwhelms the antioxidant defense system of the cell, thereby setting in motion the activation of redox signaling cascades that lead

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to toxicity [1, 130–133]. The extent of oxidative stress can vary from the very mild to the most extensive depending on the insulting agent(s) (i.e., type and size of NP), their concentration(s), as well as the duration of exposure and the cell types exposed [134]. Three major levels of oxidative stress are proposed by Nel et al. [3], known as the “hierarchical oxidative stress model.” In this model minor levels of oxidative stress are regulated mainly by the transcription/nuclear factor, erythroid 2-related factor 2 (Nrf2), which controls the transcriptional activation of about 200 antioxidant and detoxification enzymes [135, 136]. Among these genes are heme oxygenase 1 (HO-1), glutathione S-transferase (GST) isoenzymes, NADPH quinine oxyreductase (NQO1) catalase, superoxide dismutase, and glutathione peroxidase (GPx) [135–138]. Interestingly, our recent studies show remarkable changes in antiapoptotic and pro-apoptotic genes induced by QDs [43]. Among the antioxidant defense systems, GPx was the most susceptible to the treatment with QDs and was completely abolished after 24 hours. When this preventative antioxidant defense system is insufficient, for instance, if long-term NP exposure is overwhelming the system, a pronounced increase in ROS can result and subsequently cause the release of proinflammatory cytokines [139–141]. This proinflammatory stage is mediated by the redox-sensitive MAPK and NF-κB cascades. Further increase in the severity of oxidative damage leads to morphological changes in mitochondria and also, as we have recently demonstrated, to an enlargement of lysosomes and activation of lysosomal enzymes [122]. Our pilot studies using a mouse-derived microglia cell line (N9 cells) show that a large number of NPs are internalized by these cells and, subsequently, glial cells are activated. Enzymatic activity in this activated state is significantly enhanced, supporting the view that activation of microglia leads to acidification of lysosomes and consequent activation of lysosomal proteases [142]. Evidence has been accumulating that among other free radicals, reactive nitrogen species (RNS) have been associated with the etiology and progression of inflammation in different neurodegenerative disorders and environmental insults [143, 144]. Nitric oxide (NO), a bioactive free radical, plays an important role in the development of neurodegenerative diseases such as Alzheimer disease (AD), Huntington disease, Pick disease, epilepsy, schizophrenia, and cerebral ischemia [145]. At low concentrations, NO plays a role in neurotransmission, synaptic plasticity, and vasodilatation, while at higher concentrations it is implicated in neurodegenerative disease pathologies [146, 147]. NO is enzymatically formed from l-arginine by the enzyme nitric oxide synthase (NOS). The calcium-dependent NOS present in neurons is constitutively expressed, whereas inducible nitric oxide (iNOS) expressed in astrocytes and microglia is inducible on the transcriptional level by various stimuli [143, 148]. Resident glia cells of the brain express iNOS and produce high levels of NO in response to a wide variety of proinflammatory and degenerative stimuli [149, 150]. Few cytokines, such as IL-β and IFN-γ, alone can induce iNOS; TNF-α is usually required in conjunction with IL-β and IFN-γ. In fact,

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the inducibility of NO depends on the cytokine, the cell type, and the species, as well as where the cells are derived from [149]. When NPs enter the cell, mechanisms involved in cell and tissue repair [151], including enzymatic systems maintaining normal cellular redox status, can be activated. These defensive systems include glutathione (glutathione/ glutathione peroxidase or glutathione/glutaredoxin) and thioredoxin (thioreduxin/thioredoxin reductase or thioredoxin/peroxiredoxin) systems. These systems complement each other and sometimes overlap in cytoprotection. Thioredoxin reductase is a member of the selenium-containing oxidoreductase family [152] and its role in regulating levels of oxidative stress is well recognized [153].

6.6

CONCLUSION

Inadequate drug delivery is one of the most important factors limiting the utilization of molecular medicines today. Effective targeting of drugs toward specific cell types and subcellular compartments has proved difficult but significant advances were made in achieving this goal, mainly in cancer therapy. Quantum dots seem to be particularly promising components of the highly sensitive and relatively simple diagnostic devices for the analyses of biological specimens ex vivo and in living cells due to their unique photophysical properties. GNPs are not only excellent markers of subcellular site distribution but also serve as extremely useful components of sensors for testing functional cellular responses and delivering anti-inflammatory and regeneration-promoting drugs. We should carefully examine possible side effects of NPs, particularly those with metallic cores. We are looking forward to new clinical trials with biocompatible and biodegradable polymeric NPs, delivering therapeutics with improved efficacy, as well as additional synergistic chaperoning activity, ultimately promoting the recovery of the compromised central nervous system.

ACKNOWLEDGMENTS We gratefully acknowledge the granting agencies National Science Research Council and Canadian Institutes of Health Research for support of the research carried out by the authors of this chapter.

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CHAPTER 7

Intracellular Fate of Plasmid DNA Polyplexes KEVIN MAIER and ERNST WAGNER Pharmaceutical Biotechnology, Munich Center for System-Based Drug Research, and Center for NanoScience, Ludwig-Maximilians-Universität, Munich, Germany

7.1

INTRODUCTION

Gene delivery by synthetic vectors started almost 50 years ago, when Smull and Ludwig [1] demonstrated that uptake of poliovirus ribonucleic acid is enhanced by interaction with plaque-forming basic proteins. From this point of time, up to date many cationic polymers were developed with the ability to condense plasmid DNA (pDNA) and promote its cellular uptake (e.g., diethylaminomodified dextran, poly-l-lysine (PLL), polyethylene imine (PEI), chitosan, cationic dendrimers). These complexes, built up from synthetic polymers and DNA, have been named polyplexes [2]. Shortly after their discovery it became clear that polyplexes possess advantages and disadvantages at the same time as compared to viral gene vectors. Flexibility concerning the payload is one of the major benefits of polyplexes. The most severe handicap of the polyplexes is their far lower transfection efficiency. To improve this, it is necessary to understand the barriers these polyplexes are confronted with, during their journey from the extracellular environment to the site of action—the nucleus in the case of pDNA, the cytosol in the case of RNA and siRNA. Cell entry, endolysosomal escape, cytoplasmic trafficking, DNA polyplex dissociation, and transport to the nucleus have been identified as the main hurdles that have to be overcome by the polyplexes. Numerous efforts were taken to address all these problems. Polymers used at the beginning of synthetic gene delivery were quite simple but tend to become more and more complex and chemically dynamic in structure. According to the different cellular compartments (each with a different chemical environment) in which the polyplexes Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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find themselves during the delivery of their cargo, they are supposed to change their chemical properties to react in the required manner. Since these new polyplexes behave almost like their natural viral counterparts, they are already called “synthetic viruses.” This chapter reviews state-of-the-art concepts and insights into the intracellular behavior of various polyplexes used for gene delivery.

7.2

CELLULAR BINDING AND UPTAKE

The first step of gene delivery by synthetic vectors is binding of the polyplex to the plasma membrane. If no targeting ligand is used, this binding is mediated through electrostatic interaction between the negatively charged cell surface and the positively charged surface of the polyplex. Little is known about the exact role of different cell surface components like glycosaminoglycans (GAGs), proteoglycans, integrins, and phospholipids. Mislick and Baldeschwieler [3] showed in 1996 that binding of PLL polyplexes to proteoglycan-deficient CHO mutant cells is strongly reduced and therefore suggested that heparin sulfate proteoglycans are involved in binding. Another evidence for their hypothesis was the limited binding of polyplexes in the presence of competitive soluble heparin and heparan sulfate in the medium. Chondroitin sulfate seems to play a minor role and the addition of hyaluronic acid had no effect on binding. A special role is supposed for adhesion of the polyplexes to syndecans (transmembrane heparan sulfate proteoglycans). Binding should effect enclosing the particle by lateral diffusion of the syndecans, clustering into cholesterol-rich rafts. Furthermore, these clusters cause activation of protein kinase C (PKC) and linker mediated binding of actin to the cytoplasmatic end of the syndecan molecule. The resulting fibers were supposed to pull the particle into the cell [4].

7.2.1

Uptake Mechanisms

Due to their size and charged surface, polyplexes cannot diffuse passively through the plasma membrane. All types of polyplexes share endocytosis as a common uptake means [5–7]. In literature at least five different ways of endocytosis are described—phagocytosis, clathrin-mediated endocytosis, macropinocytosis, caveolae-mediated endocytosis, and clathrin-caveolae-independent endocytosis [8]. It is necessary to understand these internalization pathways, as they vary in size, coating, and most importantly in further processing of the internalized vesicles after uptake. The different uptake mechanisms and further processing of the vesicles are shown in Figure 7.1. For example, uptake through caveolae-mediated endocytosis has the advantage that the polyplex is not internalized in vesicles, which can lead to lysosomal degradation [9]. Which endocytotic mechanism by which the polyplexes are

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Figure 7.1. Schematic illustration of polyplex uptake pathways and further processing of the internalized vesicles. Except for caveolae-mediated endocytosis, all other cellular entries lead at least partly to fusion of the vesicles with lysosomes and therefore to acidic degradation of the cargo.

internalized is dependent on many parameters, like particle size, the polymer employed, and the cell line [10–13]. 7.2.1.1 Nonspecific Uptake As described above, standard polyplexes are bound to the plasma membrane by electrostatic interaction. They are supposed to be taken up mainly by clathrin-dependent endocytosis (CDE) and caveolae-mediated endocytosis (CME). This assumption is based on many transfection experiments with different chemicals inhibiting different uptake pathways. Colocalization studies with transferrin (evidence for CDE) and choleratoxin B (evidence for CME) also revealed that linear PEI (LPEI) and branched PEI (BPEI) polyplexes are included via both mechanisms [13]. Moreover, by inhibiting one specific pathway, for example, CDE with chlorpromazine or caveolae-mediated endocytosis with filipin III, cell line dependence for the favored uptake mechanism could be shown. The main player for successful transcription in COS-7 cells with LPEI and BPEI was CDE, whereas in HUH-7 cells both pathways are equally important for transfection with BPEI. For LPEI polyplexes again CDE was the main contributor to the transcription process. In HeLa cells, independent of the polymer employed, both internalization pathways led to effective transcription with a small preference for the caveolae-mediated route. Only a few studies have been done to

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reveal the mode of endocytosis for PLL polyplexes. Lühmann et al. [14] showed that in COS-7 cells the internalized PLL-g-PEG polyplex did not colocalize with the early endosome antigen-1 (EEA-1), which is a marker for the endolysosomal pathway. Although the cellular uptake of the nanoparticle was inhibited about 50% with genistein (inhibitor of CME), there was no decrease in transfection efficiency. Furthermore, no colocalization of the polyplexes with caveolin 1 protein could be observed. This result led to the assumption that the impact of caveolin-mediated uptake is low. As inhibition of the clathrin-mediated endocytosis and macropinocytosis decreased transfection efficacy about 17% and 24%, they assumed that several pathways were participating in the uptake of the polyplexes and led to transfection. Cholesterol depletion experiments demonstrated the main uptake mechanism for the commercially available polyamidoamine (PAMAM) dendrimer superfect (Qiagen) in an endothelial cell line. Without cholesterol the binding of the dendriplexes as well as the transfection efficiency was strongly decreased [15]. After reinstatement of the membrane cholesterol the efficacy was rebuilt. In addition, confocal microscopy studies showed colocalization with the membrane raft ganglioside GM1. There is strong evidence that the uptake is provided mainly through caveolae-dependent processes. Another widely used polymer for gene delivery is the polysaccharide chitosan, which is built up from repeating units of glucosamine. In 2009 Nam et al. [16] suggested after endocytotic inhibitor experiments and colocalization studies in HeLa cells that at least three uptake mechanisms (macropinocytosis, clathrin-mediated endocytosis, and caveolae-mediated endocytosis) are involved in the internalization of hydrophobically modified glycol chitosan polyplexes (HGC). The uptake of alginate chitosan polyplexes seems to be cell line dependent [17]. HEK 293T cells internalize the particles through CDE, whereas the uptake in CHO cells is predominantly provided through CME. Interestingly, the latter led to no gene expression, because the cargo was entrapped in caveosomes and could not escape. In contrast, gene expression could be observed in HEK 293T cells, consistent with release from acidified endosome provided through the socalled proton sponge effect (see Section 7.3.1). Another important parameter influencing the uptake mode is the size of the nanoparticles. Medium sized particles (50–200 nm) are internalized through a clathrin-dependent and/or a caveolae-mediated mechanism, whereas nanoparticles larger than 200 nm are taken up mainly by macropinocytosis. For very small particles a caveolaemediated pathway could be observed [10–13, 18]. 7.2.1.2 Targeted Uptake In contrast to untargeted polyplexes, a targeting ligand causes binding of the polyplex to a special membrane region, where the appropriate receptor is located. This should lead to a selective uptake pathway as well as a special further processing and intracellular trafficking. Indeed, in general, ligands, except for a few, result in clathrin-dependet endocytosis [19, 20]. So far many different endogenous and exogenous ligands for targeting of polyplexes have been investigated (Table 7.1). Lactose targeted PLL is inter-

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TABLE 7.1 Targeting Ligands Used for Polyplexes Ligand Carbohydrates Lactose Galactose Mannose Peptides Tet1 HIV-TAT Penetratin Antibodies anti-EGFR anti-CD3 anti-Her2 Endogenous ligands EGF Transferrin TGF-α

Reference 21 23 24 25 26 27 28 29 30 22 6 31

nalized through clathrin-dependent processes, whereas glycosylated BPEI is taken up mainly through caveolae-mediated mechanisms [21]. To avoid unspecific binding of polyplexes (caused by electrostatic interaction of the cationic polyplexes with the overall negatively charged cell surface) and trigger ligand– receptor mediated binding, the polyplexes have to be shielded. The commonly used reagent for shielding is polyethylene glycol. While the binding of the shielded polyplexes to the cell surface without a targeting ligand is highly reduced, targeting is rebuilding effective binding in the presence of shielding. A widely used targeting ligand is transferrin. It was shown that transferrin targeting increases cellular binding and the uptake rate as well as transfection efficiency in K562 cells [6]. The 7-kDa protein EGF is another well-studied endogenous targeting ligand, which enhances cellular binding and therefore transgene expression in human adenocarcinoma (KB) cells [6]. EGF targeting leads to faster binding and uptake compared to untargeted polyplexes in HUH-7 cells. As detected by novel single-particle tracking microscopy, 5 minutes after incubation 50% of targeted polyplexes were internalized, whereas only 5% of untargeted polyplexes were taken up at this time point [22]. 7.2.2 Phases of Polyplex Internalization and Further Intracellular Routing Single-particle experiments revealed an internalization process that can be divided into three different phases. During phase 1, which includes the extracellular binding and internalization process, the particles moved slowly with a drift mediated by the actin cytoskeleton. In this phase the endosomes are

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(A) targeted Polyplex

Membrane protein

(B)

3 μm Phases 1+2

Phases 3 Actin

Endosome

Microtubule Phase 1

Phase 2

Phase 3

instantaneous velocity [μm/s]

(C) Phases 3 3 2,5 2 1,5 Phases 1 Phases 2 1 0,5 0 0 50 100 150 200 250 time [s]

Figure 7.2. Cell binding, cellular entry, and further trafficking of an EGF-PEG-PEI polyplex. (A) Scheme of the three internalization phases. Phase 1 includes cellular binding and uptake, mediated by the actin cytoskeleton. Phase 2 shows free diffusion of the internalized vesicle. Phase 3 is characterized by fast directed motion along microtubules. (B) Trajectory plot of the particle with a frame rate of 300 ms. (C) This panel shows velocity of the vesicle during the different phases. The corresponding video can be found at http://www.nature.com/mt/journal/v15/n7/suppinfo/6300176s1 .html. (Adapted from de Bruin et al. [22].)

formed and tensioned by actin over membrane proteins. In phase 2, the vesicles are waiting to combine with the microtubule transport system. Normal diffusion in the cytoplasm with random movement is characteristic for this phase. During phase 3, a dramatic increase in particle velocity and active, directed transport of the vesicles along microtubules could be observed. Except for a shortening of phase 1 the same phases for EGF targeted as well as for untargeted polyplexes could be monitored [22]. Figure 7.2 shows the trajectory of the vesicles during these three different phases. These observations are in accordance with previous studies done by Kulkarni et al. [32] in 2005. They showed two different phases of intracellular polyplex movement by spatiotemporal image correlation spectroscopy: a long term phase (about 60 seconds) with low velocity and random motions of the particles, and a short-term phase (about 10 seconds) characterized by a high velocity (0,05 μm/s) and directed motion of the vesicles. They also suggest that this directed movement is correlated with transport of the endosomes along microtubules. After the polyplexes have overcome the hurdle of the cytoplasmic membrane and the vesicles have reached the cytoplasm, further processing can involve

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recycling back to the cell surface, fusion to degenerative vesicle forms like lysosomes, as well as transport to organelles like endoplasmic reticulum or Golgi apparatus. As shown in Figure 7.1, this is primarily dependent on the internalization pathway.

7.3

ENDOSOMAL ESCAPE

In Figure 7.1 it becomes apparent that four of the five internalization pathways for polyplexes lead at least partly to fusion of the internalized vesicles with lysosomes. In this cell compartiment with a typically acidic pH between 4 and 5, the cargo of the vesicles is degraded. It is generally believed that cellular uptake of polyplexes has a lesser impact on transfection efficacy than further processing [33]. Dinh et al. [34] predicted when using PEI polyplexes that only 1% of the internalized DNA molecules reach the nucleus, the final aim for DNA in gene delivery. Endosomal escape was identified as one of the major bottlenecks in gene delivery by synthetic vectors [35, 36]. In contrast to lipoplexes, classical cationic polyplexes do not exhibit hydrophobic regions and therefore cannot release their cargo from the endosomes by membrane fusion processes. A lot of different strategies that help the polyplexes to get over this hurdle have been investigated so far [37]. The most common ones are presented in the next sections. 7.3.1

Protonable Amines

Polyplexes built up from cationic polymers like BPEI, LPEI, and histidinylated PLL possess protonable primary, secondary, or tertiary amines. Such protonable amines possess a pKa between 5 and 7 and therefore can buffer acidification of the late endosomes by taking up protons. This buffered acidification of late endosomes goes along with concurrent influx of chloride to maintain charge neutrality and results in an increased ionic strength inside the endosome. To balance this, water is accumulating passively inside the endosome. Therefore the pressure inside the endosome increases more and more until the membrane bursts and the content of the endosome is released into the cytosol [38]. Today this mechanism is known as the “proton sponge effect.” Akinc et al. [39] showed that the transfection efficacy of N-quarternized PEI (lacks protonable amines) is reduced by approximately 100-fold. The same observation was made by using the vacuolar proton pump inhibitor bafilomycin A1. Chloroquine is an osmotically active lysosomal buffer reagent, which increases the transfection efficiency of PLL polyplexes between twofold and more than 100-fold [40, 41]. Other neutralizing but not osmotically active inhibitors did not enhance gene transfer. This reveals that blocked lysosomal degradation of DNA (due to prevention of acidification) and endosomal escape contributed to the beneficial effect. No effect of chloroquine could be observed with PEI polyplexes that can buffer the endosomal pH by

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themselves. More recent findings suggest that the osmotic effect by the proton sponge action of PEI is not sufficient to explain endosomal escape; the protonated cationic PEI residues appear to be directly involved in the membrane disruption process. For example, irreversible shielding of PEI polyplexes by PEGylation strongly interferes with this process and pH-sensitive PEG attachment recovers the activity [42]. 7.3.2

Photoinduced Endosomal Release

Photochemical disruption of the endosomal membrane [43, 44] is another strategy to overcome lysosomal degradation. In this case the polyplexes are internalized by the cells together with amphiphilic photosensitive reagents (e.g., disulfonated meso-tetraphenylporphine [TPPS2a]) that were present in the transfection medium (Figure 7.3). After internalization the membranebound photosensitizer (PS) is activated by illuminating the cells. The PS mediates energy transfer to molecular oxygen, generating radical singlet oxygen inside the endosomes. Subsequently, these radicals lead to oxidative damage of the endosomal membrane, resulting in rupture of the vesicular membrane and release of the polyplexes into the cytosol. Using this so-called photochemical internalization (PCI) technique, the transfection efficiency of EGF targeted PEI polyplexes could be increased in HUH-7, HepG2 and A431 cells between twofold and 600-fold [45]. For in vivo application the PS could be

Photosensitizer (PS)

Polyplex

early Endosome

light

h*v

1

PS*

PS 1O 2 3PS*

O2

Figure 7.3. Photochemical internalization and operating mode of a photosensitizer for promoting endosomal escape. After light activation, the photosensitizer mediates energy transfer to molecular oxygen, resulting in radical singlet oxygen inside the endosome. The built radicals attack the endosome membrane and lead to its rupture. Thus liberation of the polyplex into the cytosol is accomplished.

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internalized inside the polyplexes [44]. In the case of PLL polyplexes, which lack proton sponge mediated liberation, de Bruin et al. [46] showed a fast release of the cargo into the cytosol (within about 50 seconds) after activation of the photosensitizer. 7.3.3 Membrane Disruptive Peptides The use of membrane destabilizing peptides was the first technique to promote effective release of polyplexes entrapped in endosomal vesicles. Up to date a lot of natural and synthetic peptides have been used (Table 7.2) to trigger this endosomal release. These peptides can be divided into two classes. Group one contains peptides with an inherent membrane lytic activity (e.g., melittin, KALA), whereas the other peptides only show lytic activity at acidic pH (e.g., GALA, INF). The selective lytic activity of the latter class is explained by pH-dependent conformational changes in peptide structure. Only at acidic pH can the peptides form an alpha-helix, which is required for lytic activity. The alpha-helical conformation is necessary for amphiphilic interaction with the membrane phospholipid bilayer and building pores in it that lead to rupture [47]. Wagner et al. [48] showed that the transfection efficiency of PLL-transferrin polyplexes could be enhanced about 100 times when coupling influenza virus hemagglutinin peptide HA2 to the polymers. Furthermore, Rittner et al. [49] designed new basic amphiphilic peptides (ppTG1, ppTG20) that can be used as singlecomponent vectors by combining DNA complexing ability and endosomal release function in one molecule. The bee venom peptide melittin also enabled release of polyplexes into the cytoplasm [50]. As melittin has an inherent membrane disruptive property, not only at acidic pH inside the endosome but also at neutral pH inside the cytosol, it increases cellular toxicity of the polyplexes. To overcome this problem investigators either introduced acidic residues into melittin [51] or composed a chemically modified melittin by coupling dimethylmaleic anhydride (DMMAn) to the lysine residues of melittin [52, 53]. When the polyplexes are acidified inside the endosome, this pH labile

TABLE 7.2

Membrane Disruptive Peptides

Peptide Natural peptides Melittin INF-HA2 HRV2-VP1 PFO Synthetic peptides NMA-3 KALA GALA ppTG1 ppTG20

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binding of the protection groups is cleaved and the lytic activity is regained. This resulted in less toxicity compared to PLL-melittin polyplexes and even to single PLL polyplexes. Gene expression was enhanced about 1800-fold compared to simple PLL polyplexes [53]. The concept has also been adapted for efficient cytosolic delivery of siRNA [54, 55].

7.4

DNA POLYPLEX DISSOCIATION

For various reasons the release of the DNA from the polyplex is necessary for successful transcription. It remains controversial whether this has to occur before or after nuclear entry. By using quantum dot/FRET assays, the kinetics of unpackaging has been monitored and occurs in several compartments [59]. On the one hand, flexible small size for nuclear import and accessibility of transgene DNA in the nucleus by the transcription machinery (transcription factors, polymerases) is important. For example, a standard 50–100-nm polyplex would be too large to enter the nucleus through a nuclear pore, either passively (10-nm pore limit) or actively (maximum pore diameter of around 30 nm) [60]. On the other hand, the dissociation of the vector from the DNA in the cytosol is a critical step in efficient gene delivery, because after release free cytosolic DNA has no protection against degradation enzymes like nucleases. Therefore the dissociation step should occur as late as possible, optimally just before penetration of the DNA into the nucleus; this happens, for example, in the case of adenoviral infection. The release of the electrostatic bound nucleic acid from positively charged polymers is provided by anionic cellular components like overall positively charged proteins, lipids, and last but not least RNA molecules [61–63]. Microinjection experiments revealed that the dissociation of PEI polyplexes in contrast to lipoplexes can also happen in the nucleus, presumably because of the exchange of the transgene with cellular DNA [64]. Schaffer et al. [65] demonstrated an enhancement of transfection efficiency when using low molecular weight PLL for transfection in comparison with the high molecular weight counterpart. They predicted that this enhancement arises from easier dissociation of the polyplexes in the cytosol. This has also been postulated for the more effective LPEI in comparison to BPEI [66]. Most recently, intranuclear FRET analysis has been used for analysis of pDNA decondensation from nonviral carriers [67]. pDNA polyplex decondensation after nuclear entry was the major determining factor for transgene expression. Besides better DNA release, lower cytotoxicity is another great advantage of smaller or degradable polymers. Again, different strategies have been developed to overcome these obstacles. 7.4.1

pH-Sensitive Polymers

Polyplexes that are cleaved into smaller metabolites after acidification of the endosome were developed by Knorr et al. [68,69]. These polymers are based

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on small 800-Da PEI subunits. These subunits were chemically covalently conjugated to larger molecules over pH-sensitive acetone ketal crosslinkers such as 2,2-bis(N-maleimidoethyloxy)propane (MK) or 1,1-bis-(2-acryloyloxy ethoxy)-[4-methoxy-phenyl]methane) (BAA). These polyplexes showed better transfection efficiency than analogs built up with noncleavable linkers at high vector to plasmid ratios. Moreover, the cytotoxicity of the pH-sensitive polyplexes was greatly reduced as shown with methylthiazoletetrazolium (MTT)/thiazolyl blue assay in Neuro 2a and B16F10 cells. 7.4.2

Biodegradable Esters

Linkage of small subunits to larger polymers over biodegradeable ester bonds is another promising strategy [70–72]. Ester bonds are hydrolyzed by water perhaps and even intracellular esterases can cleave the bonds. In contrast to the above-mentioned pH-sensitive acetal bonds, ester bonds are more stabile with a longer half-life. Many parameters can be varied (e.g., changes in structure and molecular weight of the subunits) to obtain the desired degradation kinetic. Furthermore, amine density of the subunits as well as the grade of crosslinking can influence the release of DNA [73–75]. Russ et al. [74] described a polymer based on generation two PPI dendrimers as the core, grafted with PEI 800 Da and crosslinked with hexane-1,6-diol diacrylate, that exhibits a transfection efficiency comparable to or even better than PEI, but with lower cytotoxicity. 7.4.3

Intracellular Reducible Polyplexes

These polyplexes capitalize for dissociation on the different redox potentials between the oxidizing extracellular environment on the one hand and the reducing cytosolic environment (mediated by glutathione) on the other hand. The polyplexes are stable until they are released into the cytosol. Once they reach the cytosol the disulfide bonds between the subunits are cleaved and thus the DNA release is facilitated and toxicity is reduced. For cleavable PEI (CLPEI50%) polymers built up from smaller 1800-Da PEI subunits and crosslinked with 3,3′-dithiopropionimidate dihydrochloride (DTBP), cleavage in the presence of 3-mM glutathione—comparable to intracellular concentrations—was evident [76]. Furthermore, these conjugates exhibited lower toxicity than PEI 25 kDa. These data are in accordance with that of another group that reveals high transfection efficacy of an analogous biodegradeable polyethylene imine sulfide polyplex and also low cytotoxicity [77]. Moreover, Lee et al. visualized the intracellular degradation of the polyplexes in HeLa cells using fluorescence quenching of a conjugated BODIPY-dye. All these studies to enhance the cytosolic release of the DNA into the cytoplasm are quite promising, but as already mentioned, once free in the cytosol, the DNA is no longer protected against degredation enzymes like proteases. At this stage, perhaps an even more important bottleneck for synthetic gene

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delivery by cationic polyplexes is reached—the transport of the DNA into the nucleus.

7.5

CYTOSOLIC TRAFFICKING TOWARD THE NUCLEUS

The short half-life of free cytosolic DNA makes clear why a fast transport of the DNA into the nucleus is desired. In HeLa and COS-1 cells pDNA halflives are between 50 and 90 minutes, as monitored by fluorescent in situ hybridization (FISH) experiments [78]. Thus it is not surprising that intranuclear microinjection of plasmids encoding for β-galactosidase into COS-7 cells results in much higher gene expression than when the same amount of plasmid is injected into the cytosol [79]. Highly impeded velocity of the DNA in the cytoplasm is one obstacle for fast nuclear import [80]. As microinjected oligonucleotides are distributed homogeneously in the cytoplasm, size is the main factor of impeded velocity rather than interaction with cytosolic proteins [81]. The cytoskeleton built up by microtubules, intermediate filaments, and microfilaments forms a dense network with narrow meshes, which prevents free diffusion of plasmids larger than 250 base pairs [82]. For this reason strategies to actively transport the DNA toward the nucleus (either within vesicles or after intracytosolic release) [83] are highly desired.

7.6

NUCLEAR IMPORT

The next hurdle that has to be overcome by the plasmids on their way into the nucleus is the nuclear envelope. This double membrane is interspersed by nuclear pore complexes (NPCs) with a diameter of about 10 nm. Molecules, smaller than 40 kDa, can diffuse passively through these pores. Larger molecules have to be transported by an active energy-dependent process, which goes along with conformational changes of the NPCs and widening of the pores to a diameter of about 30 nm [84]. Now the pores are able to transport compact macromolecules as large as 50 MDa [85]. Plasmids used for gene delivery have a molecular weight between 2 and 10 MDa and a flexible size, with an extended length of around 1 μm but a small helix diameter of 2 nm. To initiate this energy-dependent uptake mechanism, a targeting signal is necessary [86]. For all these reasons it becomes clear why only 1 out of 1000 cytosolic cDNA molecules is estimated to reach the karyoplasm [78]. A number of efforts were made to facilitate this critical step in gene delivery by polyplexes. 7.6.1

Cell Cycle Dependence

Microinjection of naked DNA in HeLa cells revealed that transfection efficiency was dependent on the cell cycle stage [87]. After injection of 10 ng naked plasmid DNA, 28% of the nondividing cells showed transgene expres-

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sion, whereas 50% of the cells that divided after injection expressed the gene encoded on the plasmid. Although even growth arrested cells could be transfected with PEI, the main entry of the transgene into the nucleus happens passively, when the nuclear envelope is temporarily degraded during mitotic cell division [88]. Brunner et al. [89] demonstrated that transfection efficiency for transferrin-polylysine and transferrin-polyethylenimine is strongly dependent on the cell cycle stage at the time of transfection. Compared to cells transfected during G1 phase, cells transfected during S or G2 phase exhibited 30–500-fold higher transgene expression in K562 cells. Further studies showed that this cell cycle dependence is not valid for all cationic polymers. In contrast to polyplexes formed with BPEI, polyplexes built up with LPEI do not display such strong cell cycle dependence in transfection efficiency [90]. Therefore it can be suggested that LPEI polyplexes exhibit improved nuclear import characteristics and/or improved nuclear unpackaging compared to BPEI and PLL formulations. 7.6.2

Nuclear Localization Signal (NLS)

As already described, macromolecules over 40 kDa can pass the NPCs only in an energy-dependent process. This mechanism is activated by nuclear localization signals, built up from small, mostly cationic, peptide sequences. Large numbers of different NLS sequences are known (e.g., SV40 large T-antigen, M9 human nuclear ribonucleoprotein A1, HIV type-1 viral protein R [Vpr], human T cell leukemia virus type 1 [HTLV]). Macromolecules containing such sequences are bound to import proteins (importins) that mediate the translocation of the complexes into the karyoplasm. In contrast to the passive cellcycle-dependent nuclear import of DNA, this is an active transport process. Peptides containing such NLS sequences can be conjugated either to the polymer vector [91, 92] or, alternatively, to the plasmid. Covalent conjugation of an NLS sequence derived from simian virus 40 to the DNA over a cyclopropapyrroloindole crosslinker induced nuclear accumulation of the plasmid DNA in digitonin permeabilized HeLa cells. It seems that the amount of NLSs per kilo base pair is an important factor [93]. Sebestyen et al. [93] used at least 100 NLS sequences per kilo base pair to achieve high nuclear accumulation of the plasmids. Because of that high modification grade, transgene expression was completely abolished [93]. A second binding possibility is mediated just by electrostatic interactions between cationic NLS sequences and the negatively charged nucleic acid [94]. Although it is a promising idea to enhance nuclear import with the usage of NLSs containing peptides, it now becomes clear that its application is limited by many different factors [95, 96]. 7.6.3

DNA Sequences

In contrast to the above described technique to transport DNA via coupling to NLS peptides (trans-acting), a further mechanism of nuclear DNA import is DNA sequence dependent (cis-acting). Some sequences of plasmids are

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supposed to have a certain affinity to proteins like transcription factors. Such proteins often carry NLS sequences, which lead to an energy-dependent nuclear import of the associated DNA [97]. An example for such a DNA sequence is the 10 base pair long κB binding site that can be found in the promotor and enhancer regions of many plasmids. NF-κB is an NLS containing transcription factor with high affinity to this κB sequence. By microinjection of plasmids into the cytoplasm of HeLa cells, Mesika et al. [98] demonstrated that these κB sites not only facilitated the nuclear penetration of the plasmids but enhanced their migration through the cytoplasm toward the nucleus in a dynein-dependent manner. Futhermore, sequences from the SV40 viral genome containing the SV40 origin of replication, early and late promoters, supported nuclear import [99]. This sequence-specific import could be inhibited by energy depletion as well as by the addition of NPC inhibitory agents like wheat germ agglutinin. Moreover, Dean [99] showed that before nuclear uptake the DNA is transported and accumulated in the nuclear periphery. The uptaken DNA was colocalized inside the karyoplasms with SC-35 splicing complex antigen and thus it is suggested that the DNA uptake is linked to transcription. Indeed, the inhibition of transcription blocked nuclear uptake. Despite these interesting results, nuclear import of DNA into the nucleus is one of the least understood mechanisms of synthetic gene delivery and remains a major bottleneck.

7.7

CONCLUSION

Although due to new techniques, like single-particle tracking, great advances in elucidating the different steps of internalization and intracellular processing of polyplexes were made, some mechanisms still remain quite unclear. These not well understood steps, for example, the penetration of DNA into the nucleus, seem to be major limiting factors in efficient gene delivery. No polyplex has been designed so far to address all known hurdles at the same time, while combining high efficiency (independent from cell line) with low cytotoxicity. Nevertheless, substantial progress has been made for reaching this final aim in the last decade.

REFERENCES 1. Smull, C. E. and Ludwig, E. H. Enhancement of the plaque forming capacity of poliovirus ribonucleic acid with basic proteins. J. Bacteriol. 84: 1035–1040 (1962). 2. Felgner, P. L., et al. Nomenclature for synthetic gene delivery systems. Hum. Gene Ther. 8: 511–512 (1997). 3. Mislick, K. A. and Baldeschwieler, J. D. Evidence for the role of proteoglycans in cation-mediated gene transfer. Proc. Natl. Acad. Sci. U.S.A. 93: 12349–12354 (1996).

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58. Wyman, T. B., et al. Design, synthesis, and characterization of a cationic peptide that binds to nucleic acids and permeabilizes bilayers. Biochemistry 36: 3008–3017 (1997). 59. Chen, H. H., et al. Quantitative comparison of intracellular unpacking kinetics of polyplexes by a model constructed from quantum dot-FRET. Mol. Ther. 16: 324– 332 (2008). 60. Melchior, F. and Gerace, L. Mechanisms of nuclear protein import. Curr. Opin. Cell Biol. 7: 310–318 (1995). 61. Huth, S., et al. Interaction of polyamine gene vectors with RNA leads to the dissociation of plasmid DNA-carrier complexes. J. Gene Med. 8: 1416–1424 (2006). 62. Iida, T., et al. Overall interaction of cytosolic proteins with the PEI/DNA complex. J. Control. Release 118: 364–369 (2007). 63. Wolff, J.A. and Rozema, D.B. Breaking the bonds: non-viral vectors become chemically dynamic. Mol. Ther. 16: 8–15 (2008). 64. Zabner, J., et al. Cellular and molecular barriers to gene transfer by a cationic lipid. J. Biol Chem. 270: 18997–19007 (1995). 65. Schaffer, D. V., et al. Vector unpacking as a potential barrier for receptormediated polyplex gene delivery. Biotechnol. Bioeng. 67: 598–606 (2000). 66. Itaka, K., et al. In situ single cell observation by fluorescence resonance energy transfer reveals fast intra-cytoplasmic delivery and easy release of plasmid DNA complexed with linear polyethylenimine. J. Gene Med. 6: 76–84 (2004). 67. Matsumoto, Y., et al. Intranuclear fluorescence resonance energy transfer analysis of plasmid DNA decondensation from nonviral gene carriers. J. Gene Med. 11: 615–623 (2009). 68. Knorr, V., et al. Acetal linked oligoethylenimines for use as pH-sensitive gene carriers. Bioconjug. Chem. 19: 1625–1634 (2008). 69. Knorr, V., Ogris, M., and Wagner, E. An acid sensitive ketal-based polyethylene glycol-oligoethylenimine copolymer mediates improved transfection efficiency at reduced toxicity. Pharm. Res. 25: 2937–2945 (2008). 70. Zhong, Z., et al. A versatile family of degradable non-viral gene carriers based on hyperbranched poly(ester amine)s. J. Control. Release 109: 317–329 (2005). 71. Forrest, M. L., Koerber, J. T., and Pack, D. W. A degradable polyethylenimine derivative with low toxicity for highly efficient gene delivery. Bioconjug. Chem. 14: 934–940 (2003). 72. Akinc, A., et al. Synthesis of poly(beta-amino ester)s optimized for highly effective gene delivery. Bioconjug. Chem. 14: 979–988 (2003). 73. Wong, S. Y., Pelet, J. M., and Putnam, D. Polymer systems for gene delivery—past, present, future. Prog. Polym. Sci. 32: 799–837 (2007). 74. Russ, V., et al. Novel degradable oligoethylenimine acrylate ester-based pseudodendrimers for in vitro and in vivo gene transfer. Gene Ther. 15: 18–29 (2008). 75. Russ, V., et al. Oligoethylenimine-grafted polypropylenimine dendrimers as degradable and biocompatible synthetic vectors for gene delivery. J. Control. Release 132: 131–140 (2008). 76. Wang, Y., Chen, P., and Shen, J. The development and characterization of a glutathione-sensitive cross-linked polyethylenimine gene vector. Biomaterials 27: 5292–5298 (2006).

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CHAPTER 8

Intracellular Trafficking of Membrane Receptor-Mediated Uptake of Carbon Nanotubes BIN KANG and YAODONG DAI Nanjing University of Aeronautics and Astronautics, Nanjing, Peoples Republic of China

8.1

INTRODUCTION

Over the last decade, newly synthesized nanomaterials (NMs), exhibiting distinctive optical, magnetic, and conducting properties, have been found to exhibit fascinating potential applications in biomedical fields. Their small size, increased surface area, and reactivity allow them to easily penetrate cell membranes, efficiently bind molecular species, and catalyze chemical reactions [1–3]. Due to their enhanced properties, NMs have been proposed for use in the fabrication of various devices such as catalysts, sensors, composite fillers, actuators, transistors, drug and gene deliverers, biosensors, virus inhibitors, and protein immobilizers [4–13]. Among such nanomaterials, carbon nanotubes (CNTs) are among the most promising materials for such applications because CNTs have better biocompatibility and easy cell internalization [14– 18]. Also, CNTs can easily be functionalized and linked to fluorescent beacons, nucleic acids, or other small molecules, which make CNTs an ideal candidate for a nanoparticle-based system for drug delivery and bioimaging [19–22]. The cellular uptake of CNTs is a very fundamental issue that needs to be understood before using CNTs in various biological applications. Although a lot of research has been done on the interaction of CNTs and living cells, current published data on cell penetration mechanisms and the intracellular dynamic are still particularly inconsistent and widely disputed. Some studies suggest that CNTs can pass directly through the cell membrane via an energyindependent nonendocytotic mechanism [23, 24]: that is, the CNTs may cause Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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localized damage on the cell membrane through which cargos enter the cells. The one-dimensional structure of CNTs has been shown to enhance this “holepunching” process and improve the efficiency of delivery into mammalian cells [24]. On the other hand, other groups propose that cells take up the CNTs through an energy-requiring endocytosis process [8]. The underlying reasons for the discrepancies might be attributed to two causes: first, the wide variability in sample preparation and “purification” methods, including incomplete characterization of the CNT materials following purification, such as minimal description about their postpurification solution behavior; and second, the use of nonuniform characterization methods and materials with different preparative protocols, viability assessment methods, and cell populations. For CNTs, the different physical characteristics, such as tube diameter, length, surface chemistry, and biological coatings, may result in various interactions in the biological context. Definitive discrimination of relative and synergistic effects with respect to these differences will continue to be impossible without implementation of precise measurements, complete characterization, and the use of well-defined materials. Another unaddressed question is the intracellular distribution of internalized CNTs after penetrating the cytoplasm membrane and further fate. Previous reports focused mainly on the uptake of the CNTs and little work has been done with respect to the behaviors of CNTs after their penetration into cells. For example, how long can the CNTs stay in the cells? Can CNTs be released from the cells through exocytosis? Can CNTs pass through the cellular nuclear envelope? All these questions are still being debated. While most studies have shown that CNTs can penetrate through the cell membrane and be internalized into the cell cytoplasm, few studies show conclusive evidence that CNTs can or cannot get into the cell nucleus, even through some studies have given indirect evidence, such as a G-band Raman signal in the nuclear region or successful use for plasmid DNA delivery. Single-particle tracking (SPT) using fluorophores is a recently developed technique for answering the above questions in cellular systems, although with small fluorophores and QDs, photobleaching is a major limitation to real-time measurements. The photobleaching time constrains the observation window during tracking so that events occurring on the order of several hours must be observed through multiple and distinct incubation periods, with each observation starting at a different time after incubation. However, SPT is considered a promising tool to study the uptake of nanoparticles; in particular, special SPT methods using the NIR fluorescence of CNTs show potential to achieve a better understanding of the cellular uptake dynamics of nanotubes. Therefore the focus of this chapter is to summarize briefly the important results of cellular uptake and tracking of CNTs conducted to date, with emphasis on the uptake pathway and the subcelluar distribution as well as the recent status on uptake dynamics by using single-particle tracking. A description of the current in vitro analysis techniques and materials characterization

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methods is given within the context of the ongoing challenges, and recommendations are made for achieving more reliable measures of CTN–living cell interactions.

8.2

RELATION OF NANOTUBE SIZE AND FUNCTIONALIZATION

There is broad agreement that biological studies should put more emphasis on detailed characterization of test nanomaterials. For cellular uptake of CNTs, one of the main sources of the variation in the published data is the wide variability in CNT dispersion from different preparative protocols and methods. The differences in dispersion ranging from clusters, bundles of multiple nanotubes, or individually dispersed nanotubes will dramatically affect the absolute size and amount of surface area of the nanotube material to which the cells are exposed. Dispersion protocols involving surfactants are attractive, as the incorporation of the nanotubes inside a surfactant shell does not appreciably alter the graphitic structure and desirable physicochemical properties of the single-walled carbon nanotubes (SWCNTs) [25]. Recently, DNA, peptides, and carbohydrates have been used in this surfactant/ wrapping polymer role and have been demonstrated to suspend SWCNTs, with high individual dispersion of the nanotubes [26, 27]. These dispersions, in the case of DNA, are even stable enough to allow for separation of the dispersed material into well-defined subpopulations of the SWCNTs. Another point is the physical properties of the materials used for CNTs, especially the length. Recent reports have highlighted successes in separating polydisperse SWCNT populations into well-defined length and chirality fractions using gel chromatography [25, 28–30], size exclusion chromatography (SEC) [31–35], ion chromatography (IC) [36, 37], or various forms of electrophoresis [28, 38–40]. This sorting affords opportunities to explore the uptake and fate of CNTs with different lengths. Some typical work has been done by Becker et al. [41]. They used well-dispersed, separated-length CNT fractions by SEC, and gave an exhaustive characterization of these fraction populations. Their results indicated a threshold on the length and corresponding toxicity of SWCNTs that are taken up into cells. They found that the longer tubes are excluded from the cell interior while the shorter SWCNTs are able to access the cytosol. Through the length-selective uptake of nanotubes by the cell populations, they determined an approximate uptake threshold of approximately 189 ± 17 nm, indicating that nanotubes shorter than this length are consumed and likely induce more toxicity. Also, the size threshold concept has been suggested for multiwalled carbon nanotubes, which influence cell uptake, retention, and biodistribution in other nanoparticle systems [42–44]. Another fundamental question is whether CNTs with different functionalized groups have the same behavior and are capable of cell binding, uptake, and internalization. Note that the functionalization here only means general chemical groups, in the absence of any coating or conjugation with biological

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O

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(FITC) H N O

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Figure 8.1. Molecular structures of CNT covalently functionalized with different types of small molecules: 1, ammonium-functionalized CNT; 2, acetamido-functionalized CNT; 3, CNT functionalized with fluorescein isothiocyanate (FITC); 4, CNT bifunctionalized with ammonium groups and FITC; 5, CNT bifunctionalized with methotrexate (MTX) and FITC; 6, shortened CNT bifunctionalized with amphotericin B (AmB) and FITC; 7, shortened CNT bifunctionalized with ammonium groups and FITC (through an amide linkage). (Copyright Nature Publish Group. Reproduced with permission from Ref. 45.)

macromolecules such as antibodies and others. This question highlights the systemic investigation of the behavior of different functionalized CNTs (fCNTs). Kostarelos and co-workers [45] did excellent work showing the effect of different functionalized groups on the interaction with cells. They carried out a series of experiments using functionalized single-walled (f-SWCNTs) and multiwalled (f-MWCNTs) CNTs with a wide variety of functional groups (Figure 8.1). Intrinsically luminescent or fluorescently labeled f-CNTs were directly tracked and imaged intracellularly by epifluorescence and confocal laser scanning microscopy (CLSM). The imaging of f-CNTs and not their adsorbed macromolecules, as previously reported by other groups [11, 46], is imperative in order to elucidate their interaction with cellular compartments.

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Even in studies tracking the intracellular localization of nonfunctionalized pristine CNTs (pCNTs) solubilized by polymer or single-stranded DNA molecules by the IR spectral characteristics of the pCNT backbone [47–49], one should be very cautious when extrapolating conclusions about pCNT–cellular compartment interactions. Indeed, the effect of the macromolecules acting as their solubilizing agent can play a critical role in determining the type of ensuing interactions with cells and the mechanisms of cellular uptake, as has been shown for other polymer-coated nanostructures [50]. The functionalization of SWCNTs and MWCNTs in the Kostarelos et al. [45] study was mainly performed using the 1,3-dipolar cycloaddition of azomethine ylides [51–53]. This approach allows insertion of amino functions around the side walls and at the tips of the CNTs, which renders the tubes highly soluble in aqueous environments. The amino groups were further modified by covalently linking a variety of small molecules including fluorescent probes and anticancer and antibiotic agents [18, 54]. The addition of functional groups was carried out in a modular fashion, gradually increasing the molecular complexity of the groups covalently linked onto the nanotube side walls (Figure 8.1). In agreement with other studies that used differently functionalized CNTs, f-CNTs 1 and 2 were found to be intrinsically luminescent in the ultraviolet/visible region [55, 56]. Meanwhile, f-CNTs 3–7 were conjugated with fluorescein isothiocyanate (FITC) to obtain high levels of fluorescence signal. The interaction between f-CNTs and a wide variety of live cells was studied. All f-CNTs were allowed to interact with different cell types. It could be observed that the nature of the functional group on the CNT surface did not determine whether f-CNTs were internalized or not. Even in cases where the functional groups were electrostatically neutral or negatively charged in physiological conditions, nanotubes were consistently taken up by cells. The intracellular trafficking of individual or small bundles of f-CNTs occurred, and the transportation of nanotubes toward the perinuclear region was observed a few hours following initial contact with the cells, even under endocytosis-inhibiting conditions. Other mechanisms (such as phagocytosis)—depending on cell type, size of nanotube, and extent of bundling—may also be contributing to or be triggered by the ability of f-CNTs to penetrate the plasma membrane, and therefore be directly involved in the intracellular trafficking of the f-CNTs. From the systemic research of cellular uptake of CNTs with different kinds of groups, they draw a conclusion that the cellular uptake of functionalized carbon nanotubes is independent of functional group.

8.3

UPTAKE PATHWAY AND MECHANISM

Generally, eukaryotic cells internalize extracellular materials inside the cytoplasm through two uptake pathways: an energy-independent pathway and an energy-dependent pathway. Low molecular weight solutes directly transport through the plasma membrane via the energy-independent pathway without

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energy consumption. The energy-dependent pathway, also known as endocytosis for uptake of relatively large particles, can be hindered when incubations are taken at 4 °C or in ATP (adenosine triphosphate)-free medium. The ATP production in cells is disturbed with a well-known reagent, NaN3. The endocytosis is divided into at least two subcategories, that is, clathrin-dependent and clathrin-independent endocytosis. Extracellular materials or solutes that were taken into cells through the clathrin-dependent pathway were first enclosed within clathrin-coated vesicles derived from folds or invaginations of the plasma membrane, and then were brought inside the cells. On the other hand, clatherin-independent endocytosis can occur through the caveolae or lipid-raft pathway. The two subcategories can be separated by hindering one of them. Such treatment consisted of pretreating the cells with sucrose to disturb the formation of clathrin-coated vesicles on the cell membrane, thereby hindering clathrin-dependent endocytosis. As the caveolae-dependent or lipid-rafts-dependent cell internalization relies on the presence of cholesterol domain, the chemical filipin III can be used to disturb the cholesterol distribution within the cell membrane, thus hindering the formation of caveolae. For CNTs, detailed work to establish the cellular uptake mechanism and pathway is still lacking. Pantarotto et al. [16] have suggested an energy-independent nonendocytotic mechanism that involves insertion and diffusion of nanotubes through the lipid bilayer of the cell membrane. However, the Dai group [8] proposed a different mechanism, that is, energy-dependent endocytosis progress. They have systematically investigated the cellular uptake mechanism and pathway for carbon nanotubes. They present evidence showing that clathrindependent endocytosis is the pathway for the uptake of various SWCNT conjugates with proteins and DNA. They also discuss the differences between the nanotube materials and the experimental procedures used in their work and by Pantarotto et al. [16], who suggested an energy-independent nonendocytotic uptake of nanotubes. Their work seems to clearly establish the intracellular uptake mechanism of SWCNTs in the form of individual and small bundles with lengths of <1 μm and is very helpful to avoid any confusion and controversy over the cellular uptake mechanism for these materials. Next, they carried out a systematic investigation of the cellular internalization mechanism and pathway for SWCNT conjugates. Cellular incubations were carried out at 4 °C and with cells pretreated with NaN3 in parallel with our regular incubation conditions. They found that the fluorescence levels observed with confocal microscopy from cells, after incubation in SWCNT conjugates at 4 °C (Figure 8.2B) and ATP depletion by NaN3 (Figure 8.2C), were low. This therefore indicates endocytosis as the internalization mechanism for the uptake of SWCNT conjugates at 37 °C. This was further confirmed by flow cytometry measurements, which indicated a significant reduction in cellular uptake at 4 °C and under the azide ATP depletion conditions (Figure 8.2D). To assess the role of clathrin in the internalization of SWCNTs, they carried out incubations under conditions that are known to disrupt the formation of clathrin-coated vesicles on the cell membrane. Their

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Figure 8.2. Confocal microscopy images of HeLa cells after incubation in fluorescently labeled DNA–SWCNT at (A) 37 °C, (B) 4 °C, and (C) after pretreatment with NaN3. (D) Flow cell cytometry data for HL-60 cells that were incubated in fluorescently labeled pure DNA or protein solutions without nanotubes (labeled “DNA” and “protein,” respectively), DNA–SWCNT and BSA–SWCNT at 37 °C, and for protein– SWCNT at 37 °C in cells incubated at 4 °C or pretreated with sodium azide. (E) Flow cell cytometry data obtained after incubation in protein–SWCNT solutions for untreated cells, cells pretreated with 0.45 M sucrose, and K+-depleted medium, respectively. (F) A confocal image that shows cellular uptake of a rhodamine-labeled transferring protein in HL-60 cells at 37 °C. The inset shows the lack of uptake of the transferrin protein after pretreatment of cells in sucrose. (G) Flow cytometry data of cells after incubation in choleratoxin B (black bars) and BSA-SWCNT (gray bars) for HeLa cells without any pretreatment (control) and cells pretreated with filipin and nystatin, respectively. (Copyright Wiley-VCH. Reproduced with permission from Ref. 8.)

experimental data suggest that cellular internalization of SWCNT conjugates with proteins and DNA is through the clathrin-dependent endocytosis pathway (Figure 8.2). Receptor-mediated endocytosis through clathrin-coated pits is the most common pathway of endocytosis. It provides a means for the selective and efficient uptake of macromolecules and particles that may be present at relatively low concentrations in the extracellular medium. Cells have receptors for the uptake of many different types of ligands, including hormones, growth factors, enzymes, and plasma proteins. For active targeting to cancer cells, nanomaterials were functionalized with ligands such as antibodies, peptides, nucleic acid aptamers, carbohydrates, and small molecules. Among these, folic acid (FA) is appealing as a small molecular ligand for targeting cell membrane and allowing nanomaterial endocytosis via a folate–folate receptor-mediated

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Figure 8.3. Multichannel confocal images of Hep G2 cells after incubation with FASWCNTs for 1 h (left, first column), 3 h (left, second column), and 5 h (left third column). The green fluorescence from the outside to the inside of cells shows the uptake pathway of FA-SWCNTs into cancer cells. Right-hand column: multichannel confocal images of Hep G2 cells incubated in FA-SWCNTs for 3 h with pretreatment of free FA blocking. The concentrations of FA-SWCNTs in all groups are 20 μg/mL. (Copyright IOP. Reproduced with permission from Ref. 57.) (See color insert.)

pathway for higher uptake yields. Also, the high affinity of folate to bind its receptor allows its use for specific cell targeting. A typical result of cellular uptake of folate conjugated CNTs is shown in Figure 8.3 [57]. After incubation in FA-SWCNTs for 1 hour, the SWCNTs initially bonded on the plasma membrane of the cells. After 3 hours, stronger fluorescence was observed in the cytoplasm, indicating that SWCNTs entered into the cells. For 5 hours, confocal images revealed decreased and spotted fluorescence inside cells, corresponding to redistribution and discharge of SWCNTs out of the cells. To further elucidate this pathway, we blocked the folate receptor on the surface of Hep G2 cell membrane with free folate acid before incubation with FA conjugated nanotubes. The uptake of FA-SWCNTs was blocked, as evidenced by the small amount of fluorescence in the cells with the pretreatment of free folate acid. These results demonstrated that Hep G2 cells were able to internalize folate-conjugated SWCNTs via the folate receptor-mediated pathway.

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SUBCELLULAR DISTRIBUTION

After cell penetration, the intracellular fate of the internalized CNTs appears to vary. The subcellular distribution of SWCNTs inside Hep G2 cells have shown that the FA-SWCNTs are located in the cytoplasm but not in the nuclei, indicating that the nanotubes under study cannot transport through the nuclear envelope. It is well known that the nuclear pore on the nuclear envelope allows the mRNA and tRNA to move across the nuclear envelope, and the diameter of the pore is estimated to be about 20 nm in various papers. After functionalization of SWCNTs, the diameter of the nanotubes might be larger than that, or the nanotubes might be too long to pass the nuclear pore, resulting in failure to transport inside the nuclei. The lack of nanotubes in nuclei indicates that studying the subcellular distribution of SWCNTs is very important for understanding the uptake and toxicity of carbon nanotubes. In particular, since SWCNTs are currently under consideration as transporters of DNA and RNA for gene therapy, the presence or absence of carbon nanotubes in nuclei is a vital question that needs to be addressed before the practice of gene therapy. Although most related studies have reported that water-soluble CNTs modified with functional groups resided in the cytoplasm upon incubation, nuclear accumulation was observed. One study has reported that an intense G-band signal was observed in the nuclear region in exposed HeLa cells using confocal Raman imaging analysis; however, the nuclear accumulation of CNTs was not inferred because of the lack of conclusive evidence of CNT-like structures in the nucleus through ultrasections analysis under transmission electron microscopy. The nuclear accumulation of CNTs has been demonstrated by Bianco and co-workers [58]; furthermore, they reported that peptide functionalized multiwalled CNTs could cross the nuclear membrane and be successfully used for plasmid DNA delivery [16]. A better understanding of the intracellular properties of CNTs is therefore essential for the development of this nanomaterial as a drug delivery vector. Recent work by Cheng et al. [59] revealed a new insight into the nuclear accumulation of SWCNTs conjugated with polyethylene glycol (PEG) and labeled with fluorescent dye FITC (FITC-PEG-SWCNTs) in several mammalian cell lines. Cellular uptake and release of FITC-PEG-SWCNTs were observed using time-lapse fluorescence microscopy. This work extends our current understanding of the interactions between CNTs and biological systems, such as the dynamic fate, intracellular behavior, and the uptake and release mechanism. They investigated the cell-penetrating property of PEG-SWCNTs by incubating human cervical carcinoma HeLa cells with FITC-PEG-SWCNTs for 48 h. After extensive washing, the treated cells were analyzed by flow cytometry. As shown in Figure 8.4A, FITC fluorescence was detected in virtually all treated cells, suggesting that a substantial amount of FITC-PEG-SWCNTs had become associated with HeLa cells. Free FITC did not associate with

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Figure 8.4. The cell penetration of FITC-PEG-SWCNTs and its nuclear accumulation in mammalian cells. (A) FACS analysis of HeLa cells treated with FITC-PEG-SWCNTs for 48 h showed that the treated cells took up the FITC-PEG-SWCNTs uniformly. (B) Live cell imaging showed that FITC-PEG-SWCNTs accumulated in the nucleus, mainly in the nucleolus (arrows), of HeLa cells. (C) DIC image of panel B. (D–G) U2OS cells (D, E) and MEF (F, G) were incubated with FITC-PEG-SWCNTs and then organeller markers. Signals from PEGylated SWCNTs (green) did not coincide with mitochondria (red, D–G) but did with nuclei (blue, E, G). Note that the distributions of FITC-PEG-SWCNTs between the nucleolus and the nucleoplasm are different in these three cell lines. Scale bars: (B, C) 20 μm; (D–G) 10 μm. (Copyright ACS. Reproduced with permission from Ref. 59.) (See color insert.)

HeLa cells. The cellular association of FITC-PEG-SWCNTs was time and dose dependent, with cellular green fluorescence detectable as soon as after 1 h of incubation. The detection of cellular FITC by flow cytometry suggested that PEGylated SWCNTs could either enter the cells or adhere to the cell surface. The intracellular distribution of FITC-PEG-SWCNTs in a number of mammalian cell lines was revealed by live cell fluorescence microscopy. Figure 8.4 panels B and C, show HeLa cells incubated with FITC-PEG-SWCNTs. The green fluorescence signal was predominately in the cell nucleus with a weaker cytoplasmic staining in some cells. In most cells, FITC-PEG-SWCNTs were observed to be enriched in a few intranuclear structures. Bright-field DIC images indicated that these intranuclear structures were nucleoli (Figure 8.4B,C, arrows). To further characterize the subcellular distribution of FITCPEG-SWCNTs, CNT-treated cells were counterstained by a mixture of cellpermeable dyes, Hoechst 33342 and MitoTracker, to label the nucleus and mitochrondria, respectively. The dyes were used at concentrations that did not cause detectable cell toxicity and did not interfere with normal mitosis for at least 24 h [16, 17]. FITC-PEG-SWCNT was observed to coincide with Hoechst

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33342 (Figure 8.4E,G), indicating the internalized SWCNT was predominately nuclear. This confirms that the green fluorescence detected by flow cytometry represented intracellular SWCNTs and supports previous observations that modified SWCNTs can effectively cross cell membranes. While the nuclear targeting mechanism of CNTs is not known, it is possible that the internalized PEGylated SWCNTs are retained in the nucleus through binding with specific nuclear proteins or nucleic acids. Future investigations will include the study of the effect of the molecular weight and surface coverage of PEG on the cellular uptake, intracellular distribution, and dynamic fate of CNTs. It seems PEGylated CNTs are one of the few nanomaterials that accumulate in mammalian cell nuclei without the need of cell-penetrating and nuclearlocalizing tags. Several types of nanoparticles can also enter cells without any conjugation, but these nanomaterials are trapped in the endosome [60]. Many studies reported a predominantly cytoplasmic localization of CNTs [46, 61] instead of a nuclear accumulation. The intracellular location of different kinds of CNTs has not been systematically studied and the cause of this discrepancy is not clear.

8.5

SINGLE-PARTICLE TRACKING

Although much effort has been expended to understand the pathway and mechanism of cellular uptake of nanoparticles, some central questions are still unclear: How is this process carried out? What are the dynamics? How long can the particles stay inside the cells? Does exocytosis occur for nanoparticles? A discussion on these issue is nearly absent from the literature. In recently published works on Au nanoparticles, exocytosis was observed after the extracellular nanoparticle concentration gradient was removed. Studying exocytosis involves incubations where separate specimens are prepared at various time points after incubation, largely to avoid photobleaching [62–66]. There are several limitations on this approach, particularly in capturing nanoparticle dynamics and precisely quantifying endocytosis or exocytosis rates simultaneously. Single-particle tracking (SPT) using fluorophores is a recently developed technique for answering the above questions in cellular systems [67, 68], although with small fluorophores and QDs, photobleaching is a major limitation on real-time measurements. The photobleaching time constrains the observation window during tracking so that events occurring on the order of several hours must be observed through multiple and distinct incubation periods, with each observation starting at a different time after incubation. Strano’s group developed a new technique for SPT of carbon nanotubes, that is, use of the NIR fluorescence signal of SWCNTs [69]. By using this method, they tracked over 10,000 individual trajectories of nonphotobleaching SWCNTs as they are incorporated into and expelled from NIH-3T3 cells in real time on a perfusion microscope stage. Also, an analysis of mean square

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displacement allows the complete construction of the mechanistic steps involved from single duration experiments. They observed the first conclusive evidence of SWCNT exocytosis and showed that the rate closely matches the endocytosis rate with negligible temporal offset. They identified and studied the endocytosis and exocytosis pathway that leads to the previously observed aggregation and accumulation of SWCNTs within the cells. By using image processing algorithms [70, 71], over 10,000 SWCNT trajectories have been tracked from the sequence of images from both data sets. A snapshot of the detected trajectories from ImageJ is shown in Figure 8.5A. Direct proof of endocytosis (Figure 8.5B–I) and exocytosis (Figure 8.5J–M) is repeatedly observed from the NIR image sequences, represented as independent trajectories, confirmed by the MSD features of endocytosis and exocytosis. For instance, MSD of endocytosis has the signature presented later in this chapter. By using a halogen illuminator (100 W), they could observe

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Figure 8.5. (A) A subset of observed trajectories extracted using ParticleTracker. (B–I) Endocytosis (confirmed by MSD feature) of a single particle (arrow) is observed between t = 1029 and 1112 s, identified as RME (see text). (J–M) Evidence of exocytosis (confirmed by MSD feature) of a single particle (arrow) observed from t = 2218 to 2250 s. (N–Q) Aggregation and movement inside the cell. The cell is illuminated by the halogen lamp and DNA-SWCNTs is illuminated by laser at 785 nm. (Copyright ACS. Reproduced with permission from Ref. 69.)

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the scattering of the cell periphery and the PL signal from DNA-SWCNTs simultaneously, which allows for unambiguous classification. An example image sequence of the movement of internalized particles is shown in Figure 8.5N–Q (the dark spot in the middle is from the shade of the light filter). This technique can also verify the overlap between phase contrast cell images and measured trajectories. In order to explain endocytosis and exocytosis mathematically, recurring events of temporal and spatial coordinates of each particle are then analyzed using an algorithm we have written in Matlab to classify the trajectories using their starting and ending locations relative to the cell, as well as the functional form of its MSD. Briefly, the coordinates of the particle on the stage relative to the location of the membrane, cell interior, and the flow direction along the stage, together with the functional form of its MSD, assist in classifying the trajectories according to a number of commonly observed steps within the complete pathway. While 49.2% of trajectories were purely convective diffusion in the flow field with no cellular interaction, the remaining 50.8% demonstrated membrane surface adsorption (6.2%), surface diffusion (18.4%), endocytosis (12.7%), exocytosis (5.9%), or desorption (7.4%).

8.6

CONCLUSION

Increased efforts should be made on the basic characterization of well-defined CNTs, including the diameter, length, surface chemistry, dispersity, and functionalization procedure. It is necessary to develop a standardized process for the characterization and application procedure of CNTs, even though there is a long, long way to go. Also, we emphasize the need for research on the fate of CNTs inside live cells, that is, “what will happen when CNTs are inside cells” not only just “if CNTs can get into cells or not.” New techniques like single-particle tracking should be continuously developed and improved to provide appropriate tools for studying the dynamics of cell–CNT interaction in real time. Only if these central issues are addressed can we have a good understanding of the cellular effect of CNTs; thus application of CNTs in biomedical fields such as DNA transporter and drug delivery will be more reasonable and safer.

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57. Kang, B., et al. Intracellular uptake, trafficking and subcellular distribution of folate conjugated single walled carbon nanotubes within living cells. Nanotechnology 19(37): 375103–375111 (2008). 58. Pantarotto, D., Briand, J. P., Prato, M., and Bianco, A. Translocation of bioactive peptides across cell membranes by carbon nanotubes. Chem. Commun. 1: 16–17 (2004). 59. Cheng, J. P., et al. Reversible accumulation of PEGylated single-walled carbon nanotubes in the mammalian nucleus. ACS Nano 2(10): 2085–2094 (2008). 60. Duan, H. W. and Nie, S. M. Cell-penetrating quantum dots based on multivalent and endosome-disrupting surface coatings. J. Am. Chem. Soc. 129(11): 3333–3338 (2007). 61. Monteiro-Riviere, N. A., Nemanich, R. J., Inman, A. O., Wang, Y. Y. Y., and Riviere, J. E. Multi-walled carbon nanotube interactions with human epidermal keratinocytes. Toxicol. Lett. 155(3): 377–384 (2005). 62. Chithrani, B. D. and Chan, W. C. W. Elucidating the mechanism of cellular uptake and removal of protein-coated gold nanoparticles of different sizes and shapes. Nano Lett. 7(6): 1542–1550 (2007). 63. Panyam, J., et al. Polymer degradation and in vitro release of a model protein from poly(d,l-lactide-co-glycolide) nano- and microparticles. J. Control. Release 92(1-2): 173–187 (2003). 64. Panyam, J., Sahoo, S. K., Prabha, S., Bargar, T., and Labhasetwar, V. Fluorescence and electron microscopy probes for cellular and tissue uptake of poly(d,l-lactideco-glycolide) nanoparticles. Int. J. Pharm. 262(1-2): 1–11 (2003). 65. Panyam, J. and Labhasetwar, V. Biodegradable nanoparticles for drug and gene delivery to cells and tissue. Adv. Drug Deliv. Rev. 55(3): 329–347 (2003). 66. Panyam, J. and Labhasetwar, V. Dynamics of endocytosis and exocytosis of poly(d,l-lactide-co-glycolide) nanoparticles in vascular smooth muscle cells. Pharm. Res. 20(2), 212–220 (2003). 67. Jin, S., Haggie, P. M., and Verkman, A. S. Single-particle tracking of membrane protein diffusion in a potential: simulation, detection, and application to confined diffusion of CFTR Cl-channels. Biophys. J. 93(3): 1079–1088 (2007). 68. Jin, S. and Verkman, A. S. Single particle tracking of complex diffusion in membranes: simulation and detection of barrier, raft, and interaction phenomena. J. Phys. Chem. B 111(14): 3625–3632 (2007). 69. Jin, H., Heller, D. A., and Strano, M. S. Single-particle tracking of endocytosis and exocytosis of single-walled carbon nanotubes in NIH-3T3 cells. Nano Lett. 8(6): 1577–1585 (2008). 70. Sbalzarini, I. F., Mezzacasa, A., Helenius, A., and Koumoutsakos, P. Effects of organelle shape on fluorescence recovery after photobleaching. Biophys. J. 89(3): 1482–1492 (2005). 71. Sbalzarini, I. F. and Koumoutsakos, P. Feature point tracking and trajectory analysis for video imaging in cell biology. J. Struct. Biol. 151(2): 182–195 (2005).

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CHAPTER 9

Real-Time Particle Tracking for Studying Intracellular Transport of Nanotherapeutics CLIVE CHEN and JUNGHAE SUH Department of Bioengineering, Rice University, Houston, TX

9.1

INTRODUCTION

The use of nanotherapeutic agents, such as drug and gene carriers, has offered distinct advantages, such as targeted delivery and controlled release, over systemic infusion of therapeutic drugs [1]. The efficacy of these nanotherapeutics is often affected by their ability to reach target organelles upon cellular entry. Intracellular transport of nano-sized carriers can be a difficult process as the cytoplasm, packed with a dense network of cytoskeletal filaments and organelles, is a highly complex biological medium to traverse [2, 3]. There is great interest in understanding the intracellular transport mechanisms involved in the trafficking of nanotherapeutics so that current designs can be improved to facilitate the delivery of cargo drugs and genes to their sites of action [4, 5]. Through advances in fluorescence microscopy, the movement of fluorescently labeled components can be monitored in real time in live cells. Various techniques have been developed to quantify the mobility of labeled molecules and among these, single-particle tracking (SPT) has emerged as a particularly powerful tool for measuring the transport properties of individual transporting entities [6]. In SPT, the positions of tens to hundreds of particles are monitored simultaneously over time and subsequent image analysis provides information on the transport properties for each individual particle. In contrast to other widely used mobility measurement techniques, such as fluorescence recovery after photobleaching (FRAP) and fluorescence correlation Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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spectroscopy (FCS), SPT not only can be used to measure the ensembleaveraged transport properties of a large number of particles, but it can also quantitatively delineate the mobility of individual particles. The ability to obtain detailed quantitative information on the transport properties of individual particles adds greatly to our understanding of the complex intracellular trafficking faced by our engineered nanotherapeutics. In this chapter, we will first discuss the two major stages involved in SPT: locating and tracking of individual particles over time, and analysis of tracking data. Since the majority of SPT studies on nanotherapeutics have been performed in two dimensions, our discussion will be limited to two-dimensional (2D) particle tracking. Readers are referred to a review elsewhere [6] for a discussion on how calculations involved in 2D particle tracking can be translated to three dimensions. In the second part of the chapter, we will review and discuss studies conducted using SPT to investigate the intracellular transport of a variety of nanoscale platforms, including polymeric nanoparticles, quantum dots, and viruses.

9.2

LOCATING AND TRACKING OF INDIVIDUAL PARTICLES

In SPT, the motions of particles are recorded as movies, which are a series of microscopy images taken at a fixed time interval. The trajectories of individual particles are then determined by tracing their changes in position over time. Spatial resolution, or the minimum distance required between two particles in order to resolve them as being separate, is the well-known diffraction limited resolution of about 200 nm. Tracking resolution, or the minimum displacement that can be measured with SPT, can be much smaller [7]. Tracking algorithms are used to locate individual particles precisely and to monitor their trajectories in a high-throughput manner. While many tracking algorithms have been developed over the years, they generally revolve around two basic approaches. The first approach focuses on approximating the absolute positions of particles within individual images while the second approach is correlation-based and requires the comparison of signal intensity patterns between successive frames of images [8, 9]. The absolute position of a particle can be estimated by the center of its point spread function (PSF) when imaged with a microscope. The PSF of the particle can in turn be approximated with a Gaussian function [10, 11]. The central peak of the Gaussian fit is taken to be the coordinates of the particle. Alternatively, a center of mass, or centroid, is identified by weighing the signal intensity within a region of interest selected from a microscopic image [7, 12], and its location is taken to be the position of a particle. The centroid of a spherical particle of uniform fluorescence intensity will also be the center of its PSF. After a particle has been located in every frame of the recorded movie, its change in position between successive frames is traced to determine the trajectory of the particle.

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In the correlation-based approach, images of a particle between consecutive frames are assumed to have the most similar intensity patterns [13, 14]. An image frame with an intensity pattern that represents a particle is selected and overlaid on top of its subsequent frame to begin comparison of the patterns of interest. The selected frame is shifted at one-pixel increments and the similarity between the patterns is calculated using a correlation matrix. The location with the best match in the intensity pattern can be determined by finding which cell in the correlation matrix has the maximum value. The displacement of the particle is then calculated from the number of pixels being shifted to best align its images from two consecutive frames. Each tracking algorithm has its own set of advantages and disadvantages. For example, while centroid-based algorithms are computationally efficient, they are susceptible to background noise [13]. On the other hand, correlationbased methods are not as affected by noise but can be prone to erroneous matching of particles [9, 13]. More recent algorithms often combine elements from both approaches to exploit their respective strengths. Choice of tracking algorithm usually depends on the microscope, signal detector (camera or photomultiplier tube), and fluorescent probe being tracked since the resulting signal-to-noise ratio is one of the most important factors that needs to be considered. Figure 9.1 shows how trajectories of individual nonviral gene delivery vectors would appear in an intracellular setting when monitored through SPT. Readers are referred to reviews elsewhere for a more comprehensive discussion and comparison of commonly used tracking algorithms [8, 9, 13, 15].

9.3

ANALYSIS OF TRACKING DATA

The first transport data that is calculated in a particle tracking study is the mean squared displacement (MSD) of the tracked particles. In 2D particle tracking, the MSD of tracked particles can be calculated as follows: MSD( t ) = ( x( t ) − x ( t + Δτ ) + ( y( t ) − y( t + Δτ )) 2

2

(9.1)

The MSD can then be used to find the apparent rates of transport, or effective diffusivities (Deff), of individual particles: Deff = MSD ( t ) 4t

(9.2)

It is worth noting that the MSD described here is not a function of time, but that of time interval, t (also known as time scale). As a tracked particle follows a particular trajectory within a predetermined period of imaging time, multiple MSD values can be calculated from the same particle trajectory across different t, with t being dependent on the frame rate at which the particle is

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Figure 9.1. Trajectories of nonviral gene vectors in a live cell tracked using SPT. The observed trajectories are highly heterogeneous, which suggests that different transport mechanisms may be involved. (Reproduced from Ref. 53 with permission.) (See color insert.)

monitored. Readers are referred to a review by Suh et al. [6] for a more detailed discussion of time scale and MSD calculation. While MSD itself mathematically describes the motion of the tracked particles, the changes of MSD, or Deff, across different time scales are indicative of the different transport modes, such as diffusive, subdiffusive, and active transport, which describe the motion of the particles. Various theoretical equations that relate MSD with t have been derived for the different transport modes [16]. Particles undergoing random diffusion are expected to exhibit a constant MSD across different time scales. Their motions can be described by MSD( t ) = 4 Do t

(9.3)

where Do is the diffusion coefficient independent of time scale. For particles experiencing subdiffusive transport, their impeded motion can be described by

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MSD

Active: MSD = 4Dτ + ν2τ2

Diffusive: MSD = 4Dοτ Subdiffusive: MSD = Kτα

τ

Figure 9.2. Characteristic MSD curves for different transport modes.

MSD( t ) = Kt α

(9.4)

where K is a constant and a is an anomalous exponent <1. For particles undergoing active or convective transport, their motions can be described by MSD( t ) = 4 Dt + v2 t 2

(9.5)

where v represents the velocities of the particles. The v2t 2 term originally described convectional drift, but is commonly associated with active transport in an intracellular setting. At short t, the diffusive term dominates, and at long t, the velocity term dominates. As seen from the equations above, the MSD of tracked particles vary with t in distinct manners depending on the mode of transport involved. By plotting the MSD of individual particles against t (Figure 9.2), their mode of transport can be assessed graphically. For the case of simple diffusion, Do can be found from the slope of the line of best fit in a MSD versus t plot. Other coefficients such as D, K, α, and v can be determined by fitting the appropriate curve to the MSD data using equations that correspond to either subdiffusion or active transport. Suh et al. [17] developed a quantitative method to categorize tracked particles into their respective transport mode based on the relative change (RC) in their effective diffusivities. The diffusivities of purely diffusive particles remain constant at all time scales while particles undergoing active transport or subdiffusive motions display diffusivities that increase or decrease with time scale, respectively. Through the use of a standardized algorithm, a large MSD data set can be analyzed in a high-throughput manner. An RC value is calculated for each tracked particle according to the following equation:

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0.5

2 1.5 1

Active Upper RC Bound RC Bound

Diffusive Hindered

0 0 0.5 1 1.5 2 Time Scale (s) 2

Hindered

4 5 6 7 8 9 10 11 12

Time Scale (s)

A D H A A A A D A D H H A H H

Active Diffusive

1

0.5

0.5 0

1

1.5

Diffusive Lower

Short Time Scale

Active

1.5

0.5

0 0 0.5 1 1.5 2 Time Scale (s)

Classification

Lhort Time Scale

Diffusive

RCShort Time Scale

1

Measured RC 2

RCLong Time Scale

RCLong Time Scale

RCShort Time Scale

Monte Carlo Simulations 2 Upper RC 1.5 Active Bound

% Active Fractions % Diffusive Fractions % Hindered Diffusive Fraction

Hindered

0

4 5 6 7 8 9 10 11 12

Time Scale (s)

Figure 9.3. Steps involved in the RC method to categorize quantitatively individual particles into different transport modes. (Adpated from Ref. 18.)

RC = Deff ( t comp ) Deff ( t ref )

(9.6)

where t comp is the comparison time scale and t ref is a reference time scale that is shorter than t comp. RC values of purely diffusive particles are normally distributed around 1; thus statistical RC value boundaries for diffusive transport are first established through either Monte Carlo simulation or particle tracking in purely homogeneous medium (e.g., glycerol). RC values are then calculated for the experimental particles of interest. Particles with RC values larger than the theoretical upper bound are classified to be undergoing active transport, while particles with RC values less than the theoretical lower bound are classified as subdiffusive. The analysis procedures are often performed at both short and long time scales to find possible changes in transport modes across different time scales. The overall mode of transport can be determined by taking into account the transport modes involved at short and long time scales. The steps involved in RC analysis and the categorization of overall transport mechanism is summarized in Figure 9.3. While mathematical criteria such as RC value have been set forth to sort out the dominant mode of transport of tracked particles, qualitative aspects of the analysis remain indispensable. For example, particles exhibiting “pearlon-a-string” trajectories that are not characteristic of simple diffusion can be misclassified as diffusive unless their trajectories are visually examined. The choice of time scale when using the RC method also requires a considerable amount of understanding and experience in particle tracking. For technical details on performing particle tracking experiments and its data analysis, readers are referred to a guide written by Lai et al. [18].

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APPLICATIONS

9.4 9.4.1

167

APPLICATIONS Drug Carriers

Various drug carriers have been developed over the years to improve the efficacy of therapy and to minimize systemic toxicity through targeted delivery. Biodegradable polymeric nanoparticles (NPs) are increasingly popular drug carriers since they can achieve targeted delivery with a readily tunable release profile [19]. The intracellular fate of nanosized polymeric drug carriers has been studied by loading the NPs with fluorescent dyes and examining their subcellular localizations using fluorescence microscopy at fixed time points [20]. Unfortunately, SPT studies on NPs made of materials used in actual therapeutic applications are rare. Instead, model fluorescent probes are often used to explore how size and surface chemistry can affect the intracellular transport of NPs in real time. 9.4.1.1 Model Polymeric NPs Polymeric NPs, such as fluorescent polystyrene (PS) beads, have been used as model probes to investigate the intracellular trafficking behavior of nanosized polymeric drug/gene carriers. These model particles have high fluorescence output, allowing for easy tracking with fluorescence microscopy. They are also relatively monodisperse, thereby minimizing heterogeneity in transport properties due to inherent differences amongst the NPs. Using particle tracking, Suh et al. [17] quantitatively assessed the effect of surface modification with polyethylene glycol (PEG) on the cytoplasmic transport of PS beads in live cells. After cell entry through endocytosis, polymeric drug carriers may need to escape from endosomes to travel to their site of action, such as the nucleus, to exert their maximal therapeutic effect. The highly crowded cytoplasm can hinder the transport of drug carriers to their target organelles, therefore limiting their therapeutic efficacy. Surface modification with PEG, a hydrophilic polymer, was hypothesized to improve particle transport through the cytoplasm by reducing the nonspecific adhesion of particles to subcellular structures, such as cytoskeletal elements. PEGylated or unmodified NPs were microinjected into live cells and their transport properties were quantified by particle tracking. The fraction of PS particles that exhibit unhindered diffusive transport in the cytoplasm increases by 100% upon PEGylation, showing quantitatively how PEG can improve cytoplasmic transport of polymeric NPs. Model polymeric NPs have also been used to characterize new endocytotic pathways. Lai et al. [21] have shown that NPs can be sorted into different endocytotic pathways based on differences in particle size. Through inhibition of classical endocytosis pathways, it was found that 43-nm NPs are likely to undergo clathrin-mediated endocytosis while 23-nm NPs enter cells in a nonclathrin-, non-caveolae-dependent manner. Also, as opposed to 43-nm PS NPs, which are mostly sequestered within acidic vesicles (AVs) such as the

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late endosomes or lysosomes, 23-nm PS NPs appear to be sorted into nonacidic vesicles (NAVs), a pathway that could potentially reduce the degradation of polymeric drug carriers. While particles of both sizes are found to accumulate in the perinuclear region within 2 hours, their transport properties are revealed to be significantly different. The ensemble average MSD of NPs in NAV is about four fold slower than that of NPs in AVs [21, 22]. While around 45% of AVs undergo microtubule-dependent active transport, only 15% of NAVs display active transport, accounting for their slower overall transport. The slower transport of particles within NAVs can potentially increase residence time of drug carriers in the perinuclear region. This may in turn allow for extended release near the perinuclear region and may potentially be beneficial for drug carriers that target organelles near the nucleus. 9.4.1.2 Quantum Dots Quantum dots (QDs) are fluorescent nanocrystals made of semiconductor materials. They are much brighter than most conventional fluorophores and are highly stable against photobleaching [23]. Moreover, their narrow, easily tunable emission spectra allow them to be used in multicolor experiments [24]. Together, these optical properties make QDs excellent probes for fluorescence microscopy. The intrinsically hydrophobic QDs are poorly soluble in aqueous solution, making them ill-suited for biological studies. Development of biocompatible, amphiphilic surface coatings [23, 25, 26] for QDs has led to an exponential increase in the use of QDs as probes for biological processes [27]. To facilitate the entry of QDs into living cells, ligands that induce receptor-mediated endocytosis, such as folate [28] and transferrin [23], are conjugated onto the surface of QDs and these functionalized QDs can be used to investigate the effects of different ligands on endocytotic events [29–31]. While functionalized QDs are now relatively established biological probes, their role as model drug carriers in intracellular transport studies is just beginning to be explored using SPT. A study performed by Tada et al. [32] has demonstrated the applicability of QDs as model drug carriers in SPT studies. Notably, this study also set a remarkable precedent for the application of SPT in live animals. In this study, PEG-coated QDs were conjugated with monoclonal anti-HER2 antibody and their movements from tumor blood vessels to the perinuclear region of tumor cells were tracked in real time after administration into live mice models. After cellular uptake, the functionalized QDs are found to exhibit two distinct stages of directed motion, possibly along two different types of cytoskeletal filaments, before undergoing Brownian motion in a confined area in the highly congested perinuclear region. The velocities of the QDs suggest active transport while the “stop-and-go” phenomenon is attributed by the authors to steric hindrance within the complex biological environment of the cytoplasm. While the intracellular trafficking properties of these model NPs may vary from actual drug carriers due to fundamental differences in the materials used, their usefulness has nonetheless been demonstrated in the characterization

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of a nonclassical endocytotic pathway and establishing the effects of physiochemical parameters, such as size and surface modification, on the intracellular routing of NPs. 9.4.2

Gene Carriers

Another area of research that benefits from particle tracking is the field of gene therapy. In gene therapy, therapeutic genes are introduced into cells to restore functionality or elicit new responses. Carriers are frequently used to package the therapeutic genes, facilitating their cellular uptake and protecting them from degradation within the cytoplasm. The ability for gene delivery vectors to reach specific organelles, in particular, the nucleus, directly affects the efficacy of gene therapy. Hence the intracellular routing and transport properties of gene vectors have been extensively studied [33, 34]. By monitoring the trajectories of gene delivery vectors in real time with SPT, researchers are able to quantitatively define their transport characteristics, gaining additional insights that could help guide the rational design of delivery vehicles toward better gene delivery efficiencies. 9.4.2.1 Viral Vectors The evolved abilities of many viruses to deliver genetic material into the nucleus of human cells have been harnessed for therapeutic applications. Viral vectors are highly efficient at delivering their genetic cargo and the observed efficacy is often attributed to the expedient transport of viral vectors toward the nucleus [35]. By tracking the journey of individual viral particles in live cells, the endocytotic pathways and cellular components involved in the intracellular trafficking of viral vectors can be precisely and quantitatively characterized. Such knowledge can be used to improve the design of synthetic gene delivery systems through biomimicry. To study the intracellular transport of viruses in live cells, they first need to be fluorescently labeled. The surface of viruses can be chemically labeled with low molecular weight fluorescent dyes, such as Cy5 [36] and Alexa Fluor [37]. While viruses need to be sufficiently labeled so that their motions can be captured with fluorescence microscopy, it is also important to ensure that the degree of labeling does not affect the infectivity of the viruses [36, 38]. Another common approach is to genetically fuse fluorescent proteins to viral capsid proteins [39–41]. However, since fluorescent proteins can be relatively large attachments to the viral capsid subunits, such fluorescent viruses need to be carefully characterized to ensure capsid integrity and infectivity before they can used for SPT studies. The work by Seisenberger et al. [36] is a landmark SPT study that was performed on a popular gene delivery vector, the adeno-associated virus (AAV). It was found that multiple modes of transport are involved in the intracellular transport of AAV. The majority of the tracked viruses either (1) exhibit diffusive behavior with two distinct diffusivities (depending on whether the viruses are vesicle-bound) or (2) hindered transport. A small

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subset of viruses is also observed to be traveling at velocities that are in close agreement with microtubule-dependent active transport. Unexpected directed motions within the nucleus are also observed and it was postulated that AAV could be moving along tubular cytoplasmic structures that have extended into the nucleus. The influenza X-13 virus is another virus that has been studied extensively with SPT. By tracking the motions of individual endocytosed virus particles, Lakadamyali et al. [42] found that the trafficking of influenza viruses toward the nuclei can be divided into stages with distinct signature trajectories. Further quantitative analysis on these trajectories revealed characteristic velocities that are associated with transport along different cytoskeletal components. Similar multistaged trafficking patterns have also been observed for other viruses such as the simian virus 40 (SV40) [43], suggesting that this could be a prevalent feature among different viruses. Although the influenza virus is not a popular gene vector, the studies on influenza virus provide another clear demonstration of how SPT can be used to investigate and characterize viral infectious pathways. Intracellular trafficking pathways of other gene therapy related viruses, such as adenovirus [44, 45], herpes simplex virus (HSV) [46, 47], and human immunodeficiency virus (HIV) [41], have also been examined with SPT. A vast diversity of viruses exist in nature and there are considerable variations in how different viruses deliver their genomes into the cell nucleus. There are, however, some common features to the infectious pathways among different viruses, such as receptor-mediated endocytosis [38, 48, 49] and motor protein-guided active transport along cytoskeletal filaments [41, 42, 45, 50]. The intracellular transport of viruses can often be divided into distinct stages and different transport processes. SPT has been demonstrated to be an invaluable tool for discriminating and characterizing these different transport processes. Future investigations on how the infectivity of a virus particle is affected by its intracellular transport will continue to shape the design of advanced gene delivery vectors. 9.4.2.2 Nonviral Vectors Nonviral, synthetic vectors are attractive gene delivery systems since they are capable of delivering large genetic cargos with relatively little biosafety or immunogenicity concerns, especially when compared to viral vectors. However, the gene delivery efficiencies of synthetic vectors are generally considered to be lower than their viral counterparts. Inefficient intracellular transport is thought to be a critical barrier that lowers the effectiveness of synthetic vectors [2]. Particle tracking has been used to understand the intracellular transport properties of these vectors and to identify any rate-limiting steps involved. One particular nonviral carrier that has been studied extensively with SPT is polyethylene imine (PEI). PEI-based vectors are considered to be some of the most efficient synthetic vectors [51]. Being polycationic, PEI condenses negatively charged cargo genes into PEI/DNA nanocomplexes, protecting the genes and facilitating their intracellular transport. The intracellular trafficking

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171

trajectory of these nanocomplexes can be tracked when either the PEI [44, 52] or DNA [53] component is fluorescently labeled. The intracellular trafficking of PEI/DNA nanocomplexes prior to nuclear entry has been quantitatively characterized using particle tracking. A hypothesis in the field was that nonviral gene vectors reach the cell nucleus via random diffusion, a slow and potentially rate-limiting step due to the highly crowded nature of the cytoplasm [2, 3]. However, Suh et al. [52, 54] observed that PEI/DNA nanocomplexes accumulate in the perinuclear region within 30 minutes of transfection. By analyzing the trajectories of individual nanocomplexes, it was found that a substantial population of PEI/DNA particles exhibit MSDs associated with active transport. When PEI/DNA nanocomplexes are applied to cells treated with nocodazole, a microtubule depolymerizing agent, active transport of nanocompelexes is inhibited, suggesting that the active transport is microtubule dependent. By tracking the movement of PEI/DNA nanocomplexes along fluorescently labeled cytoskeletal filaments, Bausinger et al. [53] further confirmed that vesicle-bound PEI/DNA nanocompelxes indeed travel along microtubules at velocities comparable to those involved in kinesin- and dynein-mediated transport. While most of the aforementioned studies have been conducted in readily available model cell lines, Suk et al. [44] have shown that similar results are also obtained in primary neurons. In addition, by comparing the intracellular trafficking between adenovirus and PEI/DNA, it was found that failure of synthetic vectors to escape late endosomes and lysosomes may account for the lower gene delivery efficiencies of nonviral vectors. Taken together, particle tracking studies have shown that the cytoplasmic transport of nonviral gene carriers toward the nucleus is not a limiting factor. Instead, design of synthetic vectors should focus on overcoming bottlenecks that emerge subsequent to perinuclear accumulation, such as endosomal escape and nuclear uptake. There is an apparent trade-off in efficiency and biosafety between viral and nonviral gene vectors. Studies suggest that viral vectors enjoy higher gene delivery efficiencies because they are more proficient at overcoming certain intracellular barriers, such as endosomal escape and nuclear translocation. Much research is focused on endowing virus-like features onto nonviral vectors in order to improve nonviral gene delivery efficiency. Further comparisons in the intracellular transport mechanisms between the two systems with SPT should help contribute to the development of highly efficient gene delivery vectors with limited biosafety concerns.

9.5

CONCLUSION

SPT is a powerful technique capable of tracking the movement of individual particles at high spatial–temporal resolution. Analysis of individual particle trajectories within the complex intracellular environment has revealed highly heterogeneous and unique transport properties that are likely overlooked by

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ensemble average-based mobility assays. Continual advances in the development of imaging technologies and tracking algorithms should enhance the knowledge obtained from SPT studies. The expanding use of SPT to characterize intracellular trafficking pathways of different nanotherapeutics will continue to elucidate the link between intracellular transport properties and therapeutic efficacy, thereby guiding the future designs of nanoscale therapeutic devices. ACKNOWLEDGMENT This work was supported by the NSF Center for Biological and Environmental Nanotechnology (EEC-0647452).

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13. Cheezum, M. K., Walker, W. F., and Guilford, W. H. Quantitative comparison of algorithms for tracking single fluorescent particles. Biophys. J. 81: 2378–2388 (2001). 14. Gelles, J., Schnapp, B. J., and Sheetz, M. P. Tracking kinesin-driven movements with nanometre-scale precision. Nature 331: 450–453 (1988). 15. Jenkins, M. C. and Egelhaaf, S. U. Confocal microscopy of colloidal particles: towards reliable, optimum coordinates. Adv. Colloid Interface Sci. 136: 65–92 (2008). 16. Saxton, M. J. and Jacobson, K. Single-particle tracking: applications to membrane dynamics. Annu. Rev. Biophys. Biomol. Struct. 26: 373–399 (1997). 17. Suh, J., et al. PEGylation of nanoparticles improves their cytoplasmic transport. Int. J. Nanomed. 2: 735–741 (2007). 18. Lai S. K., et al. Design and characterization of gene transfer vectors. In: Gene Therapy Protocols, Vol. 3. Humana Press, Totowa, NJ, 2008, pp. 81–97. 19. Panyam, J. and Labhasetwar, V. Biodegradable nanoparticles for drug and gene delivery to cells and tissue. Adv. Drug Deliv. Rev. 55: 329–347 (2003). 20. Cartiera, M. S., et al. The uptake and intracellular fate of PLGA nanoparticles in epithelial cells. Biomaterials 30: 2790–2798 (2009). 21. Lai, S. K., et al. Privileged delivery of polymer nanoparticles to the perinuclear region of live cells via a non-clathrin, non-degradative pathway. Biomaterials 28: 2876–2884 (2007). 22. Lai, S. K., et al. Characterization of the intracellular dynamics of a non-degradative pathway accessed by polymer nanoparticles. J. Control. Release 125: 107–111 (2008). 23. Chan, W. C. W. and Nie, S. M. Quantum dot bioconjugates for ultrasensitive nonisotopic detection. Science 281: 2016–2018 (1998). 24. Jaiswal, J. K., et al. Long-term multiple color imaging of live cells using quantum dot bioconjugates. Nat. Biotechnol. 21: 47–51 (2003). 25. Bruchez, M., et al. Semiconductor nanocrystals as fluorescent biological labels. Science 281: 2013–2016 (1998). 26. Pellegrino, T., et al. Hydrophobic nanocrystals coated with an amphiphilic polymer shell: a general route to water soluble nanocrystals. Nano Lett. 4: 703–707 (2004). 27. Pinaud, F., et al. Advances in fluorescence imaging with quantum dot bio-probes. Biomaterials 27: 1679–1687 (2006). 28. Bharali, D. J., et al. Folate-receptor-mediated delivery of InP quantum dots for bioimaging using confocal and two-photon microscopy. J. Am. Chem. Soc. 127: 11364–11371 (2005). 29. Chen, B., et al. Transmembrane delivery of the cell-penetrating peptide conjugated semiconductor quantum dots. Langmuir 24: 11866–11871 (2008). 30. Delehanty, J. B., et al. Self-assembled quantum dot-peptide bioconjugates for selective intracellular delivery. Bioconjug. Chem. 17: 920–927 (2006). 31. Medintz, I. L., et al. Intracellular delivery of quantum dot-protein cargos mediated by cell penetrating peptides. Bioconjug. Chem. 19: 1785–1795 (2008). 32. Tada, H., et al. In vivo real-time tracking of single quantum dots conjugated with monoclonal anti-HER2 antibody in tumors of mice. Cancer Res. 67: 1138–1144 (2007).

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CHAPTER 10

Tracking Intracellular Polymer Localization Via Fluorescence Microscopy SIMON C. W. RICHARDSON

10.1 AN INTRODUCTION TO THE STUDY OF THE ENDOCYTIC AND SECRETORY PATHWAYS USING FLUORESCENT MICROSCOPY Organelles integral to the sorting and movement of material through the endocytic and secretory pathways (Figure 10.1) are critical to cellular homeostatic balance and perform a diverse array of physiological functions [1, 2]. Examples of the diversity of function performed by the two aforementioned pathways include communicating with and responding to the external environment [3], the secretion of structural material and enzymes (i.e., to remodel the cell’s environment) [4], the attenuation of external signals (i.e., regulating cell growth) [5], and the feeding of the cell through the internalization of essential nutrients such as iron [6] and albumin [7]. All of these functions require the sorting and compartmentalization of material into discrete membrane delimited structures and the regulated movement of material between these structures [1]. As this regulated movement requires not only the anterograde (forward) movement of material but also the retrograde (backward) recycling of cellular machinery involved in this regulation, there exists the possibility of exploiting not only the endocytic pathway and lysosomotrophic targeting for drug delivery [8], but also the transit of material between the secretory and endocytic pathways [9–11]. This occurs in nature and has been documented during the characterization of several bacterial and plant toxins [11, 12]. Over the last 35 years much interest has been shown in the use of synthetic, polymeric drug delivery systems that have demonstrated lysosomotrophic targeting in many types of mammalian cell (in vitro), in vivo and in the clinic Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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Secretory pathway

Endocytic pathway

Secretory Vesicles

Recycling Endosomes (TfR /RAB11a)

trans-Golgi (TGN46/38 or Ceramide)

Early Sorting Endosome (EEA1 /Rab5a)

MedialCis-Golgi (GM130) ER (BiP)

Cytosol

Early Endocytic Structures

Late Endosome (LAMP1-3 / M6PR / Rab7)

Hybrid Organelle

Lysosome (LAMP1-3)

Late Endocytic Structures

Nucleus

Figure 10.1. Generic intracellular trafficking pathways.

[8]. However, if investigators wish to optimize the intracellular movement, or kinetics of drug delivery at the intracellular level, they are obliged to investigate the intracellular fate of the delivery system [11]. As organelle intracellular localization and morphology are very diverse, it is often useful to identify subcellular compartments using physiological, immunological markers that have their roots in a fundamental understanding of organelle physiology and function [13]. Consequently, if the appropriate subcellular markers are employed, it is possible to effectively monitor the subcellular fate of nanoscale drug delivery systems in a temporal and qualitative manor [11]. It has been reported that it is possible (though extremely difficult) to use fluorescent microscopy to approximate the concentration of fluorescent material within a subcellular compartment. This difficulty is primarily due to the effects of fluorophore concentration-dependent quenching and possible pH-driven fluorescent phenomena in live cells. However, due to its inherent difficulty and complexity, this topic will not be considered further. The qualitative localization of synthetic biocompatible drug delivery systems is far from simple. One commonly used method capable of untangling

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the vast network of intracellular organelles is immunofluorescent microscopy. Typically, immunofluorescent microscopy requires the use of specific immunoreactive proteins, documented to mediate organelle-specific function. As these proteins define intracellular compartments, they can act as “landmarks” or markers. Often proteins that are responsible for regulating fusion to an organelle are chosen as these have the potential to represent “lock and key” type specificity through the very nature of their intracellular function. That said, the models used are often not perfect as there may be promiscuity associated with a particular protein’s function, that is, certain molecules may catalyze more than one fusion event (e.g., Syntaxin 6 [14, 15]), or cycle between more than one compartment (e.g., transferrin receptor [16]). Equivalently, there may be a great deal of primary protein sequence similarity between members of a particular family (i.e., Syntaxin 7 and Syntaxin 13), mediating fusion between the late endosome and early endosome, respectively [17], or the Rab family of GTPases [18, 19], which makes the immunological identification of specific family members difficult. Typical strategies used during the fluorescent identification of a subcellular compartment may be either (1) direct (i.e., by detecting a fluorescent physiological probe or fluorophore conjugated antibody) or (2) indirect (i.e., by immunolabeling the sample with a primary antibody specific to a protein enriched either within or upon the target compartment or organelle). The primary antibody is then identified using a fluorophore labeled secondary antibody (specific to the primary antibody) [16, 20, 21]. However, with respect to the subcellular localization of synthetic macromolecules, a paradox becomes apparent. Antibodies are macromolecules and are unable to penetrate cellular membranes. As the compartmentalization of macromolecules is one of the primary functions of the cellular membrane, it is obvious that, with the exception of antigens on the exofacial leaflet of the plasma membrane, an extracellular antibody applied by the researcher will not be able to assimilate its target unless some sort of membrane extraction or permeabilization is undertaken. Yet, it is also critical that intracellular macromolecules remain in place and are thus representative of the cell “in life.” Antibody intracellular penetration is usually achieved by lipid extraction using either solvents or detergents [20]. However, both forms of extraction potentially allow the escape (or movement) of poorly fixed (i.e., immobilized) synthetic material such as a soluble polymer forming part of a drug delivery system during sample preparation. This specific quandary set the basic tenet for this chapter, which will discuss ways of localizing soluble, nonfixed material to specific organelles or intracellular compartments (Section 10.4). But, before embarking on this discussion, it is critical to understand the aforementioned intracellular markers that will serve as counterstains, landmarks, and reference points, allowing the localization of a material with an unknown intracellular fate into a specific organelle or compartment over time. The following is not an exhaustive list, which would be extensive, but a list of tools that the author is familiar with and has used successfully [11].

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10.2 ENDOCYTIC AND SECRETORY ORGANELLES: FUNCTION AND MARKERS Figure 10.1 depicts both the secretory and endocytic membrane trafficking pathways. It is of note that this is a general depiction of cellular function as several cell types have evolved and differentiated to utilize these marker proteins to catalyze fusion between not one subset but many different organelles. An example of this is seen in the murine melanoma cell line B16 F10. B16 F10 cells localize Syntaxin 6 (normally found in the trans-Golgi network (TGN) and on early sorting endosomes) to lysosome-associated membrane glycoprotein (LAMP) positive structures, typically discreet and separate from the TGN and early endosome [15]. However, within the context of this work, Figure 10.1 represents a useful rule of thumb. Table 10.1 describes most of the reagents discussed herein and documents suppliers and catalog numbers. 10.2.1 Plasma Membrane At the cellular level, the first barrier encountered by a scientist wishing to enhance the bioavailability of a molecule with an intracellular target is the

TABLE 10.1

Commonly Used Reagents

Antigen/Reagent

Clone/Cat #

Source

EEA1 Syn6 TfR LAMP1 LAMP2 CD63 TGN38 TGN46 Texas Red anti-rabbit Texas Red anti-mouse Alexa Fluor 488 anti-rabbit Alexa Fluor 488 anti-mouse Cy5 anti-rabbit Cy5.5 anti-mouse Texas Red-NHS Alexa Fluor 488-NHS Lysotracker (Red) Lysotracker (Green) Texas Red-WGA Alexa Fluor 488-WGA Lysine fixable Dextran

(#E41120) (#S5542) (H68.4) (H4A3) (H4B4) (RFAC4) (T69020) (AHP500) (#T-2767) (#T-862) (#A11008) (#A11001) (611 110122) (610 713124) (T-6134) (A-20000) (L-7528) (L-7526) (W21405) (W112161) (D-1864)

Transduction Labs Transduction Labs Zymed DSHB* DSHB* Biodesign Int. Transduction Labs ABd SeroTech Invitrogen Invitrogen Invitrogen Invitrogen Rockland Inc. Rockland Inc. Invitrogen Invitrogen Invitrogen Invitrogen Invitrogen Invitrogen Invitrogen

*DSHB (Development studies hybridoma bank at the University of Iowa.

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plasma membrane. This may be the exofacial leaflet of an endothelial cell forming the lumen of a microcapillary in the lung [22], or of the endothelial cells forming part of the blood–brain barrier [23], or that of the target cell. Imaging the plasma membrane is useful when asking questions such as: “Is the ‘cargo’ or drug to be delivered inside the cell?” Even if the exact nature of the compartment occupied by the drug is not positively identified, data demonstrating that a substance has been internalized may still be informative. An excellent example of a directly labeled marker that can be used to label the plasma membrane is Texas Red-labeled Wheat Germ Agglutinin (TxR-WGA). If a fixed cell has not been subject to permeabilization (using either detergent of solvent), WGA can be used to label sialic acid and N-acetylglucosaminyl residues extending from the exofacial leaflet of the plasma membrane [24], the WGA being unable to penetrate this structure. It is also possible to image and delineate the plasma membrane living cells using WGA by keeping the cells at 0 °C. It is important to note that as WGA binds to sugar moieties on the cell surface, it will be internalized by live cells at 37 °C and, at this point, is no longer at the cell surface. 10.2.2 Early Endosome Following endocytic internalization, the first intracellular sorting station reached by exogenous material is the early endosome (EE), also called the sorting endosome. This will typically house material 5–10 min after internalization and is characterized by the presence of the small GTPase Rab5 [25], Rab5 effectors such as early endosomal antigen 1 (EEA1) [26] as well as a host of other proteins such as Syntaxin 13 [27]. In this acidic (pH typically about 6.5) organelle, material is sorted into a putative default catabolic (lysosomal) pathway or recycling pathways (Figure 10.1) [1, 28, 29]. This organelle also contains much of the sorting machinery required for the ubiquitin-mediated entry into multivesicular bodies that ultimately results in the degradation of trans-membrane proteins [29, 30]. Furthermore, the segregation of soluble material from receptors and membranes is a function of decreased pH and manipulating surface area to volume ratio, sorting the membrane into long reticular tubes [1]. EEA1 is a tethering protein that forms homo-oligomers and is responsible for bringing prospective fusion partners into proximal contact, that is, a donor vesicle and a recipient compartment, prior to soluble N-ethylmaleimidesensitive factor (NSF) attachment protein receptors (SNARE) pair formation [27]. SNARE proteins, such as the Syntaxin and vesicle associated membrane protein (VAMP) family of integral membrane proteins, are commonly held to be the proteins responsible for driving vesicular fusion [31]. Fusion is thought to be achieved by overcoming the entropy associated with pushing together two hydrated phospholipid bilayers, squeezing out water from between the two prospective fusion partners [32]. EEA1 can be

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immunolabeled using commercial monoclonal antibodies, which require an aldehyde rather than a cold solvent fix (Section 10.3.2). Unfortunately, this makes counterstaining samples fixed in aldehyde with a second antibody that require a cold solvent fix (i.e., LAMP-specific antibodies), difficult [16]. As fixation and permeabilization of the cell is required, this antibody is not usually a good choice for labeling live cells. If it is necessary to image the EEs of a live cell then the overexpression of Rab5a fused in frame to enhanced green fluorescent protein (eGFP) may be used [25]. Constitutively active Rab5 mutants (Rab5:Q79L-GFP) [33] are well characterized and should be used with care as their expression can cause the redistribution of intracellular markers (e.g., LAMP1) [34]. 10.2.3 Recycling Endosomes The recycling endosome (RE) is a Rab11 positive, reticular structure that has been documented to contain recycled material (i.e., apo-transferrin) 30 min after internalization [35]. The transferrin receptor (TfR) is an often-used marker for this organelle. The trafficking of TfR is markedly different from that of some other cellular receptors such as the epidermal growth factor receptor (EGFR), which is subject to ubiquitinylation and sorting into the lysosomal catabolic pathway following its activation [5]. From the early endosome, the TfR will follow either a kinetically fast or kinetically slow path back to the cell surface, the latter transiting through the Rab11 positive recycling endosome [18]. A consequence of this is that if TfR is used as an endocytic marker, there will be significant overlap between the Rab5 positive recycling endosome and the Rab11 positive recycling endosome [16]. TfR may be visualized indirectly by either immunolabeling using a commercially available antibody (in aldehyde fixed cells) or by using fluorescently labeled transferrin, which will be internalized and transit through the sorting and recycling endosomes in cells expressing TfR. Similar to the overexpression of Rab5a-GFP, Rab11a-fluorescent protein overexpression is well characterized and may be used to label recycling endosomes in living cells [36]. 10.2.4 Late Endosomes (LEs) LEs may be defined as LAMP positive, mannose-6-phosphate receptor (M6PR) positive structures. They are devoid of Rab5 and Rab11 and positive for Rab7, which regulates both homotypic and heterotypic fusion to LEs [34]. Morphologically, these structures have been documented by electron microscopy to contain internal membrane and are consequently also referred to as multivesicular endosomes. Material subject to endocytic capture may be documented in LEs after 60 min. There is some controversy as to the genesis of LEs with some reports documenting the transit of material from early sorting endosomes to LEs in vesicles (i.e., endocytic carrier vesicles [37]). However,

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there is also evidence documenting the maturation of early sorting endosomes into LEs [34]. LAMPs are highly glycosylated integral membrane proteins that contain a C-terminal cytosolic sorting signal that directs these proteins to late endocytic structures [38]. The luminal portion of these molecules is highly glycosylated, a physiological adaptation that is thought to help prevent the autolysis of the LE/Lys limiting membrane following lysosomal heterotypic fusion [39]. Furthermore, the primary protein structure of the LAMP family of proteins is highly species specific in terms of immunoreactivity. In primates and humans LAMP1 and LAMP2 specific antibodies will not cross-react with their rat orthologous (lysososmal glycoprotein (Lgp110 and Lgp120)). Commercially available antibodies to the above lysosomal proteins are readily available and work well after sample fixation using cold methanol. As previously mentioned, the LE is also defined by the presence of the M6PR [40]. The M6PR may be divided into two families, one being cation dependent (CD) and the other being cation independent (CI). CD-MP6R is a type 1 trans-membrane protein (having its C-terminus in the cytosol) that is responsible for transporting biogenic material destined for the lysosome from the Golgi body [41]. A well-characterized example of one such MP6R cargo would be the lysosomal protease cathepsin D. LAMP2 has also recently been reported to be involved in the process of lysosomal biogenesis, transporting molecules from the Golgi to the LE [42]. 10.2.5 Lysosomes (Lys) The lysosome is typically the terminal compartment of the endocytic system and may be thought of not as a “suicide bag” but as an enzyme storage depot. During endocytosis and the entry of material into the late endocytic compartment, material is sorted to the LE, which then fuses with Lys. The Lys then disgorge their contents into the lumen of the LE, creating a LE/Lys hybrid organelle where digestion occurs [43]. Although there is little literature describing the events after this fusion event, Lys are thought to reform from this LE/Lys hybrid organelle [44]. BSA–colloidal gold has been documented within a LAMP positive compartment after 60 min and aggregation of BSA– colloidal gold continues over the next 20 h, indicating that digestion of the BSA is occurring [45]. It is worthy of note that the catabolic nature of lysosomes makes their study difficult. However, tools exist that can help localize potentially labile material to late endocytic structures (such as enzyme inhibitors). Leupeptin (200 μM) fed to cells during and after a pulse of proteinaceous fluorescent labeled fluid phase or adsorptive marker (such as Texas Red-labeled Dextran, BSA, or WGA) can maintain the integrity of fluorophore-labeled material for up to 72 h. If the leupeptin is omitted there will be much nonspecific labeling of the cell and very little will be lysosomal (data not shown). Fluorophore-labeled WGA, BSA, and Dextran will yield almost complete colocalization with late

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endocytic markers following a 4–18-h incubation period (denoted the chase time) in the presence of leupeptin [11, 16]. This time is extended beyond the initial 60 min required for the detection of material within late endocytic structures to allow the clearing of earlier endocytic structures [16]. Although some cells express BSA-specific receptors [7], adsorptive endocytosis has been reported to drive the endocytosis of BSA (in specific instances). However, in the vast majority of cells and at high BSA concentration, the majority of BSA uptake appears to be nonspecific and concentration dependent (i.e., by fluid phase endocytic capture). This is a consequence of the relatively high concentrations of fluorophore-labeled BSA that are necessary to obtain an acceptable signal-to-noise ratio. Typically, this will be between 5 and 10 mg/mL BSA in PBS. This has been used to successfully delineate late endocytic (LAMP positive) structures in a range of developmentally different cell types derived from both fibroblasts and epithelia [11, 16, 46]. Commercially available lysine fixable Texas Red-labeled Dextran contains lysine residues. The lysine residues are intended to make this molecule susceptible to aldehyde fixation (the epsilon amine being able to react with an aldehyde). However, the charge associated with the lysine makes this molecule (in the author’s hands) very nonspecifically “sticky.” A result of this is a very high background, nonspecific, fluorescent signal. Although this molecule is less susceptible to degradation than BSA (internal signal persisting up to 72 h with or without leupeptin), it is of note that this advantage is often offset by the high background signal, lowering the signal-to-noise ratio obtained. After solubilization in PBS at 10 mg/mL, this reagent benefits from centrifugation at 100,000 g at 4 °C for 60 min. The supernatant may then be used without very bright nonsoluble aggregates interfering with the process of image acquisition. Similar to fluorophore-labeled BSA, Texas Red-labeled WGA can be allowed to bind to cell surface sialic acid and N-acetylglucosaminyl moieties and then be internalized. As this binding is absorptive (i.e., to membrane associated sugar residues, which are in abundance) [11], relatively low concentrations of WGA can be used to generate an acceptable signal-to-noise ratio. After a 1-h incubation with the fluorophore-labeled WGA in cell culture media (a pulse of WGA) at 37 °C and, following rigorous washing and a further 4-h incubation time (the chase period, also at 37 °C), WGA can be detected in LAMP positive structures (although these structures are larger than native LAMP positive structures). This phenomenon may be attributed to WGA having multiple sugar-binding sights and causing the crosslinking of membrane constituents [24]. Whether this is due to the increased trafficking of material to LEs or a reduction in the efficiency of LE reformation from the LE/Lys hybrid is not known. LysoTracker® is an aminic fluorophore that, at a neutral pH, freely permeates cell membranes. Once this reagent enters an acidic compartment, protonation impairs the molecule’s ability to cross membranes, leading to its selective

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accumulation in acidic membrane-bound intracellular structures. Consequently, in live cells with active membrane proton pumps, this reagent can be very successfully used to label acidic membrane-bound structures (i.e. EE, RE, LE, and Lys). At a 10-nM concentration, the photobleaching of the LysoTracker reagent has been an issue for the author [11]. This can be countered by increasing the concentration of the lysotracker in the cell media to 100 nM. This action also increases background noise and may interfere with membrane fusion events, which have been documented to be sensitive to vesicular luminal pH [47]. This is an issue as the accumulation of LysoTracker molecules within an acidic vesicle has been documented to cause an alkalizing effect [48]. 10.2.6 The Golgi Apparatus The Golgi body may be thought of as acting in opposition to the EE. As the EE mediates the sorting of material (and membrane components) entering the cell through the endocytic pathway (i.e., from the outside), the Golgi body sorts material that has been synthesized by the cell, directing it either to another intracellular location or into the secretory pathway. The Golgi may be divided into four subcompartments—the trans-Golgi network (TGN), the medial-Golgi (or Golgi ribbon), the cis-Golgi, and the endoplasmic reticulum to Golgi intermediate compartment (ERGIC) [49]. The TGN may be labeled using a variety of reagents in live or fixed cells. Antibodies specific to Syntaxin 6 are well-characterized Golgi markers. However, due the promiscuity of Syntaxin 6, it is not always the best choice when labeling the TGN [14, 15]. Additionally, varying expression levels of Syntaxin 6 may also result in poor signal-to-noise ratio. Monoclonal antibodies specific for TGN38 and TGN46 proteins may also be used to decorate the TGN. The species-specific expression of these proteins means they should be approached with care [50], for example, TGN38 is not expressed in Vero cells (whereas TGN46 is). Fluorescently labeled WGA may also be used to label the TGN in fixed, permeabilized cells. Ribosome inactivating proteins (RIPs) may also be chased into the Golgi and visualized by indirect immunofluorescence using polyclonal antibodies or following direct labeling [11]. If the TGN of live cells is to be imaged, then there are a variety of fluorescently labeled materials that have worked well for the author. Fluorescently labeled ceramide may be used to label the Golgi body in live cells and is suitable for use over a time course [11]. If the cis- or medial-Golgi is to be imaged, then monoclonal antibodies specific for the protein GM130 may be useful [11]. Like EEA1, GM130 is a tethering protein that helps regulate endoplasmic reticulum (ER) to Golgi transport. GM310 exists functionally within a complex of several other proteins, such as (though not exclusively) P115 and Giantin [51]. This protein may be labeled using commercially available antibodies after an aldehyde fix.

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10.2.7 The Endoplasmic Reticulum The ER is the site of lipid synthesis (smooth ER) and acts as part of the cell’s quality control mechanism, ensuring integral and secreted proteins are folded properly prior to sending these proteins to the Golgi for distribution around the cell or for secretion [52]. In order to facilitate this process, luminal proteins called “chaperones” are employed such as heat-shock protein 70 (HSP70). Monoclonal anti-immunoglobulin heavy chain binding protein (BiP) antibodies may be used to visualize the ER. BiP is an ER chaperone protein [53] and can readily be detected with a commercially available antibody. To draw this section to a close, it is of note that the researcher’s choice of fluorophore will impact on the success of imaging a given structure. If a probe is to be directly labeled and then subject to endocytic capture, Texas Red or Alexa Fluor 488 are good candidates. When chasing fluorophore-labeled BSA into cells, the author, despite exhausting Molecular Probe’s (now Invitrogen) comprehensive catalog (and obtaining consistent labeling efficiencies), was unable to image intracellular structures using a “blue” fluorophore. However, Marina Blue-labeled WGA was regularly and successfully used to label late endocytic structures.

10.3

FIXATION

Fixation is performed to ensure that the spatial location of material is not altered during the preparation of samples for microscopy. Commonly, one of two methods are employed, utilizing either solvents or aldehyde. If the immunodetection of specific markers is to be employed after fixation, a blocking step is necessary prior to antibody hybridization. This is typically performed using animal serum diluted to 2% (v/v) in phosphate buffered saline (PBS) and prevents the nonspecific adsorption of antibodies onto the sample [20]. It is important to note that it is critical not to use a blocking agent that will be recognized by the secondary or primary antibodies. For example, if the secondary antibodies have been raised in a donkey, donkey serum would be an excellent choice as a blocking agent. However, if the intended secondary antibodies were specific to rabbit IgGs, rabbit serum would not be a good choice as a blocking agent. Occasionally, unanticipated artifacts are encountered using the more standard blocking agents and if this is an issue Teleostean gelatine may be used in place of mammalian serum. If the researcher is in any doubt with regard to secondary antibody specificity, omitting the primary antibody from a negative control, processed in exactly the same way as the samples, will usually document which structures are being specifically labeled and which are not, the difference denoting specific labeling. Similarly, if a researcher using a confocal microscope is worried about bleed from one fluorophore’s channel to another, simply turning off the relevant laser and imaging as before will act as a negative control. If signal persists in the channel with

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Scheme A Formaldehyde Methylene Paraformaldehyde Glycol O H OH H (In water) HO O n OH H H H n=2-8 Scheme B Protein X R

O N N

R′

H N R′

Methylene Glycol H

+ NH2 H O

OH+ OH

Protein Y O

R″ N N

R″″

H N

O R″′

O

R′ HN

Fixed protein aggregate R″″ R″ O H N N R″′ O HN

NH

R″

O R

Figure 10.2. Aldehyde fixation.

the extinguished laser, another laser is exciting the fluorophore in question, which may be emitting light at several wavelengths, triggering a photodetector (not necessarily the intended one). Turning down any other active laser may solve this problem. 10.3.1 Aldehyde Fixation Formaldehyde is a gas that is soluble in aqueous media (the solution then being formalin) and can be used to crosslink proteins, resulting in the fixation of proteinaceous tissue [20]. Once in solution, formaldehyde exists in equilibrium with methylene glycol (Figure 10.2) [20]. As a starting material, paraformaldehyde is commonly used and is the polymeric form of formaldehyde, shown diagrammatically in Figure 10.2. However, to be used as a fixative, paraformaldehyde needs to be reduced into its monomeric components [20]. Formaldehyde solutions are typically prepared on the day of use [11, 16, 54]. Usually, for cells grown on a cover-slip, a 20-min fixation time using 2% (w/v) formaldehyde solution is adequate. If a 4% (w/v) formaldehyde solution is employed, the fixation time may be reduced to 2 min. Both fixation times are representative of fixation being performed at room temperature [11, 16, 54]. Following fixation, washing with PBS and quenching with 50-mM glycine removes or neutralizes excess unreacted aldehyde. It is common practice to perform the detergent extraction and quenching step simultaneously by adding 0.02% (v/v) Triton X-100 to the glycine solution. This will disorder cell membranes, allowing the penetration of macromolecules, such as antibodies, into the interior of the cell. Typically, a quenching and permeabilization time of 20 min at room temperature is not uncommon.

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10.3.2 Solvent Fixation Methanol cooled to −20 °C can be used to effect fixation, as this treatment will precipitate proteins and extract lipid from the cell. A variety of other solvents may be used and will also negate the need for a detergent extraction step as the lipidic portion of cell membranes will be dissolved and effectively discarded. Typically 5 min at −20 °C is adequate to achieve fixation. As methanol is volatile, it is important not to allow the samples to dry out. Consequently, it is a good idea to add PBS to the cold methanol after fixation and to act as soon as possible once the samples have been removed from the freezer. Rapid washing with an additional three charges of PBS will normally remove residual methanol [11, 16, 54].

10.3.3 Pre-fixation Methodologies Immunoreactive cytosolic pools of peripheral membrane proteins can mask a membrane-bound pool, obscuring the labeling of a target compartment. In this instance, cytosolic extraction may be desirable, in order to remove the immunoreactive pool of protein prior to fixation. There are several ways of achieving this end. Extraction with saponin prior to fixation removes membrane-bound cholesterol, effectively leaving pores in the cell membrane through which ions and proteins may escape or be washed out [11]. If this step is to be used “prealdehyde fixation,” it is vital to reduce the time of the subsequent detergent extraction step (author uses 5 min). The use of this agent prefixation may (like leupeptin) slightly alter vesicular morphology, giving an expanded “doughnutlike” appearance [11]. Streptolysin O (SLO) is a bacterial toxin that also can be used to generate pores in the plasma membrane. Typically, it is introduced to the cell at 0 °C in the presence of calcium ions. Once the calcium ions are chelated using ethylene glycol tetraacetic acid (EGTA), the SLO will form pores large enough for proteins to escape the intracellular environment while keeping large intracellular supramolecular assemblies inside the cell [55]. The use of some lipidic transfection reagents can, at times of up to 48 h post-transfection, significantly alter the distribution of intracellular markers such as the LAMP proteins, independently of the transgene being expressed. Further cellular morphology can also be altered, revealing a highly vacuolated phenotype. Care should be taken when using transient transfection and lipidic vectors that any observed effects are not attributable to the transfection vehicle. With the careful selection of transfection vehicle and the application of positive and negative controls, it is possible to control for any transfection vector induced artifacts. As has been previously mentioned [11, 16], leupeptin is a useful and necessary tool when using fluorophore-labeled biodegradable markers for temporally dissecting the endocytic pathway. However, like saponin, leu-

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peptin may also induce artifacts in the form of expanded vesicular structures [11, 16].

10.4

SAMPLE PREPARATION FOR POORLY FIXABLE MATERIAL

One solution to the “washing out” of a poorly fixed, soluble, drug delivery system (i.e., Oregon Green-labeled poly(ethylene glycol), Dextrin, or N-(2 hydroxypropyl methacrylamide) has recently been published by the author. Central to this methodology is the use and characterization of internalized markers such as fluorophore-labeled BSA (i.e., TxR-BSA) or TxR-WGA as previously discussed (Section 10.2.5). The kinetics of fluorophore-labeled BSA (fBSA) movement through early and late endocytic structures can be delineated and documented using standard fixation methodologies in conjunction with immunolabeling and may be viewed as being relatively constant under similar growth conditions. Once the temporal location of the fBSA is known, fBSA may be coincubated with a soluble, synthetic drug delivery system (DDS) (labeled with a second, nonspectrally overlapping fluorophore). By virtue of any colocalization between the DDS (an unknown quantity), and the precharacterized endocytic marker (a known quantity), the intracellular kinetics and localization of the DDS may be identified. Furthermore, this may be achieved without the need for a permeabilization step, negating the opportunities for poorly fixed or nonfixed material (i.e., the DDS) to move, distorting the data set obtained. As antibodies have already been used to characterize the localization of fBSA over time, they can be dispensed with during the characterization of the DDS. Consequently, it is also possible to utilize this methodology to document DDS localization in live or fixed cells relative to fBSA, which has already been characterized using immunofluorescent subcellular markers such as EEA1, LAMP, and GM130 [11]. This technique assumes that the unknown quantity will not alter the distribution of the marker (fBSA) and this assumption needs to be controlled for. Particular care needs to be taken regarding the osmolarity of the buffers used. PBS, in the hands of the author, has worked well.

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CHAPTER 11

Can QSAR Models Describing SmallMolecule Xenobiotics Give Useful Tips for Predicting Uptake and Localization of Nanoparticles in Living Cells? And If Not, Why Not? RICHARD W. HOROBIN Division of Integrated Biology, FBLS, The University of Glasgow, Glasgow, Scotland, United Kingdom

11.1

QUESTIONS AND THEIR CONTEXT

Nanoparticles come with a wide variety of composition and form, and possess a wide variety of surface features. When considering their biomedical applications, cellular uptake and localization is almost always of significance, whether as drug delivery agents or as biosensors. Consequently, being able to predict these behaviors from a knowledge of the physicochemical properties of nanoparticles would be very advantageous. And yet, apart from the finetuning of endocytosis (for some recent work see Hild et al. [1], Rejman et al. [2], and Sánchez-Martín et al. [3]), this is not deemed possible—and moreover what is known is rarely spelled out in any summary and explicit way. But have we been too pessimistic? The question of whether intracellular localization of nanoparticles can be predicted is worth asking again, since intracellular uptake and localization in many organelles can now be predicted for small-molecule species. This has been achieved using both first-principles and quantitative structure–activity Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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(QSAR) models. While, as discussed below, it was unlikely that such models would apply in detail to nanoparticles, nevertheless can such approaches at least give us tips for predicting the localization of such species?

11.2

SOME BACKGROUND: THE MEANING OF “NANOPARTICLES”

There are many radically different types of nanoparticles—including aggregates of hydrophobic organics such as hypericin or porphyrins; liposomes; modified proteins (e.g., albumin and gelatin); metal crystallites such as iron and gold; crystallites of iron oxide, silica, or titania; quantum dots; polymers; and dendrimers. The notion that such particles have features in common, other that is than size, is perhaps odd but is nevertheless a widely implicit assumption of “nanoscience.” In any event, nanoparticles certainly do differ radically from “small molecules.” Thus sheer size plus surface (i.e., not overall) properties may be controlling factors in the interactions of nanoparticles with living systems. Perhaps nanoparticles should not be regarded as one end of a spectrum that has small molecules at the other end; rather, they are on a different distribution altogether. Another, unavoidable, issue raised by asking “What are nanoparticles?” is that of nomenclature and definition. Defining a nanoparticle is somewhat arbitrary, one widely held position merely being that it is a particle falling between 10 and 100 nm in “diameter.” But this lacks precision and significance, both physicochemical and biological. Indeed, such a definition should be seen more as a staking out of academic territory rather than as entailing boundaries emerging from the biological and physical worlds. So while “diatom nanotechnology” [4] exceeds such a range, certain “nanocrystal lumiphores” [5] fall below it. And do we have to ignore possible data from “colloid chemistry” or indeed from “micro” particles? Indeed we do not, since the operational conclusion of researchers as shown by journal terminology is clearly less restrictive, with illuminating comparisons emerging, for instance, from comparisons of particles ranging from less than 10 nm up to more than 1000 nm. Also, beyond mere size, the effects of the amazing variety of entities we are dealing with, noted above, is significant—and yet all these variations are on occasion shoehorned into the “nano” category.

11.3 PREDICTING UPTAKE AND LOCALIZATION OF SMALLMOLECULE XENOBIOTICS USING QSAR MODELS For small-molecule xenobiotics (typically < 1000 daltons) it has been found that rules can be specified connecting the physicochemical character of the xenobiotics with their uptake and localization in live cells of many types. Two quite different and workable strategies have been investigated. First is a

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physicochemically first-principles approach, where interaction of small xenobiotics with a simplified model cell is described by utilizing the Henderson– Hasselbach and Nerst–Planck equations, Fick’s law of diffusion, and a hydrophil–lipophil partition constant [6, 7]. Although this approach is currently limited in the number of organelles that can be modeled, it has being applied successfully in several contexts of biological significance, for example, soil–water [8] and water–fish [9] partitioning of ionizable organic chemicals. The other strategy involves correlational QSAR models, formally expressed as dichotomous decision rules. The limits of this approach are different, namely, having the correct set of exemplar xenobiotics to accurately specify the models. Again, this has achieved application over a wide range of biological systems, from microorganisms to whole plants. However, neither of these approaches is universally applicable at this time, as shown in a recent comparative study [10]. Intriguingly, both strategies are based on overall molecular properties, suggesting that the widely adopted “tag” or “motif” approach is not in actuality the only game in town. In this chapter it is the second strategy, based on QSAR decision rules, that is utilized. The relevant QSAR decision rules relate the uptake of xenobiotics (e.g., drugs and fluorescent probes) into living cells to the structure of the xenobiotics as described numerically. These correlations were established following direct microscopic observation of the interaction of a wide range of fluorescent compounds with live cells. The strategy requires the numerical description of the structure of xenobiotics in terms of a few key physicochemical properties. Thus electric charge (Z) is specified directly, and acid/base strength described in terms of pKa values. Overall size is modeled using the relative molecular mass (i.e., molecular weight, MW). Overall size of the conjugated, usually substantially aromatic, system is modeled using the conjugated bond number (CBN) parameter and that of the largest planar conjugated region by the largest conjugated fragment (LCF) parameter. Lipophilicity/hydrophilicity is modeled using the logarithm of the octanol–water partition coefficient (log P), and amphilicity by the amphiphilicity index (AI). For information concerning the derivation, application, and limits of these parameters see Horobin [11] and Christensen et al. [12]. QSAR decision rule models are now available to predict the occurrence, or nonoccurrence, of a wide variety of processes: namely, membrane permeability, adhesion to the cell surface, exclusion from a cell’s interior, and passage through internal cellular barriers. Most of these processes can occur by more than one mechanism, each of which requires a separate model, as discussed in Horobin [11] and Stockert et al. [13]. A summary of the available QSAR decision rule models is provided in Figure 11.1. QSAR decision rule models are also available to predict the accumulation of xenobiotics in a variety of cellular regions and organelles. To date these include generic biomembranes [11] as well as those of the endoplasmic reticulum [14], Golgi apparatus [15], and plasma membrane [11]; the cytosol [16];

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CAN QSAR MODELS FOR SMALL-MOLECULE XENOBIOTICS GIVE USEFUL TIPS?

Membrane permeability, of plasmalemma & others

Fluid phase (pinocytosis)

Adsorptive

Lysosome or other acidic organelle Passage through gap junction or plasmadesmata Phagosome

Phagocytosis

nucleus No binding to, or passage through, plasma membrane Passage through nuclear pore

Figure 11.1. Relationships between chemical properties of small-molecule xenobiotics (e.g., drugs or fluorescent probes) and their cellular uptake and intracellular transfer behavior in live cells. Structural requirements for occurrence of the arrowed processes are summarized below; see text for explanation of terms. Adsorptive endocytosis requires AI or log P > 8 or CBN > 40. Fluid-phase endocytosis requires that if CBN < 40 then log P < 0 or if CBN > 40, then log P < −10. Membrane permeability, plasmalemma, and others require that 8 > AI or log P > 0; CBN < 40. No binding to cells requires that if pKa << 7 or >>7 and CBN < 40, then log P < 0; or if pKa << 7 or >>7 and CBN > 40, then log P < −10; or if pKa ∼ 7 and CBN < 40, then log P (most ionized species) <−5. Note that uptake into cells is still possible if pinocytosis occurs. Passage through gap junctions or plasmadesmata requires that MW < 1000 (gap junction) or MW < 700 (plasmadesmata). Passage through nuclear pores requires that MW < 5000. Phagocytosis requires particles with a “diameter” greater than ∼500 nm.

lysosomes and other acidic structures [17]; mitochondria [18, 19]; nuclear chromatin [11, 20]; and nucleoli and ribosomes (Rashid-Doubell, Stockert, and Horobin, unpublished findings). Again, there are often multiple accumulation mechanisms for each organelle, each with a separate QSAR model. A summary of the available decision rules is given in Figure 11.2. There are various limitations of such models. For instance, some needed structure parameters may be unavailable. Examples of such problems are the lack of log P fragment values for key moieties, and the lack of micro pKa values for certain polybasic compounds. Moreover, these QSAR models, being based on simple physicochemical effects, will not cope with the effects of sophisticated targeting systems, for example, the involvement of antibody–antigen, enzyme–substrate, or peptide–receptor interactions, or of nucleic acid complementarity. This is a radical limitation, given that such targeting strategies are often used with nanoparticle delivery pharmaceuticals.

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Membrane bound

Lipid domains Protein domains

Plasma membrane

Precipitation trapping of weak acids

Potential driven &/or cardiolipin complexation

Ion-trapping of weak bases

Ion-trapping of weak acids

Mitochondria

Lysosomes & acidic organelles

Fluid lipid domains, e.g., apoptotic cells

Outer, fluid, membrane

Phagosome Endoplasmic reticulum membrane

Generic biomembranes Ribosomal RNA

Golgi membrane

Nucleolus, nucleic acids Fat droplet

Cytosol Histones

Nuclear chromatin DNA

Figure 11.2. Relationships between chemical properties of small-molecule xenobiotics (e.g., drugs or dyes) and their patterns of localization in live cells. Structural requirements for localization in the named organelles and other sites are summarized below; see text for explanation of terms. Cytosol requires AI < 3.5; CBN < 40; 5 > log P > 0; Z = 0. Endoplasmic reticulum membranes require that 6 > AI > 3.5; 6 > log P > 0; Z > 0. Fat droplets require that AI < 3.5; HGS < ∼20; log P > 5.0; pKa (if relevant) < 6. Generic biomembranes, for reversible uptake, require that 8 > AI or log P > 5. Golgi membranes require that 5 > log P > 3. Lysosomes/acidic organelles, weak base ion-trapping requires that 0 > log P (cation) > −5; 10 > pKa > 6; Z > 0. Lysosomes/acidic organelles, weak acid precipitation trapping requires that log P (free acid) > 0; pKa = 7 ± 3; Z(most ionized species) < 0. Lysosomes and endosomes, membranes, lipid domains require that AI or log P > 8. Lysosomes and endosomes, membranes, protein domains require that CBN > 40. Mitochrondria, potential driven and/or cardiolipid complexation require that 5 > log P > 0; pKa > 12; Z > 0. Mitochrondria, ion trapping of weak acids requires that 5 > log P (least ionized species) > 0; pKa = 7 ± 3; Z(most ionized species) < 0. Mitochondria, outer membrane, require that 6 > AI > 3.5; log P > 0; Z > 0. Nuclear DNA, chromatin requires that LCF > 24; −2 > log P(cation) > 0; pKa > 10; Z(ionized species) > 0. Nuclear histones require that if Z(most ionized species) < 0, and CBN > 40, then log P > −10; or if Z < 0 and CBN > 16, then 8 > log P(anion) > 0. Nucleoli, nucleic acids require that LCF > 17; 8 > log P(least ionized species) > 0; pKa > 10; Z > 0. Phagosomes require particles with a “diameter” greater than ∼500 nm. Plasma membrane uptake, fluid lipid domains (e.g., in apoptotic cells) require that ∼5.5 > AI > 3.5; log P < 5.0; HGS > 400. Plasma membrane uptake, lipid domains require that AI or log P > 8 or AI > 5; HGS > 400. Plasma membrane uptake, protein domains require that log P < −10; CBN > 40. Ribosomal RNA requires that LCF > 17; 8 > log P(least ionized species) > 0; pKa > 10; Z > 0.

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11.4 MORE BACKGROUND: WHAT FACTORS CONTROL UPTAKE AND INTRACELLULAR LOCALIZATION OF NANOPARTICLES? First, a disclaimer: this chapter, as acknowledged above, does not discuss the major approaches to the targeting of nanoparticles adopted to date. Most investigators have been focused on the development and application of “smart targeting” mediated by various entities. Thus a recent review of the cellular and intracellular targeting of quantum dots gave considerable attention to work involving targeting by antibodies, by ligands for cell-surface receptors such as folate, and by peptide ligands for various tumor cell markers, with some attention also directed at ligands binding to receptors for angiotensin and neurotransmitters [1]. Other interactions of nanoparticles and cells were discussed under the heading of “nonspecific particle uptake,” which is precisely the type of phenomenon we will be considering in the present chapter. Given the focus on smart targeting, it is perhaps not surprising that there is a paucity of rigorous investigations into such “nonspecific” mechanisms of uptake and localization of nanoparticles within living cells, not only because of the issues noted above, but also because preparation of novel nanoparticles is within the technical capabilities of many chemical and related laboratories. Indeed, a wait-and-see approach to such a complex problem as nanoparticle uptake and targeting may be a valid, if not the only, way to successfully solve pressing problems of drug delivery and toxicology. However, we will now briefly review some of the available rigorous studies, first concerning uptake into cells, and then the post-uptake localization within cells. It is immediately apparent, and of course no surprise, that the most dramatic particle-associated variable influencing cellular uptake is the size of the nanoparticle. Of course, nanoparticles are (depending on definitions, see above) very large objects to pass through the plasma membrane in the same manner that is easily possible for small molecules entering a cell by passive diffusion. Instead by far the most common mode of entry is by endocytosis, whether the particles are lipoplexes [21], liposomes [22], quantum dots [23], or some other variety. At the extreme, it was observed as long ago as the early twentieth century—when India ink was used as the probe—that particles exceeding approximately 0.5 μm in diameter were extensively taken up by phagocytes [24]; for more modern accounts see Besterman and Low [25] and Haas [26]. However, for “true” nanoparticles, a discriminating and critical investigation has been carried out more recently by Hoekstra’s group [2]. This involved exposing nonphagocytic eukaryotic cells to fluorescent latex beads ranging in size from 50 to 1000 nm. All except the largest (1000-nm) particles were taken up, by energy-dependent processes. In the size range up to 200 nm this involved a clathrin-mediated process, with the particles finally arriving in late endosomal/lysosomal compartments. In the larger size range, above 200 nm to below 1000 nm, uptake involved a caveolae-mediated process, with the final destination not being lysosomal. This latter process was distinct from

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macropinocytosis, which can also favor larger particles [27]. These size criteria have been found applicable to a variety of cell types (e.g., epithelial [28] and fibroblast [29]) and to diverse types of particle (e.g., lipoplexes [30] and porphyrin aggregates [31]. Endocytic uptake, however, is modulated by other factors. One such is a particle’s surface electric charge. Thus endocytotic uptake of nanoparticles of very widely varying types is often facilitated by a positive surface charge, as reviewed, for instance, by Blau et al. [32]. This is generally considered to be due to an initial adsorption to surface negative charges, usually on the plasma membrane. Indeed, this does often result in an observed increase (short or long term) of the surface adhesion of the nanoparticles at that stage [33]. However, a positive surface charge does not always result in entry into the cell, since if a cell has an external coating of an anionic glygosaminoglycan, for instance, then a cationic nanoparticle can be trapped in that layer [34]. Another surface property dramatically modifying endocytic uptake is the lipophilicity or hydrophilicity of the particle’s surface. In vivo, it was observed many years ago that after intravenous administration nanoparticles were rapidly removed from the circulation as a result of being phagocytosed by macrophages, especially in bone marrow, liver, lungs, and spleen [35]. It was shown that this was largely due to adsorption of proteins onto the particle surface resulting in opsonization [36]. In keeping with this, occluding the hydrophobic surface with hydrophilic moieties reduces protein binding and consequently phagocytic endocytosis, as demonstrated quantitatively by Chang’s group [37]. Hydrophilic materials as varied as polyethylene glycol [38] and polysaccharides [39] have been used in this way. Of course, this reduction of competition from macrophages enhances uptake into other cell types by other endocytic mechanisms. Before moving on to consider the intracellular localization of nanoparticles, it should be noted that their entry into cells is not entirely restricted to endocytosis. For instance, membrane fusion can also enable internalization, albeit largely that of liposomes. However, even for those structures it is not the most common mechanism (although it has given rise to some wonderful electron micrographs; e.g., see Dini et al. [40], Figure 5d). Crucially, membrane fusion can result in direct delivery of hydrophilic materials into the cytosol [41]. In addition, some nanoparticles do cross the plasma membrane. Least surprising is to see this occuring with small (∼1 nm) fullerene derivatives, which include lipophilic species [42, 43]. More surprisingly, it has also been reported for particles with diameters above 10 nm. Thus Jablonski et al. [44] have developed a system involving coating the particle with a cationic species with a subsequent application of a hydrophobic counterion, pyrenebutyrate, in the published example. This is not a permeabilization process, and the negative surface charge of the particle means that the plasma membrane potential strongly influences uptake. Another process involves coating quantum dots with amphiphilic polymers [45].

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Post-uptake intracellular localization of nanoparticles will now be considered. As described above, particle size strongly influences which particular endocytotic mechanism is involved. Consequently, particle size substantially controls whether the nanoparticle will end up in a lysosome or an acidified late endosome, or instead in a nonacidified vesicle. Once again, however, a particle’s electric charge modulates this: for instance, in directing down which intracellular pathway the nanoparticles travel. A recently reported example involves the application of various anionic and cationic derivatives of poly(ethylene glycol)-d,l-polylactide nanoparticles, all with diameters between 50 and 100 nm, to cultured epithelial cells. Most were taken up via the clathrin pathway. However, whereas many anionic particles transited through the lysosomal route, cationic particles typically did not but instead diverted to the lateral plasma membrane [46]. Ionization can also play a role in escape from endosomes and related vesicles. For instance, weak bases such as protonsponge dendrimers can become ionized within acidified vesicles, thus dramatically raising the local osmotic pressure, favoring vesicle rupture [47]. Amphilicity, and the molecular features influencing this property, also has an impact on membrane rupture and so on escape of particles from endosomes. For instance, nanoparticles will escape more readily if their surfaces carry amphiphils with surfactant properties. In particular, this effect is influencial in lipoplexes, where chain length effects have been discussed using the structure parameter AI [48]. A related, but distinct, property influencing membrane rupture and fusion is lipid shape. In this case it is membrane curvature that is the key property. This is controlled both by the relative sizes of the head group and hydrocarbon tail, and by the cross-sectional area of the head group; for an accessible account of which Israelachvili’s [49] formulation is still to be recommended.

11.5 SO CAN SMALL-MOLECULE QSAR MODELS TELL US ANYTHING ABOUT NANOPARTICLE UPTAKE AND LOCALIZATION? As discussed above, the factors found to be of significance do include those considered in the small-molecule models—namely, amphilicity, electric charge, head group size, lipophilicity, and pKa. However, it is hard to see how the small-molecule models can be applied directly to such vast structures as nanoparticles. So is that all? Are there are no regularities to which the smallmolecule QSAR models can be applied, or even any useful tips? It does seem that there are two distinct ways in which numerical structure–localization rules may be usefully applied to nanoparticle uptake into cells. First, consider nanostructures at the lower end of the size range, such as C60 fullerene derivatives. Such, if lipophilic, do penetrate the plasma membrane directly rather than by endocytosis. In one study, a dicarboxylic C60 fullerene was found to localize in biomembranes including the plasma mem-

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brane and, most markedly, in mitochondria [43]. This fullerene is small enough for structure parameters to be estimated, and it was found to be a lipophilic weak acid, with some species being extremely lipophilic. Plugging the parameters into the relevant QSAR models, accumulation of the weak acid by iontrapping in mitochondria, and more general partitioning of the strongly lipophilic species into membranes, with superlipophilic species favoring the plasma membrane, matched observation. However, when a hexacarboxylic acid C60 fullerene was investigated it was observed to still accumulate in mitochondria, with cytoskeletal keratin fiber binding also occuring [42]. Again, estimation of structure parameters and application of the QSAR models gave congruent predictions, again of lipophilic weak acid ion trapping and, in this case, protein binding by the hydrophilic carboxylate anions. Morover, some larger nanoparticles do behave somewhat as suggested by the small-molecule models. Examples are provided by mitochondrial targeting of nanoparticles with lipophilic and cationic surfaces, noting that small molecules with lipophilic and cationic character are typically mitochondrially targeted. One such instance involved DQAsomes derived from dequalinium and its derivatives in intact cultured cells [16]. Another involved linking triphenylphosphonium to hydroxypropylmethacrylamide polymers [50], which nanoparticles appeared to show localization to mitochondria, but only with the smallest (<5 kDa) species, as larger particles were restricted to the lysosomal compartment. Such observations suggest that with the smallest nanoparticles available QSAR small-molecule models do have some direct applicability; and even with larger particles the models, although not rigorously applicable, can give useful tips. Perhaps further experimental studies of nanosystems of this type would have predictive value. In particular, it would be useful to know whether the loss of predictability with increase in size is merely a methodological limit of the modeling system or also involves physicochemical limits.

11.6 SIMPLE NUMERICAL SUMMARIES OF NANOPARTICLE UPTAKE AND INTRACELLULAR LOCALIZATION Second, note the striking fact that authors discussing the various physicochemical influences on nanoparticle uptake and localization have been curiously reluctant to provide simple summaries of factors involved. This seems unduely reticent: whatever its limitations such a summary would certainly have some utility. Consequently, in Figure 11.3, in a more optimistic spirit, I offer a graphical summary of the data mentioned in this chapter. One final complication must, however, be pointed out at this point. Very dramatic cell-line effects can occur when nanoparticles interact with live cells— there are indeed “smart cells” as well as “smart targeting” of nanoparticles [51, 52].

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Particle size effects

« 50nm

phagocytosis

200-500nm Macropinocytosis

Mit

caveolinmediated

Pha Lys

Ma

CyPr

50-200 nm BioM clathrin-mediated endocytosis

Uptake & localization show log P, pKa, Z effects

500-1000nm

Particle charge effects

End

Al & pKa dependent

Coat with cationic species

Coat with various hydrophils

Coat with cationic species, then a hydrophobic anion OR with an amphiphilic block copolymer

Tactics to enhance different types of cellular uptake

Figure 11.3. Graphical summary of physicochemical factors influencing cellular uptake and intracellular localization of nanoparticles. Targeted organelles abbreviated as follows: BioM, generic biological membranes; CyPr, cytoskeletal proteins; End, endosomes: Lys, lysosome; Ma, macropinosome; Mit, mitochondrium; Pha, phagosome.

ACKNOWLEDGMENT I wish to acknowledge Prof. I. McGrath, Division of Integrated Biology, FBLS, The University of Glasgow, Scotland, for provision of facilities.

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CHAPTER 12

Self-Unpacking Gene Delivery Scaffolds MILLICENT O. SULLIVAN Department of Chemical Engineering, University of Delaware, Newark, Delaware

12.1

INTRODUCTION

Gene therapy can be defined as the introduction of genetic material into specific cells of an individual for the purpose of treating disease [1]. Gene therapy has the potential to treat a broad variety of human diseases, including those that involve heritable genetic disorders such as cystic fibrosis, hemophilia, muscular dystrophy, and familial hypercholesterolemia that could be treated by the stable expression of one or a few mutant genes within a target tissue. More broadly, gene therapy has been explored for the treatment of multiple acquired diseases, including cancer, infectious diseases, cardiovascular disease, wound healing, Parkinson’s disease, and Alzheimer’s disease. In these cases, the goals of the therapy are more diverse and might entail either stable or transient gene expression within the target cell population. For example, cancer gene therapies under exploration include those in which the goal is to permanently replace the function of a missing or altered gene, such as the retinoblastoma susceptibility gene, whose inactivation has been linked to the neoplastic phenotype in human retinoblastoma and osteosarcoma cells [2]. Other cancer gene therapies under development aim to promote the transient expression or overexpression of genes that stimulate natural defense mechanisms, act as “suicide genes” to promote tumor cell apoptosis, enhance the response to drug therapies, or block the development of tumor-induced angiogenesis. Despite broad interest in the potential applications for gene therapy, its success rate has been very low. The first clinical trial for gene therapy was

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initiated in 1990 and was directed at the treatment of severe combined immunodeficiency (SCID) [3]. The first clinical successes did not occur until 10 years later, in trials involving SCID, hemophilia, and squamous cell carcinoma of the head and neck [4–6]. Since 1990, 1537 clinical trials have been approved with a disappointing overall success rate [7]. Significant hurdles remain to effective gene therapy, including the identification of therapeutic target genes, the identification and functional elucidation of target therapeutic nucleic acids, the elucidation of methods for inducing safe chromosomal integration when desired, and the development of controllable, efficient, and safe methods for gene delivery. It is clear that, in most cases, tight regulation of the levels and duration of gene expression will be critical [8]. Gene delivery has been called the Achilles heel of gene therapy [9] and, in general, is regarded as the most significant hurdle to its success [10].

12.2 METHODS OF GENE PACKAGING Two major classes of gene delivery vehicles have been widely pursued: recombinant viruses (viral) and synthetic materials (nonviral). Viral vehicles are created from recombinant viruses in which a portion of the viral genome has been replaced with a therapeutic gene of interest. Synthetic vehicles are typically comprised of polycationic lipids or polymers, which self-assemble with DNA to form lipo- or polyplexes, respectively (Figure 12.1). 12.2.1 Viral Gene Delivery Vehicles Viruses have evolved for the purpose of highly efficient gene delivery, and viral vehicles are the more widely explored class of delivery vehicles reported

Figure 12.1. Polyplex self-assembly. Plasmid DNA (grey, left) and polycationic polymers (black, left) or lipids (not shown) electrostatically self-assemble to form multicomponent complexes (right) containing multiple plasmids and thousands of polymers. Complex formation is entropically driven by the release of bound water molecules from the polyions.

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in the literature and in current clinical trials [7]. Viral vehicles have been created from viruses, including retrovirus, lentivirus, adenovirus, adenoassociated virus, vaccinia virus, and herpes simplex virus, and are designed to be replication incompetent by replacement of appropriate viral genes with the therapeutic transgene. Unfortunately, viral vehicles have faced numerous real and perceived obstacles in clinical development including their inherent toxicity and immunogenicity and concerns that they may revert to an infective form. Viruses are also limited in terms of their transgene carrying capacity by the size of the viral capsid and, in many cases, must have their native cell-specificity reprogrammed for the appropriate therapeutic target cell or tissue. 12.2.2 Nonviral Gene Delivery Vehicles Synthetic vehicles have received significant interest for their potential as safer and more controllable gene carriers than their viral cousins. These vehicles are formed by electrostatic interactions between cationic residues on the lipid or polymer carrier and negatively charged phosphate anions along the backbone of DNA (Figure 12.1). Complexation occurs spontaneously, and is an entropically driven phenomenon: interactions between the cations and anions release bound water molecules into the surrounding medium [11]. If sufficient polycation is added, the hydrophobicity of the partially neutralized DNA becomes so high that it loses its extended conformation and is forced to collapse [12]. The process compacts DNA and results in the formation of kinetically trapped spherical or toroidal complexes of roughly ten to several hundred nanometers in diameter [13, 14]. The formed complexes tend to be relatively polydisperse, with sizes and morphologies that depend on the conditions under which they are formed (i.e., mixing, order of addition) [15]. Lipoplexes and polyplexes typically contain several plasmids (or several thousand small RNA molecules) and several thousand lipids or polymers [16]. DNA complexation serves several purposes with respect to gene delivery. The DNA is condensed and compacted, which protects it from shear-induced degradation and sterically prohibits nuclease access to the DNA. The condensation process also imparts a level of control on the overall sizes of the complexes formed. The average size of these complexes depends on factors including the solution ionic strength and the formulation charge ratio [15, 17, 18] (for amine-containing polycations, this is defined as the ratio of amine nitrogens, N, to DNA phosphates, P) and can be tuned, to some extent, to promote more efficient extra- and intracellular transport. Furthermore, because complexes are typically formulated with an excess of the polycation (i.e., at N : P > 1), complexation enhances cellular internalization of DNA by promoting electrostatic interactions between the positively charged complexes and negatively charged glycosaminoglycan residues displayed on the extracellular surfaces of cells. Finally, the condensing polymers or lipids provide a scaffold onto which a variety of ligands can be attached, with the goal of enhancing the in vivo properties or appropriate trafficking of the complexes.

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For example, cell-binding and other targeting ligands have been grafted to free amine residues within the vehicle to stimulate vehicle uptake and appropriate subcellular trafficking by target cells.

12.3

VEHICLE TRANSPORT BARRIERS

Despite promising developments in vitro, nonviral vehicles have had limited success in vivo and have rarely reached clinical application. The relatively poor in vivo gene transfer efficiency of this class of vehicles has been attributed to a variety of factors, including their inadequate tissue and cellular targeting and poorly controlled intracellular trafficking and processing. One significant and understudied issue is the lack of control over the timing and location of DNA release. Polyplexes and lipoplexes can be prematurely unpackaged by exposure to extracellular components such as heparan sulfate proteoglycans (HSPGs), leading to DNA degradation by serum nucleases [19–25]. Inefficient unpackaging is also problematic, as condensed DNA has a limited ability to interact with the transcriptional machinery [26]. The following subsections will highlight major barriers in the extra- and intracellular gene delivery pathways and will briefly review material design strategies that address these barriers. The subsequent section will include a detailed discussion of the factors that determine polyplex/lipoplex stability and unpackaging, as well as new material-based strategies for programming site-targeted DNA release and activation. 12.3.1 Serum Stability and Extracellular Transport The gene delivery pathway presents formidable challenges to the delivery vehicle immediately following the introduction of the vehicle into the extracellular environment (Figure 12.2A). For in vivo gene transfer via systemic administration, the initial challenge to the vehicle is survival in the bloodstream. In this environment, the half-life of a vehicle depends on its ability to avoid aggregation and protein adsorption following contact with serum components. Charged particles like polyplexes are destabilized by electrostatic screening in the 150 mM physiological saline environment, which can cause their rapid aggregation, reduced activity, and significant adverse consequences to the host such as particle embolization in the lung [8]. Interactions between polyplexes and lipoplexes and serum proteins have been shown to initiate rapid vehicle elimination by phagocytic cells and the reticuloendothelial system [22, 27, 28]. To avoid these issues, poly(ethylene glycol) (PEG) and other hydrophilic and uncharged polymers, proteins, or oligosaccharides are frequently grafted as a brush onto free amine residues within the condensing polycation to provide a steric barrier against salt-induced aggregation [22, 27, 29–31]. These molecules can also prevent protein- or complement-induced inactivation [29, 32–34].

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(A)

(B)

Figure 12.2. Barriers to DNA transport. (A) Extracellular barriers. Poly- or lipoplex vehicles (black circles) travel through the host vasculature following administration, and must then extravasate (1) from the vasculature within the target tissue. Following extravasation, the vehicles must traverse the ECM containing collagen, proteoglycans, GAGs, and other ECM proteins to reach the target cell (2a). In some cases, interactions between the vehicle and the ECM can cause vehicle unpackaging and lead to the subsequent degradation of the DNA (2b). Inset image shown in (B). (B) Intracellular barriers. Vehicles enter cells by receptor-mediated endocytosis (3a) or macropinocytosis (3b). Subsequently, the early endocytic vesicles containing the vehicles are either recycled (4a) or mature into late endosomes (4b). Vehicles trapped within late endosomes might escape (5a) or be retained during endolysosomal maturation (5b); retention typically leads to DNA degradation. Released vehicles must travel through the cytosol to the vicinity of the nucleus (5a). Vehicles might enter the nucleus intact (6a) and subsequently unpackage (7a); alternatively, the vehicles might unpackage in the cytosol (6b), and the unpackaged DNA might enter the nucleus (7b) or be degraded by cytosolic nucleases (7c). (Author thanks John D. Larsen for assistance with figure preparation.)

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Provided the vehicle maintains sufficient serum stability, it must then begin the process of navigation to its target cell, which for nonhematopoietic and nonendothelial targets requires vascular extravasation and transport through the interstitial extracellular matrix (ECM) and/or lymphatics. As a general rule, methods to predictably control or promote vascular extravasation are not well established, given the low capillary permeability in most organs. For select tissues and organs, passive targeting strategies have been developed that take advantage of the unique physical properties of the target tissue. For example, nanoparticles such as polyplexes and lipoplexes tend to passively accumulate in the liver due to phagocytic clearance by the reticuloendothelial system [35–38]. A similar passive accumulation effect has been observed as a result of the aberrant architecture and increased vascular permeability and hydraulic conductivity of tumor endothelium [39–42]. This occurrence of tumor-specific accumulation, termed the enhanced permeability and retention (EPR) effect [43], is strongly size and molecular weight dependent. The average vascular pore sizes in various human and rodent tumors have been estimated as ∼400–500 nm in diameter [40], and macromolecules with sufficient serum stability and molecular weights greater than ∼50 kDa accumulate in tumor tissues. Transport of polyplexes and lipoplexes through dense ECM is mediated by a combination of convection and diffusion and depends on both their size and surface chemistry [44–46]. For example, in vitro studies employing hydrogels and multicellular spheroids have estimated the influence of nanoparticle characteristics on the efficiency of transport through simplified ECM. These studies suggest that particles with diameters of ∼100 nm or less can penetrate model matrices, but that penetration was relatively limited even for relatively small particles (∼20 nm) [47–49]. The surface chemistry of the nanoparticles was also critical. For example, PEGylated nanoparticles were found to move more rapidly through model ECM than unPEGylated nanoparticles, an effect attributed to reduced interactions between the PEGylated vehicles and fibrin networks within the ECM [50]. In another study, liposomes with neutral or low surface charges were found to penetrate to the central regions of spheroids, whereas charged liposomes were restricted [51]. A related issue with respect to the penetration of dense ECM by polyplexes and lipoplexes is the innate stability of the polyplexes and lipoplexes, and their tendency to unpackage and release DNA following contact with ECM components. This issue is covered in more detail in the following section on vehicle unpackaging. 12.3.2 Cellular Internalization and Intracellular Transport Upon arrival at their target cell, lipoplexes and polyplexes must navigate cellular entry and subcellular trafficking, ideally resulting in the delivery of intact cargo DNA to the nucleus (Figure 12.2B). Gene delivery vehicles typically enter the cell by either nonspecific macropinocytosis or receptor-mediated endocytosis, depending on their surface functionalization and size. Positively

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charged lipoplexes and polyplexes are generally internalized via macropinocytosis as a result of their nonspecific electrostatic interactions with the cell surface, as noted previously. Neutral or negatively charged complexes containing cell-binding ligands typically traffic through the relevant receptor-mediated pathway. The size of a lipoplex or polyplex is an important determinant of its internalization rate, and the optimal size for endocytosis has been investigated for various receptor-mediated pathways. As a general rule, previous experimental and computational studies have suggested that particles with diameters <50 nm are optimal for receptor-mediated endocytosis [52–57]. The mechanism of cellular entry largely determines the trafficking pathway employed by gene delivery vehicles. The majority of vehicles are thought to traffic into late endosomes, which acidify and can mature into acidic (pH ∼4–5) lysosomes containing a variety of lysosomal proteases and degradative enzymes [8]. Thus the endosomal membrane represents an important gene delivery barrier, and endosomal release prior to the acidification process is considered to be critical for effective gene delivery. A variety of strategies have been explored to mediate escape from the endosome into the cytoplasm, including the use of “proton-sponge” polymers such as poly(ethylenimine) (PEI) as DNA condensing agents, and the incorporation of pH-sensitive membranolytic polymers and peptides onto the delivery vehicle. Branched PEI contains a large number of secondary and tertiary amines, whose ability to buffer the endosomal acidification process has been proposed to result in osmotic swelling and rupture of the endosome [58–61]. pH-sensitive agents have been shown to enhance endosomal escape as a result of their structural transitions as acidic pH; for example, peptides that adopt an amphiphilic alpha-helical conformation at acidic pH can disrupt the endosomal membrane and promote the subsequent escape of encapsulated nanoparticles into the cytosol [62]. Once released into the cytosol, gene delivery vehicles must be transported to and into the nucleus for processing. Cytosolic transport represents a recently recognized barrier for the gene delivery process: cytosolic mobility is highly size dependent, restricting the free diffusion of DNA > 2000 bp and that of nanoscale particles [63–65]. Microtubule-dependent active transport of PEI polyplexes has been reported for a fraction of the internalized polyplexes (∼17%) [64]. Recently, anionic plasmid carriers formed from poly(lactic-coglycolic acid) and from 1,2-dioleoyl-sn-glycero-3-phosphocholine liposomes were conjugated to actin comet tails and shown to move through cytoplasmic extracts much more rapidly than their unconjugated counterparts [66]. Polyplex redistribution has also been proposed as a result of the mixing that occurs during mitosis [8]. The final intracellular transport barrier for gene delivery is the nuclear membrane. Transport into the nucleus is mediated by the nuclear pore complex, a multiprotein transmembrane assembly that permits the passage of small molecules but prohibits the free diffusive transport of macromolecules larger than 10–20 kDa [8]. The upper limit for active intranuclear transfer of proteins has been estimated as ∼50 kDa [67, 68], making it nearly prohibitive

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for DNA and nanoscale particles. Thus the majority of gene delivery vehicles are thought to accumulate in the nucleus during cell division [69], a theory that is supported by the findings that transfection of nondividing cells occurs at only very low levels, and that transfection of dividing cells is 30- to 500-fold more efficient when transfection is performed immediately prior to cell division as opposed to at the beginning of the cell cycle [8, 70].

12.4

VEHICLE UNPACKAGING

The final critical step in the gene delivery pathway for lipoplexes and polyplexes is the release of DNA from the complex, given that condensed DNA does not allow efficient access of the transcriptional machinery to the underlying DNA sequences [26]. This self-disassembly, or self-unpackaging, presents an important challenge in materials design, in that the same beneficial characteristics (i.e., a high density of positive charge [71–73]) that promote tight association of a polycation with DNA during gene transport eventually inhibit the activity of the DNA within the nucleus. A change in complex packaging state requires a change in the underlying structure or charge state of the condensing polycation. Unfortunately, there is a lack of obvious intracellular triggers (i.e., ionic strength, temperature, or pH) to stimulate this transition, given that the local pH drop in the endosome does not deprotonate the cationic amine residues that constitute the basis for many condensing agents. Indeed, the stability of various polyplexes to intracellular dissociation has been noted [26, 64, 74]. Inefficient polyplex disassembly has been identified as a critical rate-limiting barrier for gene delivery [26, 64, 74–78]. While unpackaging is a clear requirement for efficient transcription, the desired time and place for DNA release are less clear [71]. For example, premature unpackaging can be as problematic as inefficient unpackaging: interactions between lipoplexes or polyplexes and various extracellular components have been shown to cause destabilization, DNA release, and DNA degradation by serum nucleases. Intracellular components have also been shown to mediate lipo- or polyplex destabilization and DNA release, although in many cases, higher levels of intracellular release are correlated with improved transfection, despite the presence of cytoplasmic nucleases. Thus DNA packaging materials face opposing requirements, as they must protect DNA by tight packaging during transport, and release DNA prior to transcription. Materials designed with mechanisms to initiate the site-specific release of DNA in the vicinity of the cell’s nucleus would be expected to significantly enhance the efficient utilization of administered DNA. It is worth note that despite the development of numerous nonviral methods for achieving high levels of in vitro cell transfection, including those that employ lipoplexes and polyplexes, no method exists that could truly be considered efficient. For example, for a typical in vitro Lipofectamine-mediated transfection (Lipofectamine is a product of Invitrogen Corporation, Carlsbad, CA), millions of copies of

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plasmid DNA must be administered per cell (this assumes that 10 μg of plasmid DNA would be used to transfect 106 cells). Given that the accumulation of uncomplexed intranuclear DNA appears to very minimal, even when the levels of overall intracellular lipo- or polyplex accumulation are high [26, 64], it is clear that intracellular trafficking and unpackaging remain critical and unsolved barriers with respect to gene delivery. In this section, the parameters determining the innate stability of lipo- and polyplexes will be addressed, and known mechanisms that promote the unpackaging of these complexes in the extra- and intracellular environments will be explored. Subsequently, several classes of new materials that have been designed for the explicit purpose of site-specific unpackaging and DNA release within the cell will be reviewed, with emphasis on identifying the most promising new directions for further development. 12.4.1 Complex Structure and Stability Both lipids and polymers have demonstrated the capacity to compact and deliver DNA payloads in vitro. Polycation structure plays an important role in the efficiency of DNA binding and condensation. For example, branched PEI has been shown to more efficiently compact DNA then linear PEI [79], perhaps as a result of the higher density of primary amine groups on branched PEI relative to linear PEI. In general, primary amine groups within cationic lipids and polymers have been found to compact DNA more efficiently than higher order amines within corresponding molecules, presumably due to the reduced steric interference around the primary amine groups [80, 81]. The number and density of cationic groups on the polycation have also been shown to affect condensation efficiency. With polypeptide structures, several groups have found that a minimum of six to eight charged residues per polypeptide were necessary for tight polyplex packaging (i.e., packaging at a level that would reduce ethidium bromide staining of the complexed DNA) [26, 82, 83]. Different structural requirements for packaging were identified by these same groups, where Plank and co-workers found that the inclusion of tryptophan residues did not affect DNA binding affinity but reduced DNA compaction [82], and Wadhwa and co-workers found that the inclusion of tryptophan increased DNA binding efficiency [83]. Differences in the structures (branched vs. linear) and charge density of the studied polypeptides could underlie these different findings. Correlations between polycation length and packaging efficiency have also been identified for chitosan-based carriers [84, 85]. Furthermore, other work to systematically address the impact of various characteristics of polycationic polymers on DNA binding and condensation suggested that polymers with a low density of cationic residues were unable to efficiently compact DNA [86]. Random copolymers formed from neutral monomers and monomers containing a single quaternary amine residue bound to DNA inefficiently when the ratio of neutral monomer was 50% or greater [86].

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The functionalization of lipoplexes and polyplexes with stabilizing or targeting groups has also been shown to influence the efficiency of DNA compaction. For example, hydrophilic polymers such as PEG are often grafted to DNA condensing agents via reaction with primary amine groups along the backbone of the polycation [29, 31, 32, 87]. When a polycation is functionalized with PEG or another hydrophilic polymer prior to complexation, the charge density of the agent is reduced and DNA compaction is adversely affected [28, 88–91]. Furthermore, despite the fact that this surface grafting is typically performed for the purpose of reducing polyplex self-interactions and polyplex interactions with serum components [92], efficiently compacted polyplexes grafted with hydrophilic polymers have been found to be less stable to competing ions in blood than their ungrafted counterparts [19, 24, 32, 93, 94]. This effect has been attributed to thermodynamic destabilization caused by hydrophilic chains dispersed within the polyplex [24, 32]. However, even when grafting is performed on surface-exposed free amines postcomplexation, grafted polyplexes are often less stable to competitive ions [19, 23], and PEGylated polyplexes formed from lower molecular weight polycations can be destabilized by the addition of salt [28]. 12.4.2 Extracellular Unpackaging Depending on the innate stability of a lipo- or polyplex, a variety of components within the extracellular environment can drive premature unpackaging prior to cellular entry. Immediately following introduction into the bloodstream, PEG–PEI polyplexes have been shown to dissociate significantly [32, 93]. Polypeptide–DNA complexes have also been shown to dissociate within minutes upon introduction into the blood, allowing the degradation of DNA by serum endonucleases; when these same peptide-based polyplexes were crosslinked with glutaraldehyde prior to in vivo administration, their in vivo half-life was improved significantly [95]. Serum-induced destabilization may result from the enhanced electrostatic screening of DNA–polycation interactions at physiological salt levels. Alternatively, charged serum components such as albumin may destabilize these polyplexes by competitive DNA or polycation displacement. However, despite the fact that negatively charged model polyanions have been shown to displace DNA from polyplexes, albumin has been shown to bind to N-(2-hydroxypropyl)-methacrylamide/ 2-(trimethylammonio)ethyl methacrylate polyplexes without significant polyplex destabilization and only minor increases in the overall polyplex size [24]. Thus albumin-mediated polyplex clearance has been proposed to result from enhanced interactions between albumin-bound polyplexes and serum opsonins, as opposed to polyplex dissociation [24]. Following vascular extravasation, interactions with ECM proteins can also cause rapid dissociation of lipo- and polyplexes and the subsequent degradation of the DNA. For example, extracellular glycosaminoglycans (GAGs) such as heparin have been shown to cause polyplex destabilization/unpackaging,

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resulting in the degradation of the freed DNA by extracellular nucleases [19–21, 96, 97]. Burke and Pun demonstrated that both soluble and insoluble ECM GAGs and proteoglycans facilitated polyplex unpackaging in a fashion that correlated with the density of negatively charged residues, supporting the hypothesis of DNA displacement by competing anions in the ECM [19]. 12.4.3 Intracellular Unpackaging Interactions between lipo- and polyplexes and intracellular anions have also been proposed to stimulate polyplex unpackaging. For example, several groups have demonstrated the release of plasmid DNA from DNA-condensing lipids and polymers by cytosolic components [98–102]. In the case of lipoplexes, anionic HSPGs on the cell surface have been shown to interact with polycationic lipids within the lipoplex to initiate receptor-mediated endocytosis [103–106]. These interactions have been proposed to destabilize the endosomal membrane, cause the membrane lipids to flip to the cytoplasmic face of the membrane, and displace the DNA from the lipoplex into the cytoplasm [100, 102]. Polyplexes have been proposed to unpackage in the cytoplasm as a result of displacing interactions with other cytoplasmic polyanions. For example, Okuda et al. [99] demonstrated that the cytoplasmic component causing the release of plasmid DNA from dendritic poly(l-lysine) (PLL) and jetPEI was protease sensitive, as proteinase K-treated cytosolic fractions were inactive at DNA release. This effect was only observed at low PEI : DNA ratios, prompting speculation that other factors such as cytoplasmic RNA might be involved in cytosolic DNA displacement at high polycation : DNA ratios [101]. Huth et al. [101] demonstrated that the release of plasmid DNA from linear and branched PEI and PLL in cytoplasmic extracts was RNA-dependent, as RNase-treated cytoplasmic fractions did not release DNA. Furthermore, these authors demonstrated that in the absence of cytoplasmic extracts, the addition of RNA at levels similar to intracellular levels of RNA also resulted in DNA release from polyplexes formed at high polycation : DNA ratios. Chromosomal DNA has also been suggested to play a role in the displacement and release of DNA from lipo- and polyplexes [26, 107]. For example, fluorescence imaging analyses with PLL–DNA polyplexes demonstrated that a fraction of the polyplexes reached the nucleus, and that the amount of intranuclear polyplex dissociation was correlated with the length of the PLL chain [26]. Whereas long PLL chains (180 residues) remained closely associated with plasmid DNA, shorter PLL chains (19 and 36 residues) were found to be completely separated from the plasmid within the nucleus. In subsequent experiments, Schaffer et al. [26] demonstrated that double-stranded DNA could more easily dissociate the shorter PLLs from the plasmid than the longer PLL, a result consistent with cation exchange to the surrounding chromatin. Other investigators have identified a more minor role for chromosomal DNA in unpackaging. For example, Zabner et al. [103] demonstrated that lipoplexes

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injected into the nucleus showed low transfection efficiency, implying that chromosomal DNA plays only a small role in unpackaging [102]. Together, these experiments suggest that polycation structure and the innate stability of the lipo- or polyplex are important determinants of the site, mechanism, and extent of unpackaging, and support roles for multiple components in the unpackaging process. Although lipo- and polyplex materials capable of intracellular DNA release often have high in vitro gene transfer activities in comparison with their more stably packaged counterparts [26, 58, 84, 98, 108–112], cytoplasmic release has also been suggested to correlate with lower gene transfer activity [113]. For example, Oupicky et al. [113] demonstrated that cytoplasmic microinjection of free DNA resulted in lower levels of transfection than cytoplasmic microinjection of polyplexes, whereas nuclear microinjection of the same samples had the opposite effect. These findings were attributed in part to the enhanced resistance of the polyplex-contained DNA to cytoplasmic nucleases, although the authors also noted that the polyplexes might have improved cytoplasmic mobility and transport to the nucleus in comparison with the free DNA. In general, it is probable that cytoplasmic degradation of at least a portion of the released DNA occurs even with materials that exhibit high in vitro transfection efficiencies. Presumably, only a portion of the released DNA is sacrificed, and the remaining intact free DNA is utilized relatively efficiently.

12.5

SELF-UNPACKAGING MATERIALS

Several strategies have been proposed with respect to the design of materials capable of packaging and releasing DNA for efficient in vivo gene delivery. For example, the utilization of materials with an intermediate level of packaging has been suggested; ideally, these materials would be sensitive to the existing intracellular release mechanisms but still stable in serum and during contact with the ECM. However, given the similar structures and charge densities of the extracellular and intracellular GAGs and other anions that have been shown to initiate complex unpackaging, it would seem difficult to strike such a balance. Thus a variety of materials containing active DNA-releasing strategies have recently been explored. These include materials that release DNA by carrier degradation, carrier charge reduction, and crosslinker degradation (Figure 12.3). These major categories of active materials will be explored in the following subsections. 12.5.1 Release by Carrier Degradation Given that a correlation between the length of a polycationic carrier and its tendency to dissociate from DNA has been identified by several investigators [26, 82–85], as discussed above, one strategy that has been pursued for targeted unpackaging involves the use of high molecular weight degradable polycations

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(A)

(B)

(C)

Figure 12.3. Strategies for DNA release by active self-unpackaging. DNA within lipoor polyplexes can be released by hydrolytic or reductive degradation of the cationic carrier (A), charge reversal of the carrier (B), or degradation of surface crosslinkers on the carrier (C). Strategy (C) is often combined with carriers that degrade as in (A).

(Figure 12.3A). These species are typically either hydrolytically [109, 114–120] or reductively [75, 77, 110, 121] degradable, and thus are designed to gradually release DNA as they transition into low molecular weight (and fast-dissociating) species following exposure to aqueous media or reducing environments such as the endosome [108]. For example, Langer and co-workers have synthesized libraries of poly(β-aminoesters) and related biodegradable materials that condense DNA at physiological pH, but are hydrolytically degradable into nontoxic fragments [114, 115, 119, 120]. These investigators identified several candidate polymers with high in vitro gene transfection efficiencies. Hydrolytically degradable PEI derivatives have also been created and shown to have DNA-binding properties similar to high molecular weight (25-kDa) PEI with comparably low cytotoxicity and high gene transfer efficiency [109]. Forrest et al. [109] speculated that enhanced DNA release might be one determinant of the enhanced in vitro transfection efficiency of these materials. Jain and co-workers have synthesized pH-sensitive linear poly(amido amines)

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whose hydrolysis rate is significantly enhanced at pH values correspondent to the lysosomal pH [116]. Because of their capacity to selectively degrade in the intracellular environment, as opposed to in a sustained fashion, these new materials would be expected to have desirable properties for targeted DNA release. With a similar goal of targeted intracellular degradation and DNA release, reductively degradable DNA-packaging materials have also been synthesized. For example, Pichon et al. [77] synthesized high molecular weight PLL polymers for gene delivery by the reductive crosslinking of low molecular weight PLL via disulfide linkages. In a similar strategy, Gosselin et al. [75] created crosslinked PEI polymers from low molecular weight PEI linked by disulfide bonds. These reductively linked polymers had high in vitro gene transfer efficiencies relative to the corresponding uncrosslinked low molecular weight species, and were shown to release DNA upon incubation with reducing agents such as glutathione [75, 77]. More recently, reductively degradable poly(amido amines) and poly(amido ethylenimines) were synthesized and shown to exhibit high levels of in vitro gene transfection in the presence of serum, even when serum incubation times were extended for several hours [110, 121]. These results suggest that the intracellular targeted release mechanism employed by these materials had good potential for in vivo activity. 12.5.2 Release by Carrier Charge Reduction Other strategies for the targeted release of DNA have also been pursued, including the use of polycationic carriers designed to undergo controlled reductions in their net charge when exposed to physiologically relevant environments [71, 108, 111, 112, 122–125] (Figure 12.3B). For example, Hennink and co-workers have created charge-reducing polycations from poly(methacrylates) whose amine-containing side chains are attached to the polymer backbone via hydrolyzable carbonate bonds; thus hydrolytic cleavage of these polymers results in the gradual loss of cationic amine groups [123–125]. These polymers were shown to release DNA in aqueous media in a fashion correlated with the expected time-dependent changes in electrostatic interactions, and mediated high levels of in vitro cell transfection. Others have also created materials whose positive character is reduced by hydrolysis. For example, Lynn and co-workers have created charge-shifting cationic polymers whose hydrolysis results not only in the loss of cationic groups, but also in the introduction of anionic residues [71, 108, 112]. The DNA-releasing capacities of these materials have been demonstrated for surface-mediated [25] and polyplex-mediated [71, 108] delivery. Prata et al. [111] have designed chargereversal amphiphiles that transition from positively to negatively charged within cells. These amphiphiles bind to DNA to form supramolecular lipoplex complexes, but are cleaved by endosomal esterases to become anionic and release DNA. The supramolecular lipoplexes had high in vitro activity that was dependent upon esterase-mediated lipid cleavage.

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12.5.3 Release by Crosslinker Degradation Although the above approaches for targeted release are promising, the lack of evaluation (to date) of these systems in vivo combined with the increasing concerns about the stability of lipo- and polyplexes during prolonged exposure to the extracellular environment have led to recent interest in new methods to reversibly stabilize polyplexes. For example, several approaches have focused on the development of methods to covalently crosslink the surfaces of polyplexes with degradable linkages [76, 126] (Figure 12.3C). Oupicky et al. [76] explored the potential of polyplexes formed from reducible polycations and stabilized by surface crosslinking with hydrophilic polymers. Surface coating was shown to suppress the unpackaging of the polyplexes by poly(acrylic acid)-mediated displacement. The stabilized particles were capable of moderate levels of cellular transfection when cell-binding ligands were conjugated to their surfaces. In another approach, PEI–DNA polyplexes were crosslinked with disulfide bonds [126]. These crosslinked polyplexes were stable against dissociation by polyanions and high ionic strength and had improved biocompatibility against albumin and erythrocyte interactions in comparison with uncrosslinked PEI–DNA complexes, suggesting the potential applicability for systemic application. 12.6

CONCLUSIONS

Gene delivery represents the most significant hurdle to the realization of nucleic acid-based therapies, and despite years of effort directed at the design of new nonviral delivery materials, successes have been limited. Many pervasive problems with these materials are related to the lack of correlation between their in vitro and in vivo performance. With respect to lipo- and polyplex-mediated delivery, the ability to protect DNA within the extracellular environment but release it within the cell has presented a particular challenge. Many new materials are now being developed with promising potential for the targeted intracellular release of DNA. These materials offer possible solutions to a significant problem in gene delivery, and improved understanding in this area may enable the synthesis of nonviral materials with efficacies that rival those of viral delivery systems, without the related concerns of toxicity and immunogenicity. The demonstration of these new releasing materials in vivo will improve understanding of the mechanisms that control gene delivery and will pave the way for their clinical translation. REFERENCES 1. Mulligan, R. C. The basic science of gene therapy. Science 260: 926–932 (1993). 2. Huang, H., Yee, J., Shew, J., Chen, P., Bookstein, R., Friedmann, T., Lee, E., and Lee, W. Suppression of the neoplastic phenotype by replacement of the RB gene in human cancer cells. Science 242: 1563–1566 (1988).

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105. Legendre, J.-Y. and Szoka, F. C. Jr. Delivery of plasmid DNA into mammalian cell lines using pH-sensitive liposomes: comparison with cationic liposomes. Pharm. Res. 9(10): 1235–1242 (1992). 106. Felgner, P. L., Gadek, T. R., Holm, M., Roman, R., Chan, H. W., Wenz, M., Northrop, J. P., Ringold, G. M., and Danielsen, M. Lipofection: a highly efficient, lipidmediated DNA-transfection procedure. PNAS 84: 7413–7417 (1987). 107. Remy, J. S., Kichler, A., Mordvinov, V., Schuber, F., and Behr, J. P. Targeted gene transfer into hepatoma cells with lipopolyamine-condensed DNA particles presenting galactose ligands: a stage toward artificial viruses. PNAS 92: 1744–1748 (1995). 108. Liu, X., Yang, J. W., and Lynn, D. M. Addition of “charge-shifting” side chains to linear poly(ethyleneimine) enhances cell transfection efficiency. Biomacromolecules 9: 2063–2071 (2008). 109. Forrest, M. L., Koerber, J. T., and Pack, D. W. A degradable polyethylenimine derivative with low toxicity for highly efficient gene delivery. Bioconjug. Chem. 14(5): 934–940 (2003). 110. Lin, C., Zhong, Z., Lok, M. C., Jiang, X., Hennink, W. E., Feijen, J., and Engbersen, J. F. J. Novel bioreducible poly(amido amine)s for highly efficient gene delivery. Bioconjug. Chem. 18(1): 138–145 (2007). 111. Prata, C. A. H., Zhao, Y., Barthelemy, P., Li, Y., Luo, D., McIntosh, T. J., Lee, S. J., and Grinstaff, M. W. Charge-reversal amphiphiles for gene delivery. J. Am. Chem. Soc. 126(39): 12196–12197 (2004). 112. Zhang, J. and Lynn, D. M. Ultrathin multilayered films assembled from chargeshifting cationic polymers: extended, long-term release of plasmid DNA from surfaces. Adv. Mater. 19(23): 4218–4223 (2007). 113. Oupicky, D., Konák, C., Ulbrich, K., Wolfert, M. A., and Seymour, L. W. DNA delivery systems based on complexes of DNA with synthetic polycations and their copolymers. J. Control. Release 65(1–2): 149–171 (2000). 114. Akinc, A., Lynn, D. M., Anderson, D. G., and Langer, R. Parallel synthesis and biophysical characterization of a degradable polymer library for gene delivery. J. Am. Chem. Soc. 125(18): 5316–5323 (2003). 115. Anderson, D. G., Lynn, D. M., and Langer, R. Semi-automated synthesis and screening of a large library of degradable cationic polymers for gene delivery. Angew. Chem. Int. Ed. 42(27): 3153–3158 (2003). 116. Jain, R., Standley, S. M., and Frechet, J. M. J. Synthesis and degradation of pHsensitive linear poly(amidoamine)s. Macromolecules 40(3): 452–457 (2007). 117. Lim, Y.-B., Kim, S.-M., Lee, Y., Lee, W.-K., Yang, T.-G., Lee, M.-J., Suh, H., and Park, J.-S. Cationic hyperbranched poly(amino ester): a novel class of DNA condensing molecule with cationic surface, biodegradable three-dimensional structure, and tertiary amine groups in the interior. J. Am. Chem. Soc. 123(10): 2460–2461 (2001). 118. Lim, Y.-B., Kim, S.-M., Suh, H., and Park, J.-S. Biodegradable, endosome disruptive, and cationic network-type polymer as a highly efficient and nontoxic gene delivery carrier. Bioconjug. Chem. 13(5): 952–957 (2002). 119. Lynn, D. M., Anderson, D. G., Putnam, D., and Langer, R. Accelerated discovery of synthetic transfection vectors: parallel synthesis and screening of a degradable polymer library. J. Am Chem. Soc. 123(33): 8155–8156 (2001).

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120. Lynn, D. M. and Langer, R. Degradable poly(beta-amino esters): synthesis, characterization, and self-assembly with plasmid DNA. J. Am. Chem. Soc. 122(44): 10761–10768 (2000). 121. Christensen, L. V., Chang, C.-W., Kim, W. J., Kim, S. W., Zhong, Z., Lin, C., Engbersen, J. F. J., and Feijen, J. Reducible poly(amido ethylenimine)s designed for triggered intracellular gene delivery. Bioconjug. Chem. 17(5): 1233–1240 (2006). 122. Shim, M. S. and Kwon, Y. J. Controlled delivery of plasmid DNA and siRNA to intracellular targets using ketalized polyethylenimine. Biomacromolecules 9(2): 444–455 (2008). 123. Funhoff, A., van Nostrum, C., Janssen, A., Fens, M., Crommelin, D., and Hennink, W. Polymer side-chain degradation as a tool to control the destabilization of polyplexes. Pharm. Res. 21(1): 170–176 (2004). 124. Jiang, X., Lok, M. C., and Hennink, W. E. Degradable-brushed pHEMApDMAEMA synthesized via ATRP and click chemistry for gene delivery. Bioconjug. Chem. 18(6): 2077–2084 (2007). 125. Luten, J., Akeroyd, N., Funhoff, A., Lok, M. C., Talsma, H., and Hennink, W. E. Methacrylamide polymers with hydrolysis-sensitive cationic side groups as degradable gene carriers. Bioconjug. Chem. 17(4): 1077–1084 (2006). 126. Neu, M., Sitterberg, J., Bakowsky, U., and Kissel, T. Stabilized nanocarriers for plasmids based upon cross-linked poly(ethylene imine). Biomacromolecules 7(12): 3428–3438 (2006).

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CHAPTER 13

Cellular Trafficking of Dendrimers YUNUS EMRE KURTOGLU and RANGARAMANUJAM M. KANNAN Department of Chemical Engineering and Materials Science, NICHD Perinatology Research Branch, Wayne State University, Detroit, Michigan

13.1

DENDRITIC ARCHITECTURE

Dendrimers are polymeric molecules made of multiple monomers branching radially from a central core. The monomers used determine the dendrimer subfamilies, while the number of branching points between the core and the surface of dendrimers determine their generation number. The monomer used for the synthesis, generation number, and surface groups determines the chemical, physical, and biological characteristics. For biological applications various dendrimer structures have been synthesized, which include polyamidoamines [1], poly(aryl ethers) [2], polyamines [3], polyesters [4, 5], nucleic acids [6, 7], polypeptides [8], and carbohydrates [9]. These dendrimers have shown great promise in diverse biological applications such as gene and drug delivery, magnetic resonance imaging (MRI), antivirals, and antibacterials, as well as tissue scaffolds [10]. In gene and drug delivery applications, usually the aim is a combination of drug solubility enhancement, tissue targeting, increasing blood circulation time, reduction in drug/gene metabolism rate, or overcoming biological barriers. Gene delivery strategies often involve complexation of nucleic acids to positively charged dendrimers, whereas most small drug delivery approaches involve covalent attachment of active molecules to dendrimer surface groups. Alternatively, encapsulation of drug molecules to the interior space of a dendrimer has also been investigated. In general, dendrimers have lower polydispersity indices compared to linear polymers due to their stepwise synthesis schemes and highly symmetrical branching patterns. The low polydispersity index is reflected by welldefined number of surface functional groups as compared to hyperbranched Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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and linear polymers synthesized by statistically controlled polymerization reactions. As a result of the branching, higher generation dendrimers have more surface end groups and higher molecular weights. On the other hand, the surface groups get more densely packed as the generation numbers go up, which may increase the charge density on the surface depending on the end groups. Most dendrimers are not synthesized over generation 10 (G10), since the surface crowding does not leave any room for further branching and typical sizes for dendrimers vary between 2 and 10 nm. The lower generation dendrimers (up to G4) are generally ellipsoidal, whereas higher generations are more spherical in shape. Well-defined globular structure and the multivalency at their surfaces give dendritic structures some advantages over linear and hyperbranched polymers for drug delivery applications. While the well-defined structure can provide more uniform biodistribution and pharmacokinetics, the functional surface groups enable the attachment of multiple drugs, targeting ligands, and imaging agents. Due to their globular structures and surface charge densities the biological characteristics of dendrimers differ from linear polymers. This macromolecular architecture of dendrimers creates opportunities for cellular targeting and fine-tuning their uptake mechanisms and rates. The specific aim of this chapter is to discuss how the surface properties of these threedimensional (3D) globular polymers govern their cellular transport characteristics. Polyamidoamine (PAMAM) dendrimers have been most widely investigated among the other dendritic molecules perhaps due to their commercial availability and the various sizes and surface functional groups readily available. Therefore, in order to understand how the globular 3D structure and surface functionality affects the cellular uptake of dendritic structures, it is imperative to discuss how various types of PAMAM dendrimers and other dendrimers behave in a comparative manner. Mechanism and rate of cellular uptake on particular cell types and their fate after internalization are important considerations for drug delivery applications and these properties are closely related to surface characteristics. Cellular uptake of dendrimers is mainly affected by their size, the type and charge of surface groups, and the attachment of drugs or other moieties to their functional groups on the periphery. The mechanisms for transport of molecules through biological barriers include endocytosis, passive diffusion, and carrier-mediated and paracellular transport. Like other polymeric macromolecules, endocytotic mechanisms are responsible for cellular uptake of most dendrimers even though smaller and hydrophobic dendrimers may be able to enter the cells by diffusion through the cellular membrane [11]. Endocytosis is an active cellular uptake mechanism to internalize small and large macromolecules and particles. Two types of endocytotic processes have been defined: phagocytosis and pinocytosis. Phagocytosis involves the uptake of large particles and is a characteristic mechanism of white blood cells, whereas pinocytosis follows clathrin-coated pits, caveolae, or clathrin- and

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caveolae-independent pathways. The pathway by which a macromolecule is taken up can vary with the cell type and the proteins involved in the intracellular vesicles and size of these vesicles, as well as the intracellular fate of the macromolecule. Typically, after internalization polymers are transported to early endosomes followed by late endosomes and later in catalytic environment of lysosomes. The cellular uptake rates and residence times in intracellular compartments are important for designing dendrimer-based therapeutics since they often rely on release of covalently linked drugs intracellularly.

13.2 DENDRITIC VERSUS LINEAR AND HYPERBRANCHED POLYMERS The rate and mechanism of cellular uptake of G2, G3, and G4 amine terminated (cationic) PAMAM dendrimers were investigated in comparison to linear and branched polyethylene imine (PEI) polymers, which are also cationic with amine functional groups [12]. The polymers were labeled by Oregon green (through conjugation), and their cell membrane binding, endocytosis, and exocytosis characteristics were studied on B16F10 melanoma cells. While the cellular uptake mechanism of all the polymers was determined to be endocytosis, the hyperbranched PEI and the PAMAM dendrimers had cholesteroldependent pathways, whereas linear PEI uptake was cholesterol and clathrin independent. The rate of uptake was the highest for G4 PAMAM followed by branched PEI, linear PEI, and G3 and G2 PAMAM. The globular structure and the cationic surface charge density of G4 PAMAM is believed to enhance the cellular uptake by adsorptive endocytosis compared to lower generation dendrimers and PEI polymers. Particularly, the surface functional groups of the dendrimer can produce a highly localized charge density, which can have a significant influence on its interactions with the negatively charged proteoglycans in the cell membrane. All the polymers showed significant membrane binding, which suggests the cationic charge may give rise to nonspecific absorption. This type of adsorptive endocytosis usually follows curvilinear cell uptake kinetics due to saturation of the membrane binding sites. Cellular entry dynamics of amine-terminated G4 and G3 PAMAM dendrimers and hydroxyl-terminated PAMAM dendrimers were studied with human lung epithelial carcinoma cells (A549) in comparison to hyperbranched polyol polymer (Figure 13.1) [13]. Additionally, the amine-terminated G3 PAMAM dendrimer surface was modified with poly(ethylene glycol) PEG to investigate the PEG attachment effects on cellular uptake kinetics. Approximately 90% of amine-terminated PAMAM dendrimers (G3 and G4) entered the cells within 1 hour while the initial rate of cell entry of G4-NH2 was faster compared to that of G3-NH2 (Figure 13.2). In comparison, approximately 55% of G3 PEG and 75% of polyol were inside the cells in 1 hour. G4 PAMAM dendrimers were taken up by cells more rapidly when compared to G3 dendrimers with or without PEG modification and polyol. The higher cell

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234 H2N H2N H2N

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NH H2N O NH NH H2N O N O N NH N HO O N NH H2N N O NH O H2N HO N N O N NO H H NH N NO O H N NHO O NHNH2 N H2N O NH O O O O O H2N NH N O N NH N NH N H N NH N N H N NH O NHO O O NH OO O NH NH2 NH NH NH H2N O NH2 N N O N N O N NH H2N H O N O NH O NH NH O O NH O N H2N NH NH NH O NH H2N NH NH 2 N OO O 2 N NH O NH NH2 NH O NH H2N NH2 NH H2N NH2 H2N

PAMAM Dendrimer

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Figure 13.1. Schematic representation of the PAMAM dendrimer (left) and the polyol hyperbranched polymer (right). The “imperfect” branching pattern in the hyperbranched polymer can be seen [13]. 7/30/2010 2:19:13 PM

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235

% Change in absorbance

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Figure 13.2. Effect of end groups on cellular entry profile: (top) supernatant analysis, (bottom) cell lysate analysis [13]. A549 cells were treated separately with each of the dendrimers and hyperbranched polymer was tagged with fluorescent dye. The supernatant was removed at times 0, 30, 60, 120, 240, and 360 min. The amount of tagged dendrimer in the supernatant was estimated by UV/Vis absorbance at 492 nm. The cells were then washed with PBS, trypsinized, and centrifuged to obtain a cell pellet. The cell pellet was then lysed with cell lysate buffer and the amount of tagged dendrimer in the lysate was quantified. An increase in the fluorescent activity in the cell lysate along with a corresponding decrease in activity in the supernatant indicates the intracellular entry of the tagged dendrimer.

entry rate of amine-terminated G4 PAMAM compared to G3 PAMAM was explained by the larger amount of surface charges on the G4 dendrimer. The other polymers studied, namely, hydroxyl-terminated G4 PAMAM, polyol, and G3 PAMAM-PEG, do not carry cationic surface charges and their cellular uptake is associated with nonspecific adsorption to cell membrane. The hydroxyl-terminated G4 PAMAM entered the cells more rapidly compared to other uncharged polymers, suggesting that the globular dendrimer structure is contributing to the enhanced cellular uptake rates. PEG modification of amine-terminated G3 PAMAM reduced its cellular uptake rates due to reduced ionic interaction with the cell membrane and therefore reduced

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surface adsorption that would lead to endocytosis. When the noncationic polymers were compared for their cellular uptake rates, the dendritic structure enhances the cellular uptake compared to hyperbranched polyol. These studies must be viewed as somewhat qualitative and relative, since quantification would require more detailed work.

13.3

SURFACE CHARGE

G4 PAMAM dendrimers with 64 (calculated) amine (cationic), carboxyl (anionic), or hydroxyl (neutral) surface groups were studied for their cellular uptake mechanisms on A549 lung epithelial cells, which are known to have a negative surface charge [14]. The three types of PAMAM dendrimers were tagged with fluoroisothiocyanate (FITC) and their cellular uptake was investigated by flow cytometry and fluorescence microscopy analysis. The cellular uptake rates of the three dendrimers were analyzed in comparison to each other (Figure 13.3). Rate of cell uptake was highest for the cationic dendrimer, followed by anionic and neutral dendrimers. In order to identify specific uptake mechanisms, inhibitors of specific endocytotic pathways were used in each group. Regardless of their surface charge, all PAMAM dendrimers studied were endocytosed by energy-dependent, fluid-phase endocytosis while their specific endocytosis uptake mechanisms varied. Anionic dendrimers were partly taken up by caveolae, while the cationic and anionic dendrimers were 100

%Cell entry

80 60 40 COOH OH NH

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Figure 13.3. A549 cell entry profile of G4-NH2, G4-OH, and G3.5-COOH PAMAM dendrimers [14]. Rate of cell entry is determined as percent of (cell entry at time t/cell entry at 3 h). The cellular uptake was quantified by flow cytometry using fluorescence. A comparison of the time-dependent fluorescence intensity levels indicates that rate of cell uptake was highest for the cationic dendrimer, followed by anionic and neutral dendrimers. The cell uptake for cationic dendrimer plateaus off after 1 hour, unlike with the other two dendrimers where the cell entry increased more or less linearly with the treatment time, as shown in the graph. (See color insert.)

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SURFACE CHARGE

237

(A)

(B)

(C)

Figure 13.4. Confocal microscopic images (63×) of A549 lung epithelial cells after treatment with different dendrimers and LysoTracker for 5 min followed by chase for 30 min in serum-free medium devoid of any dendrimers. Images indicate cells treated with (A) FITC-labeled PAMAM-G4-NH2 dendrimer and LysoTracker. (B) FITClabeled PAMAM-G4-OH dendrimer and LysoTracker. (C) FITC-labeled PAMAMG3.5-COOH dendrimer and LysoTracker. The left-hand panel shows the green fluorescence from the dendrimer, the middle panel shows the red fluorescence from the LysoTracker, and the right panel shows the yellow fluorescence due to the colocalization of green dendrimer–FITC and red LysoTracker in the lysosomes. LysoTracker dye preferentially partitions to the lysosome, giving an intense red color in the acidic environment of the lysosomes. From the colocalization studies, it is evident that the dendrimers are in the lysosomes within 30 min of their transport from the cell membrane [14]. (See color insert.)

taken up by a non-clathrin- and non-caveolae-mediated endocytosis mechanism in A549 cells. Furthermore, the surface charge of PAMAM dendrimers seems to play a role in their intracellular trafficking. After cellular uptake, extent of cationic dendrimer that resides in the lysosomes was less than that of anionic and neutral dendrimers, whereas the cationic dendrimer was found in greater extent in peripheral vesicles (Figure 13.4). This finding is

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significantly important for dendrimer–drug conjugate therapies that rely on lysosome enzyme activity for detachment of drug from its carrier. The greater lysosomal residence time for the anionic and neutral dendrimer can provide longer release times, whereas cationic dendrimer conjugates may have less time in the lysosome compartment, thus requiring faster releasing covalent linkages. Therefore the surface charge of the dendrimers can be utilized to modulate the cell entry kinetics, mechanisms, and residence time in intracellular compartments.

13.4

SURFACE MODIFICATION

Human lung epithelial carcinoma cells were also used to investigate the efficacy of PAMAM dendrimer conjugates of ibuprofen (Ibu) [15] and methylprednisolone (MP) [16]. The cellular uptake along with their in vitro efficacy were reported. MP was conjugated to carboxyl terminated G2.5 PAMAM and hydroxyl-terminated G4-OH PAMAM dendrimers while the conjugates were also tagged with FITC. The conjugates were rapidly internalized within minutes, while the uptake continued for up to 4 hours. The localization of the conjugates was studied using fluorescence and confocal microscopy. The in vitro efficacy of MP-PAMAM conjugates was studied by inhibition of prostaglandin secretion. The efficacy of the conjugated MP was comparable to free MP treatment, suggesting that the dendritic delivery approach was successful. Even though the dendrimer may enhance the uptake of MP, the drug has to be released from the dendrimer for prostaglandin suppression. The fact that the conjugate shows comparable efficacy at short times (4 hours), when the drug release is expected to be very small, suggests that the delivery is significantly better with the dendrimer. Ibuprofen complexes of G4 PAMAM dendrimers were also studied for their cell entry. Compared to free Ibu the PAMAM complexes had higher cell entry rates, and the complexes were taken up to significant levels within 1 hour. When ibuprofen was attached to hydroxyl-terminated G4 PAMAM dendrimer and the cellular uptake rates were studied, it was determined that the conjugates entered the cells rapidly with ∼30% uptake in 15 minutes. The punctuated distribution of fluorescence in the cytoplasm determined by confocal and fluorescence microscopy images suggested that the cellular uptake was through endocytotic pathways. Intracellular trafficking of G3 PAMAM dendrimers with amine functionality in the human colon adenocarcinoma HT-29 cell line was also studied. PAMAM dendrimer surface was modified with lauroyl chains, propranolol molecules, or both among with FITC tagging [17]. Cellular uptake of unmodified and propranolol-modified G3 PAMAM dendrimers was determined to be by both caveolae-dependent endocytosis and macropinocytosis pathways, whereas when both surface modifications were present internalization of

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dendrimer was caveolae dependent, and possibly clathrin-dependent. Cell entry pathways of lauroyl chain-modified G3 PAMAM were caveolae dependent, clathrin dependent, and macropinocytosis. All the modified G3 PAMAM dendrimers were trafficked to endosomes and lysosomes after internalization. Modification of G3 PAMAM dendrimer with lauroyl moieties increased the cellular uptake rates. Furthermore, lauroyl modification of dendrimer surface reduced the extent of lysosomal accumulation. The impact of PEG modification on cellular uptake kinetics of carboxylterminated PAMAM dendrimers was also studied with Caco-2 cells with focus on transepithelial transport [18–20]. Cellular uptake and transport of G3.5 and G4.5 PAMAM dendrimers with three levels of PEG surface modification were studied. PEG modification of the surface significantly decreased the transepithelial transport for both generations, whereas the cellular uptake showed generation dependency. G3.5 dendrimer cellular uptake was largely reduced by attachment of PEG to the surface in contrast to a strong reduction in uptake for G4.5 PAMAM with PEG modification. The degree of PEG modification did not create a significant difference in cellular uptake of G3.5 dendrimers and regardless of surface groups modified the cellular uptake was decreased. On the other hand, for G4.5 PAMAM dendrimers the lowest extent of PEG surface modification showed the most improvement over the unmodified dendrimer while further PEG modification actually reduced the cellular uptake compared to less surface modification. The conformation of flexible PEG chains on the surface are believed to be determining parameters for the different characteristics for the two generations studied. The reduction in transepithelial permeability was about 65% for both G3.5 and G4.5 PAMAM dendrimers upon PEG modification as compared to permeability of unmodified G3.5 and G4.5 PAMAM dendrimers.

13.5

PERMEABILITY THROUGH BIOLOGICAL BARRIERS

For oral delivery of dendrimer drug conjugates, it is important to evaluate the permeability though epithelial barriers. For determination of absorption and cell entry characteristics human epithelial colorectal adenocarcinoma cells (Caco-2) are often employed [21, 22]. To demonstrate the influence of size and charge on transport of PAMAM dendrimers across Caco-2 cells, extensive studies were carried out using positive, neutral, and negative charged PAMAM dendrimers [23]. Permeability increased with an increase in the number of anionic surface groups, thus generation number in the carboxyl-terminated PAMAM dendrimers also increased. Cationic G2 PAMAM had greater permeability than hydroxyl-terminated G2 and carboxyl-terminated G1.5 and G2.5 dendrimers. Anionic dendrimer G3.5 PAMAM and cationic G4 PAMAM modified with FITC molecules showed the highest transport rates. Increase in the attachment of hydrophobic FITC label increased the permeability and

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reduced the toxicity of cationic dendrimers. The overall permeabilities of various PAMAM dendrimers through epithelial cells were determined as G3.5COOH > G2NH2 > G2.5COOH > G1.5COOH > G2OH. The enhanced PAMAM permeability was attributed to opening of tight junctions in cells via paracellular transport across the Caco-2 cells. Furthermore, depending on their surface properties, PAMAM dendrimers are transported across epithelial barriers to a higher extent than some of the conventional linear watersoluble polymers. Epithelial permeability of cationic dendrimers decreases with size and in contrast the permeability of anionic dendrimers increases with their size. The charged dendrimers had greater permeability than neutral dendrimers, which has no net surface charge at physiological pH to enable its interaction with cell monolayers and between the cationic and anionic, the cationic dendrimers exhibit higher permeability. Furthermore, the cationic dendrimers are transported by a combination of paracellular and endocytotic mechanisms. The trafficking of PAMAM dendrimers to endosomal and lysosomal compartments in Caco-2 cells is rapid and mediated by a clathrin-dependent endocytosis mechanism. This was confirmed from the uptake of amineterminated G4 PAMAM using endocytosis inhibitors such as brefeldin A, colchicine, filipin, and sucrose. In the presence of these inhibitors the uptake and permeability of the G4 PAMAM were significantly lowered [24, 25]. The anionic G4 PAMAM dendrimers are taken up by caveolae-mediated endocytosis in A549 lung epithelial cells and the cationic and neutral G4 PAMAM dendrimers are internalized by a non-clathrin-, non-caveolae-mediated mechanism involving electrostatic interactions or other nonspecific fluid-phase endocytosis. These studies revealed that PAMAM dendrimer internalization by endocytosis is largely dependent on the types of cells targeted in addition to the surface charge, molecular weight, and generation considerations. Therefore the differences in cell uptake mechanism of dendrimers may be cell type dependent. This dependency requires that dendritic devices may need to be tailored for each application and for ultimate cell type being targeted for optimal cellular uptake kinetics. In another study, 125I-labeled anionic G1.5 and G2.5 PAMAM dendrimers were transported across everted rat intestinal tissue much faster than other linear polymers such as polyvinylpyrrolidone (PVP), poly(N-vinylpyrollidoneco-maleic anhydride) (NVPMA), and N-(2-hydroxypropyl)methacrylamide (HPMA) copolymers [26]. While 80% of PAMAM radioactivity was transferred directly across to the serosal fluid, 20% of the anionic PAMAM dendrimer remained associated with the tissue. On the other hand, when the studies were carried out with G3 and G4 amine-terminated cationic PAMAM dendrimers, 60% of the dendrimer radioactivity was found in the tissue whereas 40% was transferred into the serosal fluid. This observation clearly demonstrates the increased interactions between the cationic dendrimers and the cell membranes.

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13.6

SIRNA

241

OLIGONUCLEOTIDE DELIVERY

siRNA therapeutics have generated a lot of interest in drug delivery research. However, their low resistance against enzymatic degradation, poor cellular uptake, and rapid renal and liver clearance have limited the in vivo applications. In order to address these issues, PAMAM dendrimers were investigated as carriers for antisense and siRNA oligonucleotides. Amine-terminated G5 PAMAM dendrimer was complexed with antisense and siRNA oligonucleotides using the cationic charge of the dendrimer surface and the negative charge of the oligonucleotides [27]. Dendrimer–oligonucleotide complexes were further modified by covalent attachment of a cell-penetrating peptide that is cationic in nature. BODIPY fluorescent molecules were attached to dendrimers for visualization, whereas Cy-5 fluorescent molecules were used for oligonucleotide tracking. The efficacies and the cellular uptake of the complexes were studied on NIH 3T3 fibroblast cells transfected with a plasmid containing the human MDR1 gene. G5 PAMAM–oligonucleotide complexes were effectively internalized and accumulated in intracellular vesicles as determined by confocal microscopy and flow cytometry. Conjugation of cellpenetrating peptide to the G5 PAMAM increased the cellular uptake by 25%, whereas this enhanced uptake did not improve the effectiveness of therapy, which was studied by inhibition of P-glycoprotein expression. G5 PAMAM delivery systems, both with and without peptide modification, were moderately effective in intracellular delivery of antisense oligonucleotide. As delivery vehicles, the effectiveness of dendrimers was comparable to Lipofectamine 2000 as a positive control. The confocal microscopy images suggested that both G5 PAMAM with and without peptide modification, complexed with either siRNA or antisense oligonucleotide, had similar intracellular distribution with most of the material associated with intracellular vesicles while a small portion of siRNA and antisense oligonucleotides were found in the cytoplasm and nucleus. Therefore amine-terminated G5 PAMAM was equally effective in carrying the oligonucleotides into the cells, but overall effectiveness varied due to the differences in siRNA and antisense nucleotide characteristics.

13.7 INTRACELLULAR TRAFFICKING AND EFFICACY Efficacy of the dendrimer–drug conjugates is usually dependent on the release of the active molecules from the dendritic carrier. The activity of small drugs are often suppressed or completely blocked in their conjugated form. It is important to design systems that will release their payload intracellularly at predefined rates. Intracellular trafficking characteristics of dendrimer-based delivery systems such as cellular uptake rates, lysosomal residence time, exocytosis rates, and ability to exit lysosomes to reach the cytoplasm determine the overall extent of drug release. The drug release rates of covalently linked

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dendrimer–drug conjugates vary with the intracellular conditions, such as availability and activity of the lysosomal enzymes and acidity of the intracellular compartment, along with time spent under these conditions. Therefore the intracellular activation of the drugs being delivered as well as the efficacy of the dendrimer conjugate system will be in direct relation with its intracellular trafficking properties. The effects of dendrimer surface functional groups on the efficacy of the dendrimer–drug conjugates were investigated with PAMAM dendrimer– methotrexate (MTX) conjugates [28]. The efficacy of amine- and carboxylterminated PAMAM–MTX conjugates were evaluated in MTX-sensitive and MTX-resistant human acute lymphoblastoid leukemia (CCRF-CEM) and Chinese hamster ovary (CHO) cell lines. Two amide-bonded PAMAM dendrimer–MTX conjugates were prepared with carboxylic acid-terminated G2.5 dendrimer and the amine groups of the MTX and another between an amineterminated G3 dendrimer and the carboxylic acid group of the MTX. Carboxylic acid-terminated G2.5 dendrimer conjugate showed an increased drug activity compared to free MTX toward both sensitive and resistant cell lines, whereas amine-terminated dendrimer conjugate did not show significant activity on any of the cell lines. The successful enhancement in cellular entry of MTX by carboxyl-terminated dendrimer conjugation was evidenced by 8and 24-fold enhancement in IC50 values toward MTX-resistant CCRF-CEM and CHO cells, respectively, which otherwise had impaired MTX transport mechanisms. The difference in efficacy of these amide-bonded conjugates was associated with the intracellular drug release from the cationic dendrimer versus the anionic dendrimer, due to the differences in lysosomal residence times dictated by the surface functionality. Carboxyl-terminated dendrimer conjugates may have longer lysosomal residence time and longer period of drug release compared to cationic PAMAM conjugate. Dendrimer–drug conjugates that depend on cytoplasmic conditions rather than lysosomal drug release pathways were also designed. An amineterminated G4 PAMAM dendrimer-N-acetyl cysteine (NAC) conjugate that contains a disulfide linkage was synthesized and evaluated for its efficacy on activated BV-2 microglial cells [29, 30]. The drug release pathway of the conjugates were glutathione (GSH) dependent, which is the most abundant thiol species in the cytoplasm functioning as the major reducing agent in biochemical processes. G4 PAMAM–NAC conjugate rapidly internalized by the cells within 15 minutes and the internalization continued for up to 4 hours. The conjugates showed an order of magnitude increase in antioxidant activity compared to free NAC, which suggests that the conjugate was successfully exiting the lysosomes and releasing its payload in the reductive cytoplasmic environment. Since the antioxidant effect of NAC is associated with its thiol group, which is occupied when in conjugated form, the conjugate would have to release the NAC to have efficacy. After the conjugates are taken up by endocytosis, they will reside in the lysosomes for a period of time, where the release of NAC may be relatively slow, because of the lower thiol content

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in lysosomes. As the conjugate escapes the lysosomal compartment, the NAC will be released into the cytoplasm. Therefore the lysosomal residence times of these types of dendrimer conjugates will ultimately affect their efficacy.

13.8

KEY ASPECTS IN DATA INTERPRETATION

Intracellular trafficking of dendrimers is often studied by attachment of some fluorescent molecules or other imaging agents followed by visualization of the uptake by confocal microscopy or quantification via flow cytometry techniques. Attachment of multiple copies of drugs to the functional surface groups of the dendrimers may alter their uptake characteristics. For a given dendrimer type, clearly the intracellular trafficking can be quite different if the dendrimer is conjugated to different drugs in two studies. The labeling of dendrimers with fluorescent molecules followed by in vitro visualization, identification of compartments, and colocalization studies via fluorescent or confocal microscopy techniques have some other limitations. Often, the fluorescent molecules have pH-dependent fluorescence or pH-dependent quenching properties. Concentration-dependent quenching is also observed in such studies. When considering the endocytotic uptake pathway of most dendrimers, such pH dependency can lead to artifacts, since the lysosomal pH is considerable different from that of the cytoplasm. Furthermore, the lysosomal accumulation could lead to a high enough concentration to introduce concentration-dependent quenching. Therefore it is important to evaluate such cellular uptake data with caution. The cellular uptake studies performed with such systems should be used as a guide for better delivery system designs, while efficacy of the ultimate therapeutic version of the dendritic delivery systems (without imaging agent) should be used for evaluating performance. Another consideration in studying cellular uptake and trafficking of dendrimers is the pathway inhibition strategies used for blocking certain pathways. These inhibition mechanisms are usually very complex and it is reported that one inhibitor can upregulate another cellular uptake mechanism under certain conditions. The experimental conditions can alter the uptake pathway of macromolecules completely even with the same cell type. The knowledge from a cell line that is used for the dendrimer uptake studies may not be readily transferable to other cell types due to variability in their membrane and intracellular trafficking characteristics.

13.9

CONCLUSION

Dendritic architecture creates opportunities for designing well-defined nanosized gene and drug delivery devices for tissue, cell, and even intracellular targeting. Due to their unique structure and advantages over linear or

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hyperbranched polymers in drug delivery applications, dendrimer research is receiving growing interest. In addition to cellular and intracellular targeting of dendritic devices, new applications are under investigation to overcome biological barriers such as gastrointestinal layers, amniotic membranes, and the blood–brain barrier. In order to create successful dendrimer-based designs, a solid understanding of intracellular trafficking and cellular uptake properties is required. The most important parameters to be optimized will be regarding surface properties of the dendrimers, such as size, overall charge, surface charge and surface charge density, the extent of drug or targeting moiety attachment, hydrophobicity, and hydrogen bonding capacity. Most of these parameters are interrelated with each other in every system; therefore researchers will have to gain strong fundamentals in engineering the surface of these globular macromolecules.

REFERENCES 1. Tomalia, D. A., Naylor, A. M., and Goddard, W. A. Starburst dendrimers: molecular-level control of size, shape, surface chemistry, topology, and flexibility from atoms to macroscopic matter. Angew. Chem. Int. Ed. Engl. 29: 138–175 (1990). 2. Hawker, C. J. and Fréchet, J. M. J. Preparation of polymers with controlled molecular architecture. A new convergent approach to dendritic macromolecules. J. Am. Chem. Soc. 112: 7638–7647 (1990). 3. de Brabander-van den Berg, E. M. M. and Meijer, E. W. Poly(propylene imine) dendrimers: large-scale synthesis by hetereogeneously catalyzed hydrogenations. Angew. Chem. Int. Ed. Engl. 32: 1308–1311 (1993). 4. Ihre, H., Hult, A., and Söderlind, E. Synthesis, characterization, and 1H NMR self-diffusion studies of dendritic aliphatic polyesters based on 2,2bis(hydroxymethyl)propionic acid and 1,1,1-tris(hydroxyphenyl)ethane. J. Am. Chem. Soc. 118: 6388–6395 (1996). 5. Grinstaff, M. W. Biodendrimers: new polymeric biomaterials for tissue engineering. Chemistry 8: 2838–2846 (2002). 6. Nilsen, T. W., Grayzel, J., and Prensky, W. Dendritic nucleic acid structures. J. Theor. Biol. 187: 273–284 (1997). 7. Li, Y., et al. Controlled assembly of dendrimer-like DNA. Nat. Mater. 3: 38–42 (2004). 8. Sadler, K. and Tam, J. P. Peptide dendrimers: applications and synthesis. J. Biotechnol. 90: 195–229 (2002). 9. Turnbull, W. B. and Stoddart, J. F. Design and synthesis of glycodendrimers. J. Biotechnol. 90: 231–255 (2002). 10. Svenson, S. and Tomalia, D. Dendrimers in biomedical applications—reflections on the field. Adv. Drug Deliv. Rev. 57: 2106–2129 (2005). 11. Duncan, R. and Izzo, L. Dendrimer biocompatibility and toxicity. Adv. Drug Deliv. Rev. 57: 2215–2237 (2005).

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12. Seib, F. P., Jones, A. T., and Duncan, R. Comparison of the endocytic properties of linear and branched PEIs, and cationic PAMAM dendrimers in B16f10 melanoma cells. J. Control. Release 117: 291–300 (2007). 13. Kolhe, P., et al. Drug complexation, in vitro release and cellular entry of dendrimers and hyperbranched polymers. Int. J. Pharm. 259: 143–160 (2003). 14. Perumal, O. P., et al. The effect of surface functionality on cellular trafficking of dendrimers. Biomaterials 29: 3469–3476 (2008). 15. Kolhe, P., et al. Preparation, cellular transport, and activity of polyamidoaminebased dendritic nanodevices with a high drug payload. Biomaterials 27: 660–669 (2006). 16. Khandare, J., et al. Synthesis, cellular transport, and activity of polyamidoamine dendrimer-methylprednisolone conjugates. Bioconjug. Chem. 16: 330–337 (2005). 17. Saovapakhiran, A., et al. Surface modification of PAMAM dendrimers modulates the mechanism of cellular internalization. Bioconjug. Chem. 20: 693–701 (2009). 18. Sweet, D. M., Kolhatkar, R. B., Ray, A., Swaan, P., and Ghandehari, H. Transepithelial transport of PEGylated anionic poly(amidoamine) dendrimers: implications for oral drug delivery. J. Control. Release 138: 78–85 (2009). 19. Kim, Y., et al. Systematic investigation of polyamidoamine dendrimers surfacemodified with poly(ethylene glycol) for drug delivery applications: synthesis, characterization, and evaluation of cytotoxicity. Bioconjug. Chem. 19: 1660–1672 (2008). 20. Kitchens, K. M., et al. Transport of poly(amidoamine) dendrimers across Caco-2 cell monolayers: influence of size, charge and fluorescent labeling. Pharm. Res. 23: 2818–2826 (2006). 21. Kolhatkar, R. B., et al. Surface acetylation of polyamidoamine (PAMAM) dendrimers decreases cytotoxicity while maintaining membrane permeability. Bioconjug. Chem. 18: 2054–2060 (2007). 22. Pisal, D. S., et al. Permeability of surface-modified polyamidoamine (PAMAM) dendrimers across Caco-2 cell monolayers. Int. J. Pharm. 350: 113–121 (2008). 23. Yang, H., et al. Stealth dendrimers for drug delivery: correlation between PEGylation, cytocompatibility, and drug payload. J. Mater. Sci. Mater. Med. 19: 1991–1997 (2008). 24. Kitchens, K. M., et al. Endocytosis inhibitors prevent poly(amidoamine) dendrimer internalization and permeability across Caco-2 Cells. Mol. Pharm. 5: 364–369 (2008). 25. Kitchens, K. M., et al. Transepithelial and endothelial transport of poly(amidoamine) dendrimers. Adv. Drug Deliv. Rev. 57: 2163–2176 (2005). 26. Malik, N., Wiwattanapatapee, R., Klopsch, R., Lorenz, K., Frey, H., Weener, J. W., Meijer, E. W., Paulus, W., and Duncan, R. Dendrimers: relationship between structure and biocompatibility in vitro, and preliminary studies on the biodistribution of 125I-labelled polyamidoamine dendrimers in vivo. J. Control. Release 65: 133–148 (2000). 27. Kang, H., DeLong, R., Fisher, M. H., and Juliano, R. L. Tat-conjugated PAMAM dendrimers as delivery agents for antisense and siRNA oligonucleotides. Pharm. Res. 2099–2106 (2005).

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28. Gurdag, S., et al. Activity of dendrimer–methotrexate conjugates on methotrexatesensitive and -resistant cell lines. Bioconjug. Chem. 17: 275–283 (2006). 29. Navath, R. S., et al. Dendrimer drug conjugates for tailored intracellular drug release based on glutathione level. Bioconjug. Chem. 19: 2446–2455 (2008). 30. Kurtoglu, Y. E., et al. Poly(amidoamine) dendrimer–drug conjugates with disulfide linkages for intracellular drug delivery. Biomaterials 30: 2112–2121 (2009).

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CHAPTER 14

Endolysosomolytically Active pHSensitive Polymeric Nanotechnology HAN CHANG KANG and YOU HAN BAE Department of Pharmaceutics and Pharmaceutical Chemistry, University of Utah, Salt Lake City, Utah

14.1

SITE-SPECIFIC NANOTHERAPEUTICS

Interest in developing pharmaceutical nanotechnology has blossomed with the desire to develop effective systems for delivering various therapeutics (e.g., from low molecular weight chemical drugs and imaging agents to high molecular weight peptides, proteins, and genetic materials) to specific sites of interest (e.g., organs, tissues, cells, cytoplasm, mitochondria, perinuclear regions, and nucleus). Unlike traditional dosage forms, site-specific nanotherapeutics are designed to maximize the bioavailability of the delivered therapeutics at the target sites and have shown beneficial therapeutic efficacy in treating diseases with reduced side effects [1]. Delivering nanosized therapeutic carriers to their target sites can be achieved by exploiting differences in anatomy, pathology, or cellular events between target and nontarget sites. First, anatomical specificity of the organs or tissues of interest can passively drive major accumulation of therapeutics. Hydrodynamic injection for liver targeting [2] and the enhanced permeability and retention (EPR) effect for tumor accumulation [3] are known examples of passive organ/tissue targeting. Second, nanosystems can selectively target solid tumors using pathological differences. For example, the local pH of solid tumors is lower than the extracellular pH of normal tissues, creating the opportunity for pH targeted delivery [4]. (Please refer to Lee et al. [4] for more detailed information.) Third, targeting specific cells has been attained mostly by utilizing particular interactions between ligands and cell-specific or

Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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overexpressed receptors located on the plasma membrane. These ligand– receptor interactions accelerate the internalization rates of therapeutics and their carriers into the target cells and enhance the intracellular accumulation [5]. However, if the final destination of therapeutics carried by nanocarriers is subcellular compartments (e.g., cytoplasm, mitochondria, perinuclear areas, and nucleus), the therapeutics or their carriers should be able to escape from the endolysosomal pathways in order to achieve maximum therapeutic effects [6].

14.2

DESTABILIZATION OF ENDOLYSOSOMAL COMPARTMENTS

Therapeutics or nanocarriers larger than 1 kDa rarely cross the plasma membrane without the aid of cell-penetrating peptides [7]. These systems use other cellular entry mechanisms such as endocytosis: that is, the cellular membrane invaginates to engulf therapeutics or pharmaceutical nanosystems and forms intracellular membrane-bound vesicles (or endosomes). The therapeuticsloaded vesicles mature from early endosomes (neutral pH to approximately pH 6) to late endosomes (approximately pH 5 to pH 6) as acidification occurs due to cytosolic protons (H+) pumped into the endosomes by vacuolar ATPase-H+ pumps. As endosomes acidify even further, late endosomes merge with lysosomes (approximately pH 4 to pH 5), which contain various lytic enzymes [1, 7]. In lysosomes, the sequestrated therapeutics may be degraded by lytic enzymes, resulting in reduced bioavailability at the target site. To avoid such a scenario, therapeutics-carrying nanosystems should recognize endolysosomal characteristics, which are easily distinguishable from the extracellular and intracellular environment. One unique trait of endolysosomal compartments is pH—endolysosomal pH is generally lower than extracellular pH and cytosolic pH. To target specific pH ranges, materials containing specific functional groups (i.e., amines/imines [8–11], carboxylic acids [12–15], and sulfonamides [16] as shown in Figure 14.1) have often been used because the functional groups enable the transition between protonation/deprotonation states. Disrupting the endolysosomal membrane may be induced by the proton buffering capacity and/or fusogenicity of the “endolysosomolytic materials.” The “proton sponge” effect suggested by Boussif et al. [8] is related to inhibiting or delaying the acidification process of endolysosomes. When materials having protonable groups are entrapped in the endosomes, protons transferred by vacuolar ATPase-H+ pumps are mostly consumed by the materials themselves and do not contribute much toward endosomal maturation/ acidification. With further proton influx into the endosomes, chloride ions (Cl−) also accumulate in the endosomes, leading to osmotic imbalances between the cytoplasm and the endosomal compartments. This imbalance causes continuous water influx, endosomal swelling, and finally leads to endosomal rupture [17–20].

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Amine/Imine-containing materials H N

H N

N H

N

N H H N

NH2 O H N CH C O

NH N

N

CH2

NH2 N

NH

NH

HN Poly (L-histidine) Branched PEI Carboxylic acid-containing materials C3H7 H2 C C

C2H5 H2 C C COOH

COOH

Poly (2-ethylacrylic acid)

Poly (2-propylacrylic acid)

Sulfonamide-containing materials CH3

S

H2 CH3 C C C=O NH

N

N

N N OCH3 Poly/oligo Poly/oligo sulfamethizole sulfadimethoxine R=

O=S=O NH R

OCH3

N

N

N

N

Poly/oligo sulfadiazine

CH3

Poly/oligo sulfamerazine

Figure 14.1. Examples of protonable oligomers/polymers that destabilize the endosomal membrane. (Copyright © 2008 Springer Science + Business Media, LLC [1].)

Fusogenic materials cause physical interactions between pH-sensitive membrane destabilizers and endosomal membranes. For example, positively charged polymers can interact with negatively charged phospholipid bilayers in the endosomal membrane, destabilizing the vesicle structure [17]. Polymers having pH-induced hydrophilic-to-hydrophobic transitions (i.e., anionic polyelectrolytes) show similar destabilization mechanisms, which were investigated using lipid bilayers (i.e., liposomes). That is, endosomal acidification increased the hydrophobicity of polymers, which could lead to interactions

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with the hydrophobic tail region of the endosomal phospholipid bilayer membrane. This interaction stimulates lateral compression and forms pores in the bilayer, allowing the release of therapeutics into the cytoplasm [21–23].

14.3

ENDOLYSOSOMOLYTIC POLYMERIC NANOCARRIERS

Various materials (e.g., polymers, liposomes, peptides, viral components, and their hybrids) can induce the destabilization of endolysosomal compartments for effectively delivering therapeutics. This destabilization process is mostly triggered by a “pH” stimulus via various physical and chemical transitions of swelling/deswelling, hydrophilicity/hydrophobicity, micellization/demicellization, complexation/decomplexation, ionic/neutral charge, and α-helix/random coils conformational changes [1]. Target pH for endolysosomal rupture can range from extracellular pH to lysosomal pH while traversing the endolysosomal pathway. However, extracellular pH can vary depending on pathology (e.g., pH 7.4 for normal blood; for solid tumors, average pH 6.8 at normoglycemia and average pH 6.4 at hyperglycemia [24]; for brain, pH 7.2 (normal) and pH 6.4 (ischemic) [25]; for heart, pH 7.5 (normal) and approximately pH 6.8 (ischemic) [26]). Thus if certain materials can disrupt the endolysosomal membranes within specific pH ranges, nanocarriers with membrane destabilizers can be designed to treat specific diseases or target certain cells. Although there are various types of endolysosomolytic materials, this chapter will focus primarily on synthetic polymer/oligomer-triggered endolysosomal disruption. A brief summary of recent endolysosomolytic strategies developed by the Bae research group is presented with representative examples of chemical drug delivery nanocarriers (for low molecular weight therapeutics) and gene delivery nanocomplexes (for high molecular weight therapeutics). (For more information regarding other systems, please refer to their extensive reviews [6, 7, 14, 17, 27].) 14.3.1 Endolysosomolytic Polymeric Anticancer Drug Nanocarriers 14.3.1.1 Poly(L-histidine)-Based Micelles Histidine is an attractive amino acid for endosomolysis because its side chain, an imidazole ring, has pKa 6.0 and sharp proton buffering at pH 6.0 [28]. Also, as shown in Figure 14.2A, its homopolymer poly(l-histidine) (polyHis; see Figure 14.1 for its chemical structure) and various copolymers (i.e., polyHis-b-poly(ethylene glycol) (PEG)) show proton buffering over a broad pH range (pH 4 to pH 9) [29]. The apparent pKb values for polyHis5kDa and polyHis3kDa were about 6.5, whereas their PEGylated copolymers (i.e., polyHis5kDa-b-PEG2kDa and polyHis3kDa-b-PEG2kDa) showed a slightly higher apparent pKb of 7.0. These differences in apparent pKb values may result from differences in water content because the hydrophilic PEG block can induce more hydration [29].

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Additional evaluation of polyHis for endolysosomolysis ability was performed using hemolysis because the membranes of red blood cells (RBCs) are similar to endolysosomal membranes. As shown in Figure 14.2B, polyHis5kDab-PEG2kDa disrupted 5–20% of total RBC number with decreasing pH from 7.4 to 6.75, whereas its hemolytic activity at pH 6.5 sharply increased to nearly 90%. However, poly(l-lactic acid) (PLLA)3kDa-b-PEG2kDa, which is used for preparing polyHis5kDa-b-PEG2kDa-containing mixed micelles, caused pHindependent membrane rupture of less than 5% in erythrocytes because the polymer does not possess pH-sensitive or endosomolytic groups [30]. To create a carrier for therapeutics (here, mostly hydrophobic chemical drugs), a polyHis block was conjugated to a hydrophilic PEG block. PolyHisb-PEG can maintain its micellar structure at basic pH due to its amphiphilicity, whereas at acidic pH its hydrophilicity can destabilize the micelles. At pH 8, micelles made from polyHis5kDa-b-PEG2kDa were more stable than those prepared from polyHis3kDa-b-PEG2kDa because the critical micelle concentration (CMC) of polyHis5kDa-b-PEG2kDa was much lower than polyHis3kDa-b-PEG2kDa (2.3 μg/mL for polyHis5kDa-b-PEG2kDa vs. 62 μg/mL for polyHis3kDa-b-PEG2kDa). However, the destabilization of polyHis5kDa-b-PEG2kDa-based micelles started at pH 7.4 and the extent of destabilization increased with decreasing pH [29]. This fact indicates that micelles fabricated from polyHis5kDa-b-PEG2kDa only are not suitable for destabilizing endolysosomes. To improve micelle stability at pH 7.4, the hydrophobic copolymer PLLA3kDa-b-PEG2kDa was introduced when preparing polyHis5kDa-b-PEG2kDa mixed micelles [31]. In mixed micelles made from polyHis5kDa-b-PEG2kDa and PLLA3kDa-b-PEG2kDa, the increased hydrophobicity (i.e., increased weight fraction of PLLA3kDa-b-PEG2kDa) lowered the pH that triggered micelle destabilization. PLLA3kDa-b-PEG2kDa (10 wt%) in the mixed micelles slightly improved micelle stability compared to micelles prepared from polyHis5kDa-bPEG2kDa. More PLLA3kDa-b-PEG2kDa (25 wt%) considerably enhanced micelle stability at pH 7.4 and initiated micelle destabilization below pH 7.0. Mixed micelles fabricated from 60 wt% of polyHis5kDa-b-PEG2kDa and 40 wt% of PLLA3kDa-b-PEG2kDa had a slightly lower destabilization pH than micelles containing 25 wt% PLLA3kDa-b-PEG2kDa. These results strongly correlated with the trigger pH for the release of doxorubicin (DOX, an anticancer drug). For the mixed micelles, DOX release could be triggered to release from pH 6.6 to pH 7.2 by modulating the weight fraction of PLLA3kDa-b-PEG2kDa [31]. These findings suggest that mixed micelles containing higher fractions of PLLA3kDa-b-PEG2kDa had weaker pH sensitivity although the micelle destabilization pH remained the same. Following the discovery that the pH at which micelle destabilization is triggered is closely linked to enhanced drug release, Yin et al. [32] investigated whether it was possible to predict pH-triggered micelle destabilization by monitoring pH-dependent changes in micelle size distribution using dynamic light scattering. The mixed micelles (75 wt% of polyHis5kDa-b-PEG2kDa and 25 wt% of PLLA3kDa-b-PEG2kDa) formed a spherical shaped secondary

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(a) 14

(b) 120

NaCl polyHis5kDa-b-PEG2kDa polyHis5kDa polyHis3kDa-b-PEG2kDa polyHis3kDa

10

100 Hemolysis (%)

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8 6 4

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1 N HCl (mL) (c)

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(f) 120

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100 80 60 40 20 0 0.0001 0.001 0.01 0.1 1 DOX concentration (μg/ml)

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Figure 14.2. (A) Acid–base titration curves of polyHis and polyHis-b-PEG. (Reproduced from Ref. 29. Copyright © 2003, with permission from Elsevier B.V.) (B) Hemolytic activity of polyHis5kDa-b-PEG2kDa and PLLA3kDa-b-PEG2kDa. (Copyright © 2008 Springer Science + Business Media, LLC [30].) (C) Schematic illustrations of micelle destabilization. (Reproduced from Ref. 32. Copyright © 2008 with permission from Elsevier B.V.) (D) The effects of PLLA3kDa-b-PEG2kDa on micelle destabilizationtriggering pH (pHt) in mixed micelles made from polyHis5kDa-b-PEG2kDa and PLLA3kDab-PEG2kDa. (Reproduced from Ref. 32. Copyright © 2008 with permission from Elsevier B.V.) (E) In vitro endosomal disruption using fluorescein DHPE-encapsulated PHSM/f and PHIM/f and the LysoTracker® dye against A2780/DOXR MDR cells. (Copyright © 2008 Springer Science + Business Media, LLC [30].) (F) In vitro cytotoxicity of free DOX (䉬), DOX-loaded PHSM/f (䊉), and PHIM/f (䊏) against A2780/DOXR MDR cells after 48-h incubation. (Copyright © 2008 Springer Science + Business Media, LLC [30].) (G) In vivo tumor growth inhibition test of s.c. A2780/DOXR MDR cells-bearing BALB/c nude mice using control (䉬), free DOX (ⵧ), DOX-loaded PHSM/f (䊉), and DOX-loaded PHIM/f (䉱). Three intravenous doses of 10 mg/kg DOX equivalent dose were administered at Days 0, 3, and 6. (Copyright © 2008 Springer Science + Business Media, LLC [30].) (See color insert.)

structure in the range of pH 7.0 to pH 7.4 because individual core–shell micelles with relatively hydrophobic cores associated together. In the pH range 7.0–7.4, the mixed micelles had a narrow and unimodal size distribution. When pH was decreased from 6.8 to 6.5, there was an obvious increase in the size and aggregation number of the micelles caused by the destabilization of the micelle core. Their size distribution was broad and unimodal, but hydrophilic polyHis5kDa-b-PEG2kDa unimers were released from the micelles. Further decreases in pH (reaching pH 6.0) resulted in a bimodal distribution of micelle size because most polyHis5kDa-b-PEG2kDa unimers had dissociated from the micelle core (see Figure 14.2C for an illustration of pH-induced micelle destabilization) [32]. As shown in Figure 14.2D, when different amounts of PLLA3kDab-PEG2kDa were introduced into the mixed micelles, pH-triggered micelle destabilization decreased with increasing weight fraction of PLLA3kDa-bPEG2kDa (e.g., pHt 6.8 for 25 wt% of PLLA3kDa-b-PEG2kDa and pHt 6.5 for 40 wt% of PLLA3kDa-b-PEG2kDa) [32]. With endolysosomal acidification, the protonated polyHis5kDa-b-PEG2kDa can cause the endolysosomes to swell due to proton buffering of the polymer. In addition, according to the destabilization mechanism of the micelles suggested by Yin et al. [32], the polyHis5kDa-b-PEG2kDa unimers released from the micelles can interact with the endolysosomal membrane. The proton buffering and fusogenic characteristics of polyHis5kDa-b-PEG2kDa may disrupt the endolysosomal compartments as shown in Figure 14.2E. When mixed pH-sensitive micelles (PHSM) containing targeting folate (PHSM/f) (i.e., micelle systems prepared from 80% of polyHis5kDa-b-PEG2kDa and 20% of PLLA3kDa-bPEG2kDa-folate) and mixed pH-insensitive micelles (PHIM) with folate

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(PHIM/f) (i.e., micelle systems prepared from 80% of PLLA3kDa-b-PEG2kDa and 20% of PLLA3kDa-b-PEG2kDa-folate) were administered to DOX-induced multidrug-resistant (MDR) ovarian carcinoma cell lines (i.e., A2780/ DOXR MDR cells), PHSM/f caused an even distribution of green fluorescent DHPE (N-(fluorescein-5-thiocarbamoyl)-1,2-dihexadecanoyl-sn-glycero3-phosphoethanoamine triethylammonium salt) and red fluorescent LysoTracker® dye in the cytoplasm. However, LysoTracker® and DHPE delivered using PHIM/f were scattered locally in the cytoplasm. This difference in the intracellular localization of DHPE and LysoTracker dye indicated that polyHis5kDa-b-PEG2kDa in the PHSM/f is an important component for the endolysosomal escape of therapeutics. Endolysosomolytic polyHis5kDa-b-PEG2kDa-containing PHSM/f effectively delivered DOX and killed both drug-sensitive tumor cell lines and MDR tumor cell lines [10, 30]. As shown in Figure 14.2F, DOX-loaded PHSM/f effectively killed in the range of tested DOX concentrations without displaying the cytotoxic effects of free DOX and DOX-loaded PHIM/f. It is noteworthy that DOX-loaded PHSM/f killed approximately 60% of A2780/DOXR MDR cells at 1 μg/mL of DOX, whereas free DOX and DOX-loaded PHIM/f killed approximately 10% and 20% of the cells, respectively [30]. For in vivo models of A2780/DOXR MDR cells-bearing BALB/c nude mice, PHSH/f treatment effectively inhibited tumor growth for 1 month, whereas free DOX and PHIM/f treatments did not suppress tumor growth (Figure 14.2G). 14.3.1.2 Poly(L-histidine-co-L-phenylalanine)-Based Micelles When PHSM/f were exposed to acidic extracellular pH (i.e., pH 6.4–6.8 for solid tumors and ischemia), micelles with up to 40 wt% of PLLA3kDa-b-PEG2kDafolate became destabilized in extracellular environments. Thus to destabilize micelles containing endosomolytic materials at pH lower than the destabilization pH of PHSM/f, polyHis was modified with the hydrophobic amino acid phenylalanine (Phe). As summarized in Table 14.1, pH-sensitive copolymers of His and Phe (i.e., poly(His-co-Phe) (PHP)) with various apparent pKb were synthesized. Their molecular weights were approximately 5 kDa and their apparent pKb values ranged from 6.7 to 4.8 as the hydrophobic Phe mole% in the copolymer was increased [33].

TABLE 14.1

Characterization of Poly(His-co-Phe)

Polymer

Phe (mol%) in Polymer

Apparent pKb

Buffering pH Range

10 16 22 27

6.7 6.3 5.7 4.8

8.2–4.8 7.6–4.7 6.5–4.0 5.8–3.2

PHP10 PHP16 PHP22 PHP27

Source: Reproduced with permission from Ref. 33. Copyright © 2008 Wiley-VCH Verlag GmbH & Co.KGaA.

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1200

100

1000

80

800

Size (nm)

120

60 40

600

Intensity (%)

(B)

(A)

Transmittance (%)

255

30 25 20 15 10 5 0 1

100 10000 Size (nm)

400

20

200

0 3

4

5

6

7 8 pH

9 10 11 12

0

8

7.4

6.5 pH

6

5.5

Figure 14.3. (A) pH-dependent transmittance transition of PHSMPhe10 (䉫), PHSMPhe16 (䉱), PHSMPhe22 (ⵧ), and PHSMPhe27 (䊉). (B) pH-dependent particle size of mPHSMPhe16(5%) (black), mPHSMPhe16(10%) (gray), and mPHSMPhe16(20%) (white). The inset shows a bimodal particle size distribution at pH 6.5 using mPHSMPhe16(5%). (Reproduced with permission from Ref. 33. Copyright © 2008 Wiley-VCH Verlag GmbH & Co.KGaA.)

Like polyHis, poly(His-co-Phe) (PHP) was conjugated with PEG for encapsulating hydrophobic drugs. The pH-sensitive micelles (PHSMPhe) prepared from PEGylated PHP (i.e., PHP-b-PEG2kDa) showed transmittance transition at pH 7.0 for PHSMPhe10, at pH 6.5 for PHSMPhe16, at pH 5.8 for PHSMPhe22, and at pH 5.2 for PHSMPhe27 (Figure 14.3A). To obtain finer tuning of the destabilization pH, PLLA3kDa-b-PEG2kDa was mixed with PHP-b-PEG to prepare mixed micelles (mPHSMPhe). In the initial trial to target early endosomal pH values, PHP16-b-PEG2kDa was selected because its micelles were stable at pH 7.0 but became destabilized at pH 6.5. The micelles “mPHSMPhe16” prepared from various weight ratios of PHP16-b-PEG2kDa and PLLA3kDa-bPEG2kDa showed pH-dependent changes in micelle size, and more dramatic size changes were seen with increasing PLLA3kDa-b-PEG2kDa content. As shown in Figure 14.3B, the size of mPHSMPhe16(5%) (i.e., mPHSMPhe16 prepared from 95 wt% PHP16-b-PEG2kDa and 5 wt% of PLLA3kDa-b-PEG2kDa) prepared at pH 8 increased as pH dropped below pH 6.5, and the sizes of mPHSMPhe16(10%) and mPHSMPhe16(20%) increased below pH 6 and sharply below pH 5.5, respectively. At pH 6.5, a bimodal size distribution of mPHSMPhe16(5%) indicated that ionized PHP blocks were not miscible with PLLA3kDa-b-PEG2kDa and dissociated from PLLA3kDa-b-PEG2kDa micelles [33]. In vitro endolysosomal escape of mPHSMPhe16(20%)/f (i.e., mPHSMPhe16 prepared from 80 wt% PHP16-b-PEG2kDa and 20 wt% of PLLA3kDa-b-PEG2kDafolate) was similar to that of the PHSM/f system as shown in Figure 14.2E. That is, mPHSMPhe16(20%)/f caused even intracellular distribution of DHPE and LysoTraker dyes. This result may be induced by the PHP16-b-PEG2kDa

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released from the mixed micelles. The micelle systems effectively killed both A2780 wild-type cells and A2780/DOXR MDR cells [33]. Based on the destabilization pH of mPHSMPhe16 with different amounts of PLLA3kDa-b-PEG2kDa, mPHSMPhe22 and mPHSMPhe27 prepared from PHP22 and PHP27 with PLLA3kDa-b-PEG2kDa could be designed to target late endosomes and lysosomes, respectively, because PHSMPhe22 and PHSMPhe27 fabricated from PHP22 and PHP27 showed micelle destabilization at pH 5.8 and pH 5.2, respectively. 14.3.1.3 Virus-Mimetic Nanogels Virus-mimetic (VM) nanogels, which can migrate from one cell to other cells, have been developed to effectively kill solid tumors. As shown in Figure 14.4A, the nanogels consist of a hydrophobic PHP core (i.e., poly(His32-co-Phe6) with nearly pKb 6.4) and two hydrophilic layers of PEG2kDa and bovine serum albumin (BSA). Like the capsid shell of a virus, the gel’s mimetic outer shell is comprised of the core and BSA bridged by multiple PEG chains. Interestingly, the nanogels have a reversible swelling/deswelling characteristic that is pH dependent. As shown in Figure 14.4B, the size of VM nanogels at pH 7.4 was approximately 55 nm with a narrow distribution. At pH 6.8, VM nanogels still showed similar size and size distribution to those at pH 7.4. However, upon exposure to pH 6.4 (close to early endosomal pH), the nanogels became bigger up to approximately 355 nm with a broader distribution due to the ionization and swelling of the PHP core. This pH-induced swelling/deswelling transition of VM nanogels modulated DOX release rates. As shown in Figure 14.4C, DOX release rates from the nanogels at pH 7.4 and pH 6.8 were similar because the solidified core at pH 7.4 and pH 6.8 entrapped hydrophobic drugs. However, pH 6.4 induced the ionization and swelling of the PHP core and allowed more DOX to release. The release rate at pH 6.4 was approximately threefold faster than at pH 7.4 and pH 6.8. In addition, acidic pH-induced volumetric expansion of VM nanogels and known proton buffering capacities of histidine-based polymers (i.e., poly(His16-co-Phe6)) induced endolysosomal disruption as shown in Figure 14.4D. Control nanopaticles (NPs) without endolysosomolytic characters were localized with LysoTracker dye, whereas VM nanogels showed even intracellular distribution. Acidic pH-induced DOX release profiles and the endolysosomolytic function of VM nanogels effectively killed A2780 wild-type cells and A2780/ DOXR MDR cells compared to free DOX and control NP treatments. As shown in Figure 14.4E, the antitumor activity of the nanogels in pH 6.8 medium demonstrated that the nanogels were stable at pH 6.8 and released DOX to effectively kill tumor cells. Interestingly, after killing tumor cells, the nanogels that still contained DOX migrated to kill other tumor cells (Figure 14.4F). This concept of a “nanogel” that is stable in an acidic extracellular environment and can be activated to release drugs and to disrupt endolysosomes

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pH 7.4 pH 6.4 DOX released F: folate F F F PEG Inner shell F F F F F F F pH F F F F F F F F F FF FF F F F BSA outer shell F FF 355 nm 55 nm

(A)

F F F F

F

F

(B) pH 7.4

pH 6.8

0 100

pH 7.4

0 100

pH 7.4

pH 6.4

pH 6.8

0 100

0 100

pH 7.4

pH 6.4

0 100 0 200 400 600 800 0 100 0 200 400 600 800

Particle size (nm)

(D) 20

FITC

Lyso Tracker

Merged

Control NPs VM-nanogels

pH pulses relative rate of DOX (μg/hour)

(C) 15 10 5 0

pH 7.4 pH 6.8 pH 6.4

: appled pH for 1 hour each

Cell viability / %

80

**

**

60 40 20 0

**

**

A2780 A2780/ A2780 A2780/ DOXR DOXR MDR cells MDR cells

Blank DOX-loaded DOX-loaded VM-nanogels Free DOX control NPs VM-nanogels

(F) pH 6.8

pH 7.4

(E) 100

Infected

not Infected

Subsequent Infections

A

B-0

B-1

B-2

B-3

A

B-0

B-1

B-2

B-3

A

B-0

B-1

B-2

B-3

A

B-0

B-1

B-2

B-3

Figure 14.4. VM nanogel: (A) Schematic presentation. (B) pH-modulating size change: A (pH 7.4), B (pH 6.8), and C (pH 6.4). (C) pH-dependent DOX release rate from DOX-loaded VM nanogels. (D) In vitro endosomal escaping activity. (E) pHdependent cytotoxicity of DOX-loaded VM nanogel (white), DOX-loaded control nanoparticles (gray), and free DOX (black) against A2780 wild-type cells and A2780/ DOXR MDR cells after 48-h incubation (DOX dose = 1 μg/mL). (F) Migration of DOXloaded VM nanogels in A2780/DOXR MDR cells. (Reproduced with permission from Ref. 38. Copyright © 2008 Wiley-VCH Verlag GmbH & Co.KGaA.) (See color insert.)

while contained in the endolysosomal pathway could be expanded to treat ischemia in the brain and heart. In this case, although migration of the nanogel toward other cells might be difficult, the pH-dependent drug release profile of the nanogel could effectively prevent the progress of ischemia over a long time period because ischemic states are initiated by drops in pH.

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14.3.2 Endolysosomolytic Polymeric Gene Nanocomplexes Kang and Bae have been interested in developing cell-customized endosomolytic agents for more effectively delivering therapeutics because endolysosomal characteristics such as endolysosomal pH [34, 35], endolysosomal membrane composition [36, 37], and acidification rate are cell specific. To investigate cell-specific endolysosomolytic agents, sulfonamides were selected. Sulfonamides have broad-range pKa values (3–11) and hydrophobicity conferred by various substituted groups, R (see Figure 14.1) [16]. For feasibility studies, sulfamethizole (SMT; pKa 5.45), sulfadimethoxine (SDM; pKa 6.1), sulfadiazine (SDZ; pKa 6.4), and sulfamerazine (SMZ; pKa 7.0) were selected because their pKa values fall within the endolysosomal pH range. Using chain transfer radical polymerization, the synthesized sulfonamide oligomers (designated as OSMT, OSDM, OSDZ, and OSMZ) had Mn = 1.8–2.5 kDa. Oligomeric sulfonamides (OSAs) showed different proton buffering and aqueous solubility transition within the endosomal/lysosomal pH range. In aqueous solubility transition studies of the OSAs (Figure 14.5A), OSMZ showed solubility transitions over a broad pH range, whereas OSMT, OSDM, and OSDZ showed relatively sharp transmittance changes within a narrow pH range. As shown in Figure 14.5B, OSMT and OSDZ demonstrated broad proton buffering over the pH ranges of 5.0–6.4 and 5.7–7.3, respectively, whereas OSDM and OSMZ buffered protons at a specific pH, 6.5 and 7.3, respectively. Their apparent pKa values were 5.7 (OSMT), 6.5 (OSDM and OSDZ), and 7.3 (OSMZ) and were slightly higher than their monomeric sulfonamides counterparts [16]. To understand the endolysosomolytic functions of anionic OSAs in polymeric nanocarriers, a well-known polymeric gene delivery system without endosomolytic function (here, poly(l-lysine) (PLL)-based polyplexes) was selected. Anionic OSAs cannot directly carry anionic genes, but OSAs can be incorporated within any polycation-based gene carriers. In intracellular distribution studies of polyplexes using pH-sensitive fluorescein pDNA (F-DNA), OSDZ-polyplexes (i.e., PLL/OSDZ/F-DNA) showed broad distribution in the cytoplasm, whereas PLL/F-DNA complexes were mostly localized (Figure 14.5C). These differences in fluorescence distribution demonstrate that OSA polyplexes caused more pDNA to escape from endosomal/lysosomal compartments than PLL/pDNA complexes. This faster endosomal release of OSDZ polyplexes resulted in faster gene expression than for PLL/pDNA complexes. In in vitro transfection studies using three different cell lines (i.e., human hepatoma HepG2 cells, human embryonic kidney HEK293 cells, and rat insulinoma RINm5F cells), OSA polyplexes showed 4–55-fold better transfection efficiency than control polyplexes (PLL/pDNA) as shown in Figure 14.5D. Interestingly, OSDM and OSDZ were more effective in transfecting HEK293 cells whereas OSMZ was the best for transfecting RINm5F cells. This study supports the need for cell-customized endosomolytic agents to achieve

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CONCLUSION

8

PLL/OSDZ/F-DNA

(D)

150 mM NaCl OSMT OSDM OSDZ OSMZ OPAA

4

(C)

6 pH

PLL/F-DNA

7

20μm

150 mM NaCl OSMT OSDM OSDZ OSMZ OPAA

100 150 200 250 300 350 Added volume of 0.1 N HCI (μL)

80 70 60 50 40 30 20 10 0

HEK293 HepG2 RINm5F

PL

L PE O I SM O T SD M O SD O Z SM O Z PA A

20μm

5

Normalized Transfection Efficiency (Fold)

Transmittance (%)

100 90 80 70 60 50 40 30 20 10 0

(B) 11 10 9 8 7 6 5 4 3 2 50

pH

(A)

259

Polyplex type

Figure 14.5. (A) pH-dependent aqueous solubility transition of OSA and OPAA (oligomeric propylacrylic acid) solutions. (B) Acid–base titration curves of OSA and OPAA solutions. (C) In vitro intracellular localization studies of PLL/pDNA and PLL/ OSDZ/pDNA complexes using pH-sensitive F-DNA. (D) In vitro transfection studies for OSA-containing PLL/pDNA complexes (OSA polyplexes) containing a luciferase gene in HepG2, HEK293, and RINm5F cells. Dose of OSA and OPPA was 5 nmol (based on their monomeric units) per 1 μg pDNA. Normalized transfection efficiency was defined as (Absolute transfection efficiency of polyplexes)/(Absolute transfection efficiency of PLL/pDNA complexes) for a specific cell. Charge ratio (+/−) of polyplexes was 3 except for PEI/pDNA (+/− = 5). (Reproduced with permission from Ref. 16. Copyright © 2007 Wiley-VCH Verlag GmbH & Co.KGaA.)

clinically effective gene delivery [16]. In addition, these anionic materials could provide an opportunity to revisit the use of other biocompatible but less transfection-efficient gene carriers.

14.4

CONCLUSION

Endolysosomolytic materials are an indispensable component for effectively targeting subcellular compartments and maximizing therapeutic benefits. Because endolysosomal destabilization is mostly mediated by “pH,” understanding extracellular pH and endolysosomal pH of various pathological states

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is important. Knowledge about the pH in the body, organs, tissues, cells, and extracellular and intracellular environments for healthy and disease states would aid the design of effective endolysosomolytic polymeric nanocarriers.

ACKNOWLEDGMENTS This work is partially supported by NIH grants (CA101850 and GM82866). The authors appreciate the help of Deepa Mishra, Dr. Haiqing Yin, and Dr. Dongin Kim for their critical reading.

REFERENCES 1. Kang, H. C., Lee, E. S., Na, K., et al. Stimuli-sensitive nanosystems: for drug and gene delivery. In: V. P. Torchilin, ed. Multifunctional Pharmaceutical Nanocarriers. Springer, New York, 2008, pp. 161–199. 2. Herweijer, H. and Wolff, J. A. Gene therapy progress and prospects: hydrodynamic gene delivery. Gene Ther. 14: 99–107 (2007). 3. Maeda, H., Bharate, G. Y., and Daruwalla, J. Polymeric drugs for efficient tumortargeted drug delivery based on EPR-effect. Eur. J. Pharm. Biopharm. 71: 409–419 (2009). 4. Lee, E. S., Gao, Z., and Bae, Y. H. Recent progress in tumor pH targeting nanotechnology. J. Control. Release 132: 164–170 (2008). 5. Nishikawa, M. Development of cell-specific targeting systems for drugs and genes. Biol. Pharm. Bull. 28: 195–200 (2005). 6. Breunig, M., Bauer, S., and Goepferich, A. Polymers and nanoparticles: intelligent tools for intracellular targeting? Eur. J. Pharm. Biopharm. 68: 112–128 (2008). 7. Bareford, L. M., and Swaan, P. W. Endocytic mechanisms for targeted drug delivery. Adv. Drug Deliv. Rev. 59: 748–758 (2007). 8. Boussif, O., Lezoualc’h, F., Zanta, M. A., et al. A versatile vector for gene and oligonucleotide transfer into cells in culture and in vivo: polyethylenimine. Proc. Natl. Acad. Sci. USA. 92: 7297–7301 (1995). 9. Pichon, C., Goncalves, C., and Midoux, P. Histidine-rich peptides and polymers for nucleic acids delivery. Adv. Drug Deliv. Rev. 53: 75–94 (2001). 10. Lee, E. S., Na, K., and Bae, Y. H. Doxorubicin loaded pH-sensitive polymeric micelles for reversal of resistant MCF-7 tumor. J. Control. Release 103: 405–418 (2005). 11. Yang, S. R., Lee, H. J., and Kim, J. D. Histidine-conjugated poly(amino acid) derivatives for the novel endosomolytic delivery carrier of doxorubicin. J. Control. Release 114: 60–68 (2006). 12. Jones, R. A., Cheung, C. Y., Black, F. E., et al. Poly(2-alkylacrylic acid) polymers deliver molecules to the cytosol by pH-sensitive disruption of endosomal vesicles. Biochem. J. 372: 65–75 (2003).

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CHAPTER 15

Uptake and Intracellular Dynamics of Proteins Internalized by Cell-Penetrating Peptides ARWYN T. JONES Welsh School of Pharmacy, Cardiff University, Cardiff, Wales, United Kingdom

15.1

INTRODUCTION

Many pathogens and therapeutic macromolecules need to enter cells to fulfill a requirement for, respectively, survival and replication or the treatment of disease. An ability to target specific cells and then overcome the barrier posed by the plasma membrane to reach the cytosol or a defined subcellular organelle, and to then mediate an effective biological response is, however, a feat limited to pathogens or pathogen proteins. An example is the HIV transactivator of transcription protein, HIV-Tat, that enters cells by endocytosis, escapes from the endolysosomal system to the cytosol, and then moves to the nucleus to mediate its effects on transcription. This protein highlights the extraordinary capacity of evolution to manufacture simple but highly efficient targeting systems. Essential for HIV-Tat to perform these feats is a basic protein transduction domain that can enter cells either alone or associated with cargo of various forms ranging from other proteins to nanoparticles. Hundereds of natural and synthetic protein transduction domains or cellpenetrating peptide (CPP) sequences have now been described and this chapter highlights the attempts that have been made to understand the cellular dynamics of proteins attached by various means to CPPs. An appreciation of the mechanisms by which these systems operate may hold answers to address a major pharmaceutical need for the design of more efficient systems for cellular delivery of therapeutic proteins. This chapter will describe endocytosis and endocytic pathways (Section 15.2), the need to deliver proteins into cells Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

263

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(Section 15.3), our current understanding of the mechanism by which the HIV-Tat enters cells and reaches the nucleus (Section 15.4), delivery of proteins linked to other CPP sequences (Section 15.5), alternative mechanisms for covalent and noncovalent attachment of CPPs to proteins (Section 15.6), delivery of CPP proteins to lysosomes (Section 15.7), and a comparison of the cellular dynamics of CPP proteins with CPP fluorophores (Section 15.8).

15.2

ENDOCYTIC PATHWAYS

The plasma membrane is a highly effective and selective boundary between the inside and outside of cells. All cells, however, have a physiological need to internalize membrane-impermeable molecules and to do this they invaginate portions of their plasma membrane to internalize membrane and accompanying extracellular fluid. This process is called endocytosis and the internalized membrane or vesicular structures are then delivered together with the fluid cargo to a sorting station that directs further membrane traffic to one of a number of cellular destinations. Only recently are we beginning to appreciate the enormous complexities of endocytosis and endocytic pathways [1]. This process, regulated by a network of proteins and lipids, is essential for several often unrelated cellular functions, and this may be why a single cell has the capacity to internalize material through several different endocytic pathways (Figure 15.1). These include those originating from clathrin-, and non-clathrin-coated structures such as caveolae through to larger structures called macropinosomes that are formed following plasma membrane ruffling. It is beyond the scope of this chapter to discuss endocytosis at great length and those wishing to gain a good foundation on the current knowledge of endocytic pathways should consult the reviews in Refs 1–9 and also this informative web site at http://endocytosis.org/. 15.2.1 Using Endocytic Pathways to Deliver Therapeutics The effectiveness of using endocytosis to deliver therapeutic macromolecules is in part constrained by the fact that the fate of the therapeutic within one of these pathways is predetermined by the cell’s trafficking mechanisms and biological barriers posed initially by the plasma membrane and then by the endolysosomal network [10–12]. Thus fragile molecules such as proteins and genes may rapidly be delivered to hostile environments such as lysosomes and inactivated before they have any chance to reach and interact with their intended targets. Pathogens and pathogen-proteins especially toxins have been shown to exploit and manipulate these pathways to reach defined subcellular destinations, thus suggesting that therapeutics can also be delivered by similar mechanisms [13, 14]. Endocytic features such as low pH and high degradative activities can, however, act as activators of bioresponsive molecules to enhance translocation of the active entity before the cell’s endocytic pathway

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Figure 15.1. Endocytic pathways and cell entry mechanisms available for proteins attached to cell-penetrating sequences or natural cell-penetrating proteins such as HIV Tat. Numbers 1–6 represent defined pinocytic pathways and not described here or in the text is phagocytosis.1, Clathrin-coated vesicles; 2, caveolae; 3, fluid-phase uptake; 4, flotillin-1-dependent pathway; 5, other clathrin- and caveolin-independent pathway(s) [5]; 6, macropinocytosis, and 7, direct penetration and entry through the plasma membrane. The named proteins represent those required for the pathways to operate and thus offer the opportunity for their mutation or depletion via siRNA. This together with pharmacological inhibitors will allow for more accurate requirement of particular pathways for entry of macromolecules designed for intracellular delivery and targeting. It is likely that an early or sorting endosome exists to accept cargo delivered through all these endocytic pathways and some have been defined. It still remains to be determined as to whether entry via any particular pathway is advantageous for allowing subsequent escape from the endolysosomal system to locate the protein in the cytosol.

traffics it to its final destination. Enhancing the delivery of therapeutics through endocytosis is therefore dependent on acquisition of a high level of understanding of specific endocytic pathways inherent in the target cell, the traffic and fate of the molecule within endocytic organelles, the effect of the macromolecule on the dynamics of endocytic pathways, the downstream effects of these on the integrity of the cell, and if required the mechanism by which escape from the endolysosomal system occurs.

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15.2.2 Identifying Endocytic Pathways Cell biologists studying endocytic pathways are consistently identifying new protein and lipid mediators and a goal for this remit that overlaps with that of drug delivery is to identify a protein that is only involved in regulating one distinct pathway. This then opens the possibility of either mutating this protein or inhibiting its manufacture to gain a better understanding of the pathway that it regulates and also the fate of the cargo that it helps to deliver. Much of the current information regarding the endocytic pathways utilized by drug delivery systems, including those using CPPs, comes from studies using microscopy and endocytic probes [15, 16] in conjunction with a library of pharmacological inhibitors of endocytic pathways [17]. Examples include methyl ß-cyclodextrin (MßCD), amiloride, chlorpromazine, and cytochalasins. At one time these agents were deemed to be specific for certain pathways but it is now clear that all suffer predominantly from a lack of specificity, and toxicity is also a common problem [17]. Increasingly, researchers are using molecular biology tools such as cell lines expressing dominant negative forms of endocytic proteins or siRNA-mediated depletion of proteins that regulate endocytic pathways. This siRNA approach has recently identified new pathways and roles in endocytosis for proteins such as flotillin-1 [18] and significantly added to the current knowledge on established endocytic proteins such as clathrin [19–21]. The use of siRNA depletion has its own drawbacks for those wishing to use the technology for analyzing endocytosis of their drug delivery systems and these include (1) the fact that a delivery system is first required to introduce the siRNA into the cytosol, (2) the possibility that the protein of interest regulates other membrane trafficking pathways, not necessarily endosomal (clathrin heavy chain and dynamin are good examples), and (3) the fact that depletion of the protein affects cell viability or dysregulates other endocytic pathways. Another important aspect of this approach is that a specific probe is required whose uptake is confined to one pathway; this allows researchers to identify whether expression of mutant proteins or siRNA depletion is having a biological response. Transferrin is a very well-established marker for uptake via clathrin-coated vesicles [22], both anti CD35 antibodies and poly(ethylenimine) complexes have been shown to be partially internalized in a flotillin-1-dependent manner [18, 23] and cholera toxin is often used as a marker for uptake via caveolae. A number of other poorly defined pathways exist and identifying specific endocytic probes for those not organized by clathrin has been difficult [5, 9]. For example, a number of studies show that cholera toxin can access through different pathways [9] and very careful analysis is required to ensure that uptake is confined to one route [24]. 15.3

DELIVERING PROTEINS INTO CELLS

Delivering a full-length protein that may have multiple domains, interactions, and effects is sometimes required to mediate a necessary biological response, whether for research or therapy. The realistic potential of delivering full-

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length proteins was probably seeded when it was realized that many diseases were caused by malfunctioning or missing proteins and the technology became available for their purification, especially from transformed microorganisms such as Escherichia coli. Most proteins are membrane impermeable and a large fraction of them function inside cells; therefore systems are required for their delivery into their site of action, often the cytosol and in some cases subcellular organelles such as the nucleus and mitochondria. This therefore requires an intracellular delivery system and potential candidates for performing this function are short peptide sequences that have the capacity to enter cells either as single entities or attached to much larger cargo such as fulllength proteins. These are called protein transduction domains (PTDs) or cell-penetrating peptides. 15.3.1 Defining Cell-Penetrating Peptides and Protein Transduction Domains The term cell-penetrating peptide gained prominence around 20 years ago as did the use of the term protein transduction domain and there is still healthy discussion in meetings over the most appropriate terminology for what could generally be described as membrane active peptides. Peptides that interact with eukaryotic and prokaryotic membranes have, however, been described in the literature for decades before either of these now familiar terms gained prominence [25]. An excellent review written by Plank et al. [26] in 1998 described a number of peptides with potential for delivering macromolecules and examples include melittin, mastoparan, GALA, and KALA; these are more familiarly known as pore-forming, membrane active, or destabilizing peptides (Table 15.1). However, as previously noted, the distinction between these and what we now refer to as CPPs or PTDs can sometimes be difficult to define [27]. Hundereds of peptides have now been described that share a common ability to deliver cargo to cells and their heterogeneity with respect to sequence and physical and biological properties suggests that neither of these commonly used terms is appropriate for all. One of the best characterized is the cell-penetrating peptide from the protein transduction domain of the HIV-Tat protein.

15.4

HIV-TAT PROTEIN

The genome of HIV encodes proteins that are important for structure, organization of its envelope, enzyme regulators, and also transactivators such as HIV-Tat. This 101 amino acid protein is manufactured inside infected cells and gains entry into the nucleus, where it binds RNA to activate viral transcription [28]. Residues 1–86 are transactivation active and a number of laboratories have used this truncated form for their studies, rather than the full-length protein. The protein also appears to be released from intact virus-infected cells and this fraction can enter other cells via endocytosis

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TABLE 15.1

Sequences of Membrane Active Peptidesa Described in This Chapter

Peptide HIV-Tat Bov-Tat JDV Tat Transportan TP10 pVEC Penetratin (pAntennapedia) MPG PEP-1 PTD-5 PDX-1 PTD VP22 GALA KALA Melittin Mastoparan HA2 HA2′

Sequence

Reference

GRKKRRQRRRPPQ RRRGTRGKGRRIRR RHDGRRKKRGTRGKGR GWTLNSAGYLLGKINLKALAALAKKIL AGYLLGKINLKALAALAKKIL LLIILRRRIRKQAHAHSK RQIKIWFQNRRMKWKK

127 54 53 83 106 84 78

GALFLGFLGAAGSTMGAWSQPKKKRKV KETWWETWWTEWSQPKKKRKV YARAARRAARR RHIKIWFQNRRMKWKK DAATATRGRSAASRPTERPRAPARSASRPRRPVE WEAALAEALAEALAEHLAEALAEALEALAA WEAKLAKALAKALAKHLAKALAKALKACEA IKITTMLAKLGKVLAHV INLKALAALAKKIL GDIMGEWGNEIFGAIAGFLG GLFEAIEGFIENGWEGMIDGWYG

89 88 81 62 128 92 92 25 25 58 64

a

Variations on these sequences are commonly used, especially the addition of terminal residues such as cysteine for conjugation to fluorophores. Bov, bovine immunodeficiency virus; JDV, Jembrane disease virus; PDX-1, pancreatic and duodenal homeobox factor-1 protein transduction domain.

[29–32]. But most importantly for those interested in drug delivery, the protein then escapes from the endolysosomal network to the cytosol and then on to the nucleus [33]. HIV-Tat has also been shown to interact with a number of other cell surface molecules to mediate biological effects, and its location on the plasma membrane and interaction with viral gp-120 may also aid in promoting viral entry [34, 35]. A crystal structure for HIV-Tat has not been published, but interestingly NMR studies reveal that the protein may be one of the many proteins that exist as an unfolded entity that may allow for multiple interactions with affectors and effectors [28, 36–38]. The basic domain is seen fully exposed on the surface (Figure 15.2) but it is likely that other parts of the molecule, perhaps the hydrophobic core, also aid in endosomal escape. 15.4.1 Cellular Dynamics HIV-Tat Protein The biology of HIV-Tat protein has been the subject of intense scrutiny and like other HIV proteins it is a target for anti-HIV drugs [28]. Its biology, either manufactured by the protein synthesis machinery inside a virus-infected cell or following uptake from the extracellular fluid, is extremely complex and over 250 HIV-Tat interacting partners have now been identified [39].

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(A)

269

(B)

Figure 15.2. NMR structures of HIV-Tat 1–86. Both (A) [37] and (B) [36] show quite different structures but common is the exposure of the basic domain, residues 48–60, shown in blue. The deposited structures were obtained from the Protein Data Bank and were processed using Pymol software.

15.4.1.1 Binding to the Plasma Membrane A requirement for HIV-Tat endocytosis, endosomal escape, nuclear enrichment, and RNA binding is a short highly basic sequence spanning residues 47–57 (Table 15.1). This region interacts with negatively charged cell surface heparan sulfate proteoglycans and is thought to gain access to the cell as an ionic complex with these sugars [40–42]. The initial cellular uptake and endocytosis of HIV-Tat protein and of functional residues 1–72 were performed using E. coli expressed protein that was then purified and labeled with 125I [30, 31]. Uptake was partially characterized in a number of cell lines and found to be dependent on temperature and could be inhibited by competing anionic charge in the form of heparin. Of note is the fact that the protein at 1 nM (∼10 ng/mL) was able to promote transactivation—an amazing feat and one that bears consideration when looking at subsequent studies aiming to delineate in much more detail its uptake and intracellular dynamics and also of HIV-Tat fusion proteins and HIV-Tat peptide linked proteins. Immediate to the finding that HIV-Tat could enter cells were studies showing that major fractions of the protein or the basic region could be linked to the N or C terminus of proteins and promote their uptake as active entities both in vitro in a number of different cell lines, and also in vivo [43–45]. There was thus an obvious interest in elucidating the mechanism by which these proteins were able to overcome membrane bilayers. The creation of a HIVTat-Enhanced Green Fluorescent protein (EGFP) fusion protein confirmed that this domain interacted strongly with heparan sulfate proteoglycans and that attenuated uptake of the fusion protein and its capacity for transactivation was observed in Chinese hamster ovary (CHO) cells deficient in proteoglycan synthesis [42, 46]. Similar reduced uptake and transactivation was noted when

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soluble glycosaminoglycans, most notably heparin, were added to cell culture media during incubations with HIV-Tat fusion proteins and with HIV-Tat alone [31]. Recent data in primary brain cells also demonstrate a degree of positive correlation between glycosaminoglycan expression and the ability of GFP-HIV-Tat peptide fusion to gain cell entry [47]. 15.4.1.2 Endocytosis The most extensive characterization of endocytosis and translocation of native HIV-Tat (residues 1–86) was performed in Jurkat T lymphocytes [35]. Uptake of the protein at 50 nM was inhibited by approximately 50% in cells treated with chlorpromazine, a well-known inhibitor of clathrin-coated uptake and it was localized, using immunocytochemistry, in clathrin-coated pits. In support of this, the ability of Tat to escape from the endolysosomal system was inhibited in cells expressing mutant proteins that affected clathrin-coated uptake. A cell-free translocation assay was developed to monitor Tat release from endosomes and this revealed that escape was dependent on endosomal acidification. 15.4.1.3 Fixing Artifacts It is fair to say that the field of cell-penetrating peptides shuddered somewhat when Lundberg and co-workers published two reports regarding the uptake and subcellular localization of GFP extended with cell-penetrating sequences from HIV-Tat and herpes virus protein VP22, octalysine and octaarginine [48, 49]. The studies highlighted the different capacities of these four sequences to label CHO cells but also suggested that fixing cells for microscopy and flow cytometry had major effects leading to an overestimation of the capacity of CPPs to deliver cargo to the inside of cells and the nucleus rather than to label, respectively, the plasma membrane and endocytic vesicles. These studies and others investigating fluorescently labeled CPPs [50] had a huge effect on this field and awoke researchers to the realization that studies with CPPs, (especially the cationic forms) needed to be performed in unfixed cells, and that stringent washing procedures with enzymes and anionic molecules were required to remove surface label. 15.4.1.4 Endocytic Pathways Utilized by HIV-Tat Protein The aforementioned study of HIV-Tat endocytosis in Jurkat cells was performed in fixed cells using anti HIV-Tat antibodies [35], but to perform live cell imaging there was a requirement to tag the protein with a fluorophore or to extend its sequence with a fluorescent protein such as GFP. In these and other studies, additional tags are added to the protein to aid in purification and these include His6 and glutathione S-transferase tags [49, 51] that may or not be linked to the protein of interest via protease sensitive linkages [52]; this minimizes the possibility of tag-mediated effects. This technology has recently been used to investigate the uptake and subcellular distribution of Tat proteins from HIV 1 and more recently other viruses [53, 54]. Comparative analysis has been performed on the uptake and intracellular localization of GST-Tat (full length)-EGFP and GST-Tat(48–60)EGFP in a

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HeLa cell derivative HL3T1 [51, 55]. Uptake of both was dependent on endocytosis and the actin cytoskeleton and fluorescence was shown to colocalize in vesicles labeled with caveolin-1 and cholera toxin that is often used as a marker of uptake via caveolae. The problems of using cholera toxin as an endocytic pathway marker have been described above. The cholesterol-sequestering agent MßCD inhibited the uptake of both fusion proteins, suggesting that lipid rafts were necessary for uptake, and very little colocalization was observed with fluorescent transferrin, a marker of uptake via clathrin-coated vesicles. This data was rather different from studies in T cells showing that uptake of HIV-Tat alone (i.e., without cargo) was through clathrin-coated pits [35]. One important aspect of the study on the uptake of HIV-Tat in the absence of cargo was the fact that it was performed in Jurkat T cells that do not express caveolin-1 [56], and thus uptake in this cell line could not have been mediated by caveolae. HeLa cells, on the other hand, express low levels of caveolin-1 but it is difficult to visualize caveolae in this cell line [57]. Overall, the studies demonstrate that caveolae are not absolutely required for cell entry and escape from endocytic pathways. A role of clathrin and caveolae was disputed when a Tat-Cre recombinase assay was used to investigate the capacity of the basic domain of Tat to deliver the 38-kDa Cre recombinase protein to mediate expression of the EGFP reporter gene [58]. The purified Cre protein was labeled with Alexa488 and its uptake into cells was shown to be inhibited after treatment with the cholesterol-interacting agent nystatin, that along with MßCD also inhibited recombination. The study showed that Tat-Cre entry and function were not dependent on caveolin expression and thus uptake via caveolae. Recombination, however, was inhibited following cytochalasin D and amiloride treatment, suggesting that macropinocytosis may be involved in peptide uptake. This endocytic process usually occurs in growth factor stimulated cells and involves extensive actin reorganization and engulfment of large volumes of extracellular liquid that are then enclosed to form internal macropinosomes [4, 6, 59]. Vesicles formed from other endocytic pathways have diameters in the range of 60–120 nm but macropinosomes can be micrometers in diameter. These enlarged structures were not observed but in support of a role for this pathway in Tat-Cre uptake was an increased fluid phase uptake of 70 kDa dextran. This was an important observation suggesting that the basic domain of Tat was interacting with the plasma membrane to cause cytoskeletal activation, ruffling, and enhanced uptake of fluid-phase markers, and possibly itself. It is important to note here that 70-kDa dextran (hydrodynamic radius 5.5 nm) is not a specific marker for macropinocytosis, as is sometimes mentioned in the literature, but rather a good probe to monitor enhanced fluid-phase uptake that is characteristic of this endocytic event. The studies did suggest, however, that entry of this protein and indeed cationic CPPs was not a passive mechanism but one that involved activation and reorganization at the plasma membrane.

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A later study using a similar Tat-Cre system also showed that energy and an intact actin cytoskeleton were required for cell entry [60]. The data also demonstrated the limitations of confocal microscopy for these kinds of analysis in so much that despite the fact that there was no visible (microscopical) evidence for cytosolic and nuclear Cre, the biological readout assay suggested otherwise. Conversely, confocal microscopy and laser-induced membrane damage and leakage can also potentially overestimate the fraction of an internalized entity that is released in the cytosol [61]. Macropinocytosis was also implicated in the uptake of the transcription factor pancreatic and duodenal homeobox factor 1 (PDX-1) [62] that has within its sequence a cell-penetrating domain (Table 15.1). Labeling recombinant PDX-1 with fluorescein isothiocyanate (FITC) and adding it to cells allowed for microscopical analysis of its distribution, and at 5-μM extracellular concentration the protein was found to label vesicles and the cytosol. Further analysis of the recombinant protein was not performed but amiloride was shown to inhibit uptake of the protein transduction domain of this protein. 15.4.2 Endocytosis of Tat from Other Viruses GFP fusion proteins of Jembrane disease virus and bovine immunodeficiency virus Tat have been generated and these also have basic rich sequences, and the Jembrane disease virus basic domain is longer than that of HIV-Tat (Table 15.1). Both of these proteins efficiently delivered GFP into cells but the uptake mechanism was not studied in detail. Interestingly, there was evidence that uptake of denatured Tat-EGFP constructs occurred via an energyindependent manner whereas uptake of the native proteins was inhibited in the absence of energy and therefore dependent on endocytosis. But this needs further analysis to reveal whether the proteins were actually located inside the cells rather than on the cell surface. 15.4.3 Escaping from the Endolysosomal Network If endosomal escape of HIV-Tat protein is dependent on acidification, then the possibility exists that the ultimate fate of the fraction that cannot escape will be degradation in lysosomes. One study utilized a mutant GFP (F64L, T203Y) that has a pH-dependent emission profile and its pH environment can then be monitored during endocytosis [63]. The mutant was extended with the HIV-Tat basic sequence and the protein was then added to cells to reveal that the fusion protein reached a compartment of pH 5.8 before the GFP fluorescence was quenched and the protein degraded. Extensive degradation had occurred within 8 hours. Degradation is a major issue if there is a requirement for the therapeutic to be delivered from the endolysosomal system, and beyond, and attempts have been made to enhance escape from endosomes. One study showed that coincubation of a membrane fusion peptide HA2 derived from the N terminus of influenza A hemagluttinin (Table 15.1) to

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incubations with Tat-Cre enhanced its capacity for recombination [58]. A longer sequence that was also called HA2 (HA2 in Table 15.1) was shown to enhance the transcriptional activity of extracellularly delivered p53RRRRRRRRR fusion protein [64]. However, the approach here was quite different as in this case the CPP, fusion peptide, and delivered cargo were linked as one protein HA-p53-R9. Subsequent labeling of this protein with FITC and confocal microscopy analysis in live glioma cells showed it to be diffusely localized and enriched in the nucleus; this was 4 hours postincubation. The same fusion protein lacking the HA2 moiety was predominantly localized to the plasma membrane. The exact mechanism by which HIV-Tat protein in the absence of a defined fusion sequence escapes from the endolysosomal system is unknown but to aid viral replication a fraction must gain access to first the cytosol and then the nucleus. The previously mentioned GFP mutant study [63] demonstrated that the protein reached a compartment of pH 5.8 but whether the “active” fraction has already escaped before this pH is reached is unknown. It is technically difficult to analyze endosomal escape of a protein and there are many unanswered questions regarding the dynamics of this protein once it has gained access into cells. One interesting method employed fluorescent resonance energy transfer (FRET) to monitor the uptake and endosomal escape of the Tat peptide sequence conjugated to fluorescent protein mCherry that formed the acceptor fluorophore for FITC [65]. FRET between these two molecules was quenched by 4-(4′-dimethylaminophenylazo)benzoic acid (Dabcyl) and the fluorescence profile of the construct was dependent on whether it was intact or degraded in endosomes or the cytosol. Although this was a complex system, it offered an opportunity to monitor in real time the fate of delivered proteins fused with penetrating sequences. 15.4.4 Entry into the Nucleus Passive diffusion should allow cytosol–nucleus exchange for molecules <45 kDa [66] and full-length HIV-Tat protein should therefore be able to access by diffusion. EGFP is a 27-kDa protein and it is expected that EGFP-HIV-Tat protein fusion proteins (masses of 36 kDa or 37 kDa, depending on whether full length or the commonly used residues 1–86 are used) should also be able to access the nucleus by diffusion. There is, however, some dispute over whether nuclear entry of EGFP-HIV-Tat is in fact an active process. Early studies in cells overexpressing HIV-Tat protein-EGFP showed the protein to be highly enriched in the nucleus and when expression was very high it was also very noticeable in the nucleolus [67]. A number of studies have subsequently aimed to determine the mechanism by which the protein reaches the nucleus and whether the basic stretch of amino acids, in addition to promoting cell entry and endosomal escape, also harbors a bona fide nuclear localization sequence (NLS) [68, 69]. Typically, NLSs are monopartite or bipartite with respect to basic sequences [66] as exemplified,

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respectively, by the NLS from SV40 T antigen PKKKRKV [70] and nucleoplasmin (KRPAATKKAGQAKKKKL) [71]. Studies using GFP and GFP dimers or tetramers containing, at the N terminus, the HIV-Tat basic domain sequence (YGRKKRRQRRR-GFP) suggest nuclear localization of GFP or GFP dimers occurs in the absence of energy and thus via passive diffusion [72]. However, by substituting the three arginines terminal to the basic sequence with glycines to give YGRKKRRQGGG-GFP, the nuclear localization of a 110-kDa GFP tetramer cargo, that was normally absent from the nucleus, could be achieved [73]. This suggests that the three consecutive arginines in the HIV-Tat sequence GRKKRRQRRRPPQ may in fact hinder nuclear entry. These studies have important implications for researchers attempting to deliver therapeutics to the nucleus.

15.5

PROTEIN DELIVERY BY OCTAARGININE

GFP variants have also been shown to be delivered to cells by extending the sequence with octaarginie (R8) that, like HIV-Tat, represents one of the most studied CPPs [49, 74]. R8-EGFP when introduced extracellularly to CHO cells was localized predominantly to the plasma membrane but the images only show data after 5 minutes uptake [49]. It is likely that longer incubations are needed to clearly visualize the protein in endosomes and possibly the cytosol. Flow cytometry analysis clearly showed that extending EGFP with R8 enhanced its cell association, but these studies did not use trypsinization to lift the cells from the plates and thus the fluorescence may have been overestimated. The addition of pyrenebutyrate, a hydrophobic negatively charged counterion, to the cell media before addition of Alexa488-R8 peptide significantly enhanced the fluorescence signal in the cytosol and there was also evidence that the same was true for R8-EGFP [74]. It was suggested that pyrenebutyrate interaction with the arginine guanidinium groups of R8, and with membranes, promoted R8-mediated delivery of EGFP to the cytosol. This effect, however, may be cell type dependent [75].

15.6 SYSTEMS FOR ATTACHING CELL-PENETRATING PEPTIDES TO PROTEINS Typically, cloning strategies are used to extend the N or C terminus of proteins to cell-penetrating peptide sequences. However, a number of other approaches have been used to link them to protein cargo (Figure 15.3). 15.6.1 Covalent Attachment 15.6.1.1 Disulfide Bonds The Tat sequence and R7 have also been attached to proteins such as tissue-specific plasminogen activator and

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7

275

1

SS

6

2

AvBi

ion

3

lat

e ch

5 4

Figure 15.3. Protein delivery strategies using CPP sequences or mimetics. 1, CPP sequence attached to cargo via a peptide bond; 2, full-length or truncated transducing proteins fused to cargo proteins; 3, CPP-mimetic–protein complex; 4, ionic or hydrophobic CPP–protein complexes; 5, CPP–protein chelates; 6, CPP–protein avidin–biotin complexes; 7, CCP sequence is linked to a protein through an S–S bond. Center shows early endosomes (red) and nucleus (blue) of a liver Hep3B cell.

asparaginase using disulfide linkages formed with the crosslinking agent N-succinimidyl 3-(2-pyridyldithio)propionate [76, 77]. FITC-labeled Tat–S-Sasparaginase was shown to be diffusely distributed in HeLa cells but the cells were fixed for microscopy and the extracellular concentration of Tat was not provided [77]. 15.6.1.2 Cell-Penetrating Peptide Mimetics Efforts have also been made to mimic the translocation capacities of cell-penetrating peptides to design new “synthetic” entities based on the well-characterized tendency for CPPs to be amphipathic and to contain guanidinium groups. An example was described using the antennapedia sequence [78] (Table 15.1) and its helical structure as a template for the design of a biphenyl mimic small molecule carrier (SMoC). This was shown to effectively deliver FITC and geminin, a 23-kDa DNA replication inhibitor protein [79]. Delivery of this protein caused growth inhibition of cancer cells, suggesting successful delivery of the protein to the nucleus. The uptake of the SMoC was inhibited at low temperatures, showing a requirement for energy, but internalization was not inhibited by a range of endocytic inhibitors including cytochalasin D, chlorpromazine, and the amiloride analog EIPA. Similar data was obtained with SMoC protein

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cargo with the exception that chlorpromazine inhibited uptake by 70%. This suggests that a large fraction entered cells via clathrin-coated vesicles. An intermediate of this SMoC could be derivatized to accommodate a wide range of attachment chemistries for conjugation to multiple types of cargo. 15.6.2 Noncovalent Linkages between Cell-Penetrating Peptides and Proteins 15.6.2.1 Avidin–Biotin Affinity GFP has been a very useful tool for analyzing the uptake and cellular dynamics of proteins containing CPP sequences. A number of studies aiming to better understand these processes have used the high affinity and specificity of avidin–biotin chemistry. One such study compared the capacity of biotinylated Tat, R4-12R, 4K-12K to deliver Alexa488-streptavidin and streptavidin-ß-galactosidase to five different cell lines [80]; also included here was a modified HIV-Tat basic sequence PTD-5 (Table 15.1) [81]. Despite the fact that these studies were preformed before the “fixing” papers came to press, some of the experiments employed trypsinization to remove surface-associated label. Tat, PTD-5, R6-12, and K6-12 were shown to deliver these cargoes but the extent by which they were able to do this was highly dependent on the cell line. Generally, the most efficient delivery sequences were either K8 or K10. This is interesting, as the literature shows that many more papers since this period have used oligoarginine and Tatbased sequences compared with oligolysines. Avidin–biotin chemistry was also used to compare the delivery capacity of HIV-Tat peptide, penetratin, transportan, and pVEC [82]. Penetratin is the protein transduction domain from the Drosophila Antennapedia protein [78], transportan is a chimera of sequences from peptides galanin and mastoparan [83], and pVEC is derived from murine cadherin [84]. Streptavidin was conjugated to the fluorophore FITC and this was preincubated with the biotinylated peptides to form avidin–biotin complexes that were then added to cells. Transportan and Tat had significantly higher delivery capacity than the other peptides but these experiments were only performed in HeLa cells. Interestingly, preincubating the cells with MßCD had only a minimal effect on uptake by HIV-Tat and transportan but despite having much lower delivering capacity, Antennapedia and pVEC mediated uptake seemed to be much more sensitive to cholesterol depletion. Extensive confocal and electron microscopy studies on biotinylated transportan, and its deletion analog TP10 [85] complexed with fluorescent or goldlabeled avidin, suggested that there was little overlap of the complexes with internalized transferrin and that labeled structures akin to macropinosomes were observed using electron microscopy. The ultimate fate of the complexes was deemed to be lysosomes but this study also raised the possibility that these CPPs could be delivering protein cargo directly across the plasma membrane and also highlighted the problems of complex aggregation in the extracellular

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media, thus making data interpretation very difficult. The main problems with using electron microscopy is that the cells need to be fixed, but parallel live cell imaging and electron microscopy can give revealing information as described here. Later studies from the same group showed reduced uptake (∼30–50%) following siRNA depletion of caveolin-1 in HeLa cells and also reduced uptake in caveolin-1 knockout mouse embryonic fibroblasts compared to control cells [86]. Again confocal microscopy showed very little evidence of colocalization of these complexes with endocytosis markers such as rab5, transferrin, and flotillin-1 but some colocalization was observed with caveolin-1 using confocal and electron microscopy. 15.6.2.2 Cell-Penetrating Peptide–Protein Complexes Cell-penetrating peptides have also been shown to deliver biologically active proteins via complexation rather than through direct peptide bond linkage or through avidin– biotin chemistry. The most well described are amphipathic peptides MPG, Pep-1 (Table 15.1), and their derivatives [87–89]. Their sequence is deemed to allow for electrostatic and/or hydrophobic interactions with a number of different proteins, but once the complex is formed the peptides still have the necessary functionalities to interact with cells and allow delivery to the cytosol. Despite observations showing that these peptides, as single entities or associated with cargo, induce actin rearrangements [90], the current thinking is that delivery may not require any form of endocytosis and that the access route is directly across the plasma membrane [87, 89]. This is thought to be mediated via pore formation; however, studies investigating siRNA delivery with MPG suggest that endocytosis is very much involved [91]. The implications of a mechanism for delivering proteins directly across the plasma membrane are clear as it could reduce or negate degradation issues that are a feature of endocytosis if the cargo is fragile to the hostile environment of late endosomes and lysosomes. Other notable pore-forming peptides with delivery capacity are the KALA and GALA peptides (Table 15.1) and related sequences [26, 92]. Comparative studies on the uptake of avidin or streptavidin either as complexes with cell-penetrating sequences or linked via biotin was revealing in that it showed among other things that the capacity of cell-penetrating sequences to deliver cargo was very much dependent on the nature of the cargo [93, 94]. An uncharacterized synthetic CPP YTA2 (Table 15.1) was analyzed against a number of well-characterized CPPs and overall the studies show that the effectiveness of any particular sequence for delivering cargo, including full-length proteins, is very much dependent on the physical properties of the cargo and most probably the choice of studied cell line or tissue. The issue of CPP-mediated toxicity and membrane leakage has been discussed in some detail [95–97] and overall the studies suggest that great care needs to be taken when interpreting data measuring uptake of cargo without parallel viability assays.

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15.6.2.3 Cell-Penetrating Peptide–Protein Chelates Linking expressed fusion proteins, with histidine residues is routine for purification of bacterially expressed proteins, but the Futaki group exploited this further when they manufactured a nitriliotraiacetic acid derivatized octaarginine peptide NTA. This forms noncovalent complexes with His residues and this affinity was shown to mediate the uptake of His6-EGFP into mammalian cells [52]. The derivatized peptide was also able to deliver a His6 tagged proapoptotic peptide PAD (KLAKLAK)2 to cells to inhibit viability. Numerous studies have shown effective in vitro and in vivo cell killing with CPP-PAD conjugates [80, 98–102] but the possibility exists in some cases that cationic CPPs and PAD, when linked together, synergistically disrupt the plasma membrane and cause necrosis rather than apoptosis [103].

15.7

TARGETING LYSOSOMES

In some situations it is beneficial to keep the protein in the endolysosomal system to allow it to reach its target or substrate in the lysosomes. This is especially true for diseases caused by a deficiency of a protein that naturally resides and functions in these organelles. An example is ßglucuronidase deficiency that causes mucopolysaccharide type VII syndrome, a lysosomal storage disease. This protein can gain access to cells via the mannose-6-phosphate receptor, which can then traffic it to the lysosome. A study investigated the uptake of this protein compared to the ß-glucuronidase sequence extended with the HIV-Tat peptide [104]. This C-terminal extension changed the endocytic uptake mechanism and generally uptake was reduced, but in tissues such as the kidney, higher levels of the chimera were observed and these correlated with enhanced clearance of glycosaminoglycans. This may be due to the reduced renal clearance that was observed for the chimera but it also showed effective intracellular delivery of this protein to the lysosomes and that the presence of the Tat domain negated the requirement for the receptor to deliver ß-glucuronidase to these organelles. Very similar results were obtained when galactocerebrosidase was extended with Tat peptide in an effort to treat Krabbe disease, another lysosomal storage condition [105].

15.8 COMPARING CPP FUSION PROTEINS WITH CPP FLUOROPHORES Despite the fact that a wealth of information exists on the capacity of CPPs to deliver active full-length or truncated proteins, only a few studies have carefully scrutinized uptake and biological activity in the same study. To analyze the uptake and intracellular traffic of CPP–protein conjugates is technically challenging as there is a requirement for cloning a cDNA for a

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fusion protein, expressing this, for example, in E.coli, purifying the protein, and then monitoring its uptake as a fluorescent entity. Here there is a further requirement that its conjugation to a fluorescently labeled large (GFP) or relatively small (FITC/Alexa dyes, <1 KDa) fluorophores does not interfere with its cellular dynamics and activity. Labeling peptides with fluorophores and analyzing their uptake is, however, relatively simple and hence the process by which CPP fluorophores interact with cells and gain access to the cell interior and beyond has been the subject of intense scrutiny. Readers are referred to numerous reviews that have recently been published on experiments in vivo, in vitro, and in artificial systems such as membrane bilayers [4, 11, 27, 106–112]. A number of the endocytic pathways described in Figure 15.1 have been implicated as being involved in entry of CPPs attached to fluorophores such as FITC and Alexa dyes. Some common themes are now emerging that should help researchers in drug delivery and cell biology maximize their translocation capacities, especially in cell-based assays. For cationic peptides there is clear evidence that they can traverse the plasma membrane of some cell lines in the presence and absence of energy [113–115]. It is also clear that at 37 °C and at relatively high peptide concentrations (=5 μM) the plasma membranes of some cell lines are permeable to these peptides but not to dyes such as propidium iodide when these are coicubated with the CPPs [114, 116–118]. However, these concentrations are very close to those that then show a general increase in membrane porosity and thus propidium iodide entry [97]. The presence of serum in the media has significant effects owing to the general affinity of the peptides to proteins such as albumin that then reduces the effective peptide concentration at the plasma membrane [117]. An interesting question is how we can relate these studies to those described above, where the cargo is much larger than a fluorophore and where a requirement for endocytosis is more likely. With different cargoes, as previously suggested, the mechanism by which membrane bilayers are breached and especially the location of this translocation may be quite different.

15.9

CONCLUSION

Following the initial revelation that a CPP could deliver a 120 kDa protein to cells and different tissues [43, 45], we have witnessed a rapidly expanding literature on the potential of this technology for the delivery of proteins as therapeutics [108, 119–122]. Important questions regarding the usefulness of this approach for delivering proteins and a library of other cargoes have been asked [123–126] but those working in the field will strive to enhance their vector capacity. Future research in this area promises the usual mixture of interesting science, controversy, and hopefully news of CPP conjugates progressing from clinical trials to the clinic.

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ACKNOWLEDGMENTS The author apologizes to those whose work was not mentioned here and who have contributed to this field, and would like to thank Nicola Zonta of Cardiff University for help in generating Figure 15.2.

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95. Saar, K., Lindgren, M., Hansen, M., Eiriksdottir, E., Jiang, Y., Rosenthal-Aizman, K., Sassian, M., and Langel, U. Cell-penetrating peptides: a comparative membrane toxicity study. Anal. Biochem. 345: 55–65 (2005). 96. Sugita, T., Yoshikawa, T., Mukai, Y., Yamanada, N., Imai, S., Nagano, K., Yoshida, Y., Shibata, H., Yoshioka, Y., Nakagawa, S., Kamada, H., Tsunoda, S. I., and Tsutsumi, Y. Comparative study on transduction and toxicity of protein transduction domains. Br. J. Pharmacol. 153: 1143–1152 (2008). 97. Tunnemann, G., Ter-Avetisyan, G., Martin, R. M., Stockl, M., Herrmann, A., and Cardoso, M. C. Live-cell analysis of cell penetration ability and toxicity of oligoarginines. J. Pept. Sci. 14: 469–476 (2008). 98. Ellerby, H. M., Arap, W., Ellerby, L. M., Kain, R., Andrusiak, R., Rio, G. D., Krajewski, S., Lombardo, C. R., Rao, R., Ruoslahti, E., Bredesen, D. E., and Pasqualini, R. Anti-cancer activity of targeted pro-apoptotic peptides. Nat. Med. 5: 1032–1038 (1999). 99. Kolonin, M. G., Saha, P. K., Chan, L., Pasqualini, R., and Arap, W. Reversal of obesity by targeted ablation of adipose tissue. Nat. Med. 10: 625–632 (2004). 100. Law, B., Quinti, L., Choi, Y., Weissleder, R., and Tung, C. H. A mitochondrial targeted fusion peptide exhibits remarkable cytotoxicity. Mol. Cancer Ther. 5: 1944– 1949 (2006). 101. Nakase, I., Niwa, M., Takeuchi, T., Sonomura, K., Kawabata, N., Koike, Y., Takehashi, M., Tanaka, S., Ueda, K., Simpson, J. C., Jones, A. T., Sugiura, Y., and Futaki, S. Cellular uptake of arginine-rich peptides: roles for macropinocytosis and actin rearrangement. Mol. Ther. 10: 1011–1022 (2004). 102. Nishimura, S., Takahashi, S., Kamikatahira, H., Kuroki, Y., Jaalouk, D. E., O’Brien, S., Koivunen, E., Arap, W., Pasqualini, R., Nakayama, H., and Kuniyasu, A. Combinatorial targeting of the macropinocytotic pathway in leukemia and lymphoma cells. J. Biol. Chem. 283: 11752–11762 (2008). 103. Watkins, C. L., Brennan, P., Fegan, C., Takayama, K., Nakase, I., Futaki, S., and Jones, A. T. Cellular uptake, distribution and cytotoxicity of the hydrophobic cell penetrating peptide sequence PFVYLI linked to the proapoptotic domain peptide PAD. J. Control. Release 140: 237–244 (2009). 104. Orii, K. O., Grubb, J. H., Vogler, C., Levy, B., Tan, Y., Markova, K., Davidson, B. L., Mao, Q., Orii, T., Kondo, N., and Sly, W. S. Defining the pathway for Tatmediated delivery of beta-glucuronidase in cultured cells and MPS VII mice. Mol. Ther. 12: 345–352 (2005). 105. Zhang, X. Y., Dinh, A., Cronin, J., Li, S. C., and Reiser, J. Cellular uptake and lysosomal delivery of galactocerebrosidase tagged with the HIV Tat protein transduction domain. J. Neurochem. 104: 1055–1064 (2008). 106. El-Andaloussi, S., Holm, T., and Langel, U. Cell-penetrating peptides: mechanisms and applications. Curr. Pharm. Des. 11: 3597–3611 (2005). 107. Fischer, R., Fotin-Mleczek, M., Hufnagel, H., and Brock, R. Break on through to the other side-biophysics and cell biology shed light on cell-penetrating peptides. Chembiochem. 6: 2126–2142 (2005). 108. Foged, C. and Nielsen, H. M. Cell-penetrating peptides for drug delivery across membrane barriers. Expert Opin. Drug Deliv. 5: 105–117 (2008).

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109. Futaki, S., Nakase, I., Tadokoro, A., Takeuchi, T., and Jones, A. T. Arginine-rich peptides and their internalization mechanisms. Biochem. Soc. Trans. 35: 784–787 (2007). 110. Gump, J. M. and Dowdy, S. F. TAT transduction: the molecular mechanism and therapeutic prospects. Trends Mol. Med. 13: 443–448 (2007). 111. Nakase, I., Takeuchi, T., Tanaka, G., and Futaki, S. Methodological and cellular aspects that govern the internalization mechanisms of arginine-rich cellpenetrating peptides. Adv. Drug Deliv. Rev. 60: 598–607 (2008). 112. Patel, L. N., Zaro, J. L., and Shen, W. C. Cell penetrating peptides: intracellular pathways and pharmaceutical perspectives. Pharm. Res. 24: 1977–1992 (2007). 113. Al-Taei, S., Penning, N. A., Simpson, J. C., Futaki, S., Takeuchi, T., Nakase, I., and Jones, A. T. Intracellular traffic and fate of protein transduction domains HIV-1 TAT peptide and octaarginine. Implications for their utilization as drug delivery vectors. Bioconjug. Chem. 17: 90–100 (2006). 114. Fretz, M. M., Penning, N. A., Al-Taei, S., Futaki, S., Takeuchi, T., Nakase, I., Storm, G., and Jones, A. T. Temperature-, concentration- and cholesterol-dependent translocation of l- and d-octa-arginine across the plasma and nuclear membrane of CD34+ leukaemia cells. Biochem. J. 403: 335–342 (2007). 115. Watkins, C. L., Schmaljohann, D., Futaki, S., and Jones, A. T. Low concentration thresholds of plasma membranes for rapid energy-independent translocation of a cell penetrating peptide. Biochem. J. 420: 179–189 (2009). 116. Duchardt, F., Fotin-Mleczek, M., Schwarz, H., Fischer, R., and Brock, R. A comprehensive model for the cellular uptake of cationic cell-penetrating peptides. Traffic 8: 848–866 (2007). 117. Kosuge, M., Takeuchi, T., Nakase, I., Jones, A. T., and Futaki, S. Cellular internalization and distribution of arginine-rich peptides as a function of extracellular peptide concentration, serum, and plasma membrane associated proteoglycans. Bioconjug. Chem. 19: 656–664 (2008). 118. Watkins, C. L., Schmaljohann, D., Futaki, S., and Jones, A. T. Low concentration thresholds of plasma membranes for rapid energy-independent translocation of a cell-penetrating peptide. Biochem. J. 420: 179–189 (2009). 119. Deshayes, S., Morris, M., Heitz, F., and Divita, G. Delivery of proteins and nucleic acids using a non-covalent peptide-based strategy. Adv. Drug Deliv. Rev. 60: 537– 547 (2008). 120. Edenhofer, F. Protein transduction revisited: novel insights into the mechanism underlying intracellular delivery of proteins. Curr. Pharm. Des. 14: 3628–3636 (2008). 121. Murriel, C. L. and Dowdy, S. F. Influence of protein transduction domains on intracellular delivery of macromolecules. Expert Opin. Drug Deliv. 3: 739–746 (2006). 122. Wadia, J. S. and Dowdy, S. F. Transmembrane delivery of protein and peptide drugs by TAT-mediated transduction in the treatment of cancer. Adv. Drug Deliv. Rev. 57: 579–596 (2005). 123. Foerg, C. and Merkle, H. P. On the biomedical promise of cell penetrating peptides: limits versus prospects. J. Pharm. Sci. 97: 144–162 (2008).

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124. Leifert, J. A. and Whitton, J. L. “Translocatory proteins” and “protein transduction domains”: a critical analysis of their biological effects and the underlying mechanisms. Mol. Ther. 8: 13–20 (2003). 125. Trehin, R. and Merkle, H. P. Chances and pitfalls of cell penetrating peptides for cellular drug delivery. Eur. J. Pharm. Biopharm. 58: 209–223 (2004). 126. Wagstaff, K. M. and Jans, D. A. Protein transduction: cell penetrating peptides and their therapeutic applications. Curr. Med. Chem. 13: 1371–1387 (2006). 127. Ratner, L., Haseltine, W., Patarca, R., Livak, K. J., Starcich, B., Josephs, S. F., et al. Complete nucleotide sequence of the AIDS virus, HTLV-III. Nature 313: 277–284 (1985). 128. Elliott, G. and O’Hare, P. Intercellular trafficking and protein delivery by a herpesvirus structural protein. Cell 88: 223–233 (1997).

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CHAPTER 16

Cargo Transport by Teams of Molecular Motors: Basic Mechanisms for Intracellular Drug Delivery MELANIE J. I. MÜLLER, FLORIAN BERGER, STEFAN KLUMPP, and REINHARD LIPOWSKY Department of Theory & Bio-Systems, Max Planck Institute of Colloids and Interfaces, Potsdam, Germany

16.1

INTRODUCTION

16.1.1 Barriers to Gene Therapy and Drug Delivery The delivery of drug carriers or viral and nonviral vectors for gene therapy is hampered by many extracellular and intracellular barriers. The drug or DNA carrier must find its way from the injection site to the target cell, avoiding sequestration and destruction, for example, by cells of the immune system. Upon reaching the target cell, the carrier must pass the plasma membrane to enter the cell, traverse the cytoplasm to its intracellular destination, possibly enter a target organelle like the nucleus or a mitochondrion, and then perform its function. Traditionally, research has focused on overcoming membrane barriers: how to get the drug carriers or transfection vectors into the cell and, once inside the cell, how to get these particles into the target organelle. However, the cytoplasm itself is also a major obstacle to delivery, especially for nonviral gene therapy: After a transfection vector has been released into the cytoplasm, it may have difficulties reaching the nucleus [1, 2]. 16.1.2 Inefficient Transport Via Diffusion One fact that is often overlooked is that drug or gene carriers are too large to diffuse through the cytoplasm at a reasonable rate. The cytoplasm is a crowded, Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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viscous medium consisting of proteins, filament meshworks, and organelles, which limit the diffusion of large particles [3]. For example, using experimentally determined cytoplasmic diffusion coefficients of DNA [4], a 1000-bp DNA needs about 30 min to diffuse a distance of 10 μm, compared to about 18 s in water. DNA strands that are larger than 2000 bp are practically immobile in the cytoplasm. Gene vectors used in gene transfer have typically several thousand base pairs. Consequently, microinjected plasmids do not diffuse far from their site of injection [5], and transfection increases dramatically if DNA is microinjected into or very close to the nucleus instead of a cytoplasmic location far from the nucleus [5, 6]. Therefore efficient transport to the nucleus cannot be achieved by diffusion and requires an active mechanism. 16.1.3 Viruses as Hijackers Some viruses face barriers similar to gene therapy: they must enter the host cell and deliver their genetic material to the nucleus. Many viruses have solved the problem of intracellular transport by hijacking the host cell’s active transport system [2, 7, 8], as shown in Figure 16.1A. After entering the cell via endocytosis or membrane fusion, the virus particle recruits molecular motors, which act as cellular “nanotrucks” and pull the virus along microtubule filaments toward the nucleus. Similarly, when leaving the cell, the virus uses molecular motors to travel from the site of assembly to the cell membrane. This active transport allows the virus to travel at a velocity of about 1 μm/s [8,

(A)

(B) entry

egress cargo dynein

tubule micro actin

MTOC

kinesin

nucleus 25nm delivery

microtubule

Figure 16.1. Intracellular trafficking. (A) Microtubule filaments (black) form long “highways” from the microtubule-organizing center (MTOC) near the nucleus to the cell periphery, while a dense meshwork of actin filaments (cyan) provides “side roads” for short-range traffic. Plus-end directed (blue) and minus-end directed (yellow) molecular motors pull various cellular cargoes along these filaments. Viruses hijack the cellular transport machinery: after entry into the cell, a virus particle (red) recruits motors from the cytoplasm to travel along microtubules toward the nucleus, where it delivers its genetic material (white). In order to leave the cell, the newly assembled virus uses motors to travel back to the plasma membrane. (B) Cargo particle (red) that is transported along a microtubule by one kinesin-1 (blue) and one cytoplasmic dynein (yellow) motor. (See color insert.)

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9] so that it can overcome a distance of 10 μm in about 10 s. For comparison, diffusion of a 100-nm virus capsid in the cytoplasm over the same distance would take a few hours. 16.1.4 Understanding Intracellular Transport Hijacking the cellular transport machinery is an elegant solution to the problem of transport through the crowded cytoplasm, which is employed by many viruses, such as human immunodeficiency virus, herpes simplex virus, and adenoviruses [2, 7, 8]. These viruses are commonly used in viral gene therapy, in which the gene vector is packaged into a virus capsid. However, also nonviral vectors and drug carriers can bind to molecular motors via adaptor proteins recruited from the cytoplasm, and then travel actively along microtubule filaments [2]. In order to improve gene and drug delivery, it is therefore essential to design carrier systems that can recruit the cellular transport machinery in an appropriate way. As a prerequisite for such a design, a good understanding of intracellular transport by molecular motors is required. 16.1.5 Overview This chapter is organized as follows. The basic features of molecular motors and cytoskeletal filaments are briefly described in Section 16.2.1. The challenges encountered by motor transport are summarized in Section 16.2.2. The different mechanisms for cooperative transport by motor teams are discussed in Section 16.3. In this latter section, we will distinguish three such mechanisms that we have recently elucidated by the construction and analysis of explicit theoretical models: (1) cargo transport by one team of identical motors [10, 11]; (2) transport by two antagonistic teams of motors that both work on the same type of filament [12, 13]; and (3) cargo transport by two teams of motors that work on different filaments [14]. At the end, we give a brief outlook on open questions and possible applications of the teamwork of molecular motors.

16.2

INTRACELLULAR TRAFFIC OF MOLECULAR MOTORS

16.2.1 Molecular Motors and Their Tracks The complex internal structure of cells depends to a large extent on active transport: vesicles, as well as RNA, filaments, and protein complexes, move between different compartments, travel from the cell center to the cell periphery or vice versa [15]. This active transport is mainly accomplished by molecular motors. These molecular motors can be compared to road trucks: they travel along a “road network” consisting of cytoskeletal filaments and consume ATP as molecular “fuel.”

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16.2.1.1 Cytoskeletal Filaments The road network for the molecular motors is formed by the cytoskeleton. The cytoskeleton is a meshwork built up from three types of filaments—microtubule, actin, and intermediate filaments—which form a complex meshwork throughout the cell. Only microtubules and actin filaments are used in motor transport. Kinesin and dynein motors walk on microtubules, while myosin motors travel along actin filaments. The microtubules form a very structured network spanning the whole cell from the center to the periphery, see Figure 16.1A, and serve as “highways” for the long-range traffic. The actin filaments, on the other hand, form a dense meshwork concentrated near the cell cortex and serve as “side-roads” for short-range traffic. Both types of filaments are polar: their building blocks are asymmetric so that each filament possesses an intrinsic directionality. Each filament has a “plus end,” at which it polymerizes faster than at its “minus end.” 16.2.1.2 Cytoskeletal Motors Each cytoskeletal motor works on a certain filament and recognizes the polarity of this filament. Thus each motor walks either to the filament’s “plus” or “minus” end. The cytoplasmic dynein motor, for example, walks to the microtubule minus end, while the kinesin-1 motor walks to the microtubule plus end; see Figure 16.1B. 16.2.1.3 Kinesin-1 The best-studied motor, kinesin-1, was first identified in the 1980s in a biochemical fractionation of squid axonal tissue [16, 17]. It is a heterotetramer composed of two identical heavy chains and two identical light chains [18]. The heavy chains dimerize to form an α-helical coiled-coil stalk. Each of the 120-kDa heavy chains includes a globular “head” at its N terminus, with both a microtubule-binding domain and an ATP-binding domain. The two heads perform the actual motor activity of the protein. The light chains associate with the heavy chains near the C terminus and form the motor tail, which is involved in cargo binding. 16.2.1.4 Kinesin Stepping When traveling along a microtubule, kinesin-1 rather “steps” than “drives” [19]. A microtubule filament consists of 13 protofilaments aligned in parallel, each of which provides a track for the motor with a binding site every 8 nm. For each step, kinesin moves the trailing head forward to the next binding site in front, while the other head remains bound to the microtubule. In this so-called hand-over-hand mechanism, each head covers a distance of 16 nm, while the center-of-mass of the motor makes a forward step of 8 nm. During each step, the motor hydrolyzes a single ATP molecule. 16.2.1.5 Cytoplasmic Dynein The structure of cytoplasmic dynein, which was discovered as a minus end microtubule motor in the 1980s [20], is more complex than that of kinesin [18, 21], as indicated in Figure 16.1B. Dynein also has two heads, which can bind to the microtubule, but its ATP-binding

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domains reside in rings of seven globular domains, which are connected to the heads by a short coiled-coil stalk. Several light and intermediate chains are attached near the cargo binding end of dynein and are involved in the binding of cargo and of accessory proteins, making dynein a huge multisubunit protein of 1.2 MDa. 16.2.1.6 Other Types of Motors Kinesin-1 and cytoplasmic dynein are the most prominent members of a whole “zoo” of molecular motors, consisting of three motor families—kinesins, dyneins, and myosins—as reviewed in Refs. 18, 22, and 23. While the dynein family consists of only two subclasses, the kinesin family has more than ten subclasses. All dyneins walk to the microtubule minus end, whereas most kinesins walk to the microtubule plus end. Several classes of myosins mediate transport on actin filaments. 16.2.2 Challenges for Motor Transport The various motors described in the last section provide a versatile toolbox for intracellular traffic [18]. These motors transport an enormous amount of vastly different cargoes through the cell, ranging from endosomes and other types of vesicles to RNAs and filaments as well as to whole organelles such as mitochondria [15]. Each cargo has to recruit the right type of motor and an appropriate motor number in order to be shuttled to the correct destination. In this section, we will discuss how the motors perform these challenging tasks. 16.2.2.1 Motor Processivity Native motor molecules have a linear extension of about 100 nm; see Figure 16.1B. Because of their relatively small size, their motion is strongly affected by thermal fluctuations and viscous forces: molecular motors continuously collide with other molecules and thus walk “in a strong storm.” Because of this thermal noise, molecular motors unbind from their track from time to time. Upon unbinding they lose their ability to perform directed motion and randomly diffuse in the surrounding solution until they finally rebind to a filament. For kinesin-1, unbinding happens on average after a run time of about 1 s, or equivalently a run length (or walking distance) of about 1 μm. A cellular cargo, however, must accomplish distances of tens of micrometers, and in some extremely large cells like neurons even up to a meter [24]. 16.2.2.2 Load Forces Molecular motors can generate forces in the range of a few piconewtons [22, 25, 26]. This has to be compared to the frictional force, Ffr = φfr v with φfr = 6πηR = kBT D

(16.1)

experienced by the cargo particle of size R as it moves through a medium with dynamical viscosity η. The parameter D is the diffusion coefficient of the

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(A) Fibroblast cell

(B) Epithelial cell apical

(C) Neuronal cell dendrite

basolateral

(mixed orientation)

unipolar orientation cell body

axon

synapse

Figure 16.2. Organization of microtubules (black lines with indicated plus ends) in three different cell types for which the nuclei are represented as black ellipses and the microtubule-organizing centers or centrosomes as white rectangles. (A) In fibroblast cells, microtubule minus ends nucleate at the centrosome and the plus ends point radially outwards. (B) In epithelial cells, microtubules form a parallel array with minus ends pointing to the apical and plus ends to the basolateral surface. (C) In long axons of neuron cells, microtubules form an isopolar array with the plus ends oriented toward the synapses at the axon tips.

(unbound) cargo, and kBT is Boltzmann’s constant times absolute temperature. In water with dynamical viscosity η = 1 mPa · s, the force needed to pull a 1-μm sized cargo at a velocity of v = 1 μm/s is equal to 0.01 pN, which is negligible for a molecular motor that is able to generate several piconewtons of force. However, since diffusion in the cytoplasm is slowed down by a factor of 10–1000 depending on the cargo size [47], it follows from Equation 16.1 that frictional forces in the cytoplasm can be in the range of 0.1–10 pN, which represents a large load for a molecular motor. For these estimates, we assumed that the diffusion coefficient D as measured for cargo diffusion in the cytoplasm also applies to motor transport; see, however, Ref. 27. 16.2.2.3 Bidirectional Transport Cells typically have an “isopolar” microtubule cytoskeleton [28]; see Figure 16.2. In a “round” cell such as a fibroblast cell, for example, microtubules are arranged radially, with their minus ends close to the nucleus at the cell center and their plus ends directed outward to the cell periphery. In epithelial cells, microtubules form a parallel array with their minus ends pointing to the apical and their plus ends to the basolateral surface. The isopolar microtubule arrangement is most pronounced in long cellular subcompartments such as neuronal axons or fungal hyphae. In these long and relatively narrow membrane tubes, the microtubules are arranged parallel to the tube axis with their minus ends pointing to the cell body and the plus ends pointing to the tip. Single molecular motors walk along these tracks only into one direction: dyneins to the minus ends, and most kinesins to the plus ends. However, many cellular cargoes travel in both directions [29, 30], a transport mode that avoids accumulation of cargo at either the cell periphery or the cell center [31, 32]. Cargoes like endosomes, mitochondria, and viruses are observed to move bidirectionally, reversing direction every few seconds [29, 30]. These cargoes have both plus- and minus-end directed motors attached. Bidirectional

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motion can lead to net transport if the runs in one direction are on average longer or faster than runs in the other direction. Therefore bidirectional transport requires an appropriate balance between the activity of plus and minus motors. 16.2.2.4 Transport Regulation Cellular regulation of cargo transport often leads to changes in the run lengths, that is, the distances covered during runs into the plus or the minus direction. This has been observed for the directional regulation of mitochondria in axons [33], of lipid droplets during embryo development [34], of virus targeting during entry and egress [35], and of melanosomes during dispersion and aggregation [36]. In the latter example, melanosome redistribution in specialized pigment cells of fish or frog allows the organism to adapt its skin color to the environment. During “aggregation,” minus-end motion dominates, and the melanosomes accumulate in the cell center. During “dispersion,” the melanosomes spread out over the whole cell because of a decrease in minus run length [36]. Aggregation and dispersion are triggered by hormonal stimulation, which is transmitted to the motors via a signal cascade involving cAMP and protein kinase A (PKA) [37]. 16.2.2.5 Transport on Two Types of Tracks Intracellular transport usually proceeds in two steps [38, 39]; see Figure 16.1A. Long-range transport from the cell center to the periphery and vice versa is mediated by kinesins and dyneins walking along the long microtubule highways, while short-range delivery is the task of myosin motors using the dense actin meshwork in the cell cortex. This “dual transport” requires that cargoes are able to switch from microtubule to actin tracks, as has indeed been observed for various cellular cargo such as mitochondria, pigment granules, and synaptic vesicles, reviewed in Refs. 38 and 39. These cargoes need to balance the activities of actin- and microtubule-based motors. A prominent example is again the traffic of melanosomes, which are carried by kinesin-2 and cytoplasmic dynein along microtubules and by myosin V along actin filaments [38]. During dispersion, myosin V is upregulated, which leads to enhanced switching of melanosomes to actin filaments [36, 40]. During aggregation, downregulation of myosin V “recollects” the melanosomes to the microtubules, on which they are then transported toward the cell center.

16.3

TEAMWORK OF MOLECULAR MOTORS

16.3.1 Different Levels of Motor Motility In general, the behavior of molecular motors involves several levels of motility [41] related to (i) the stepping of single motors, (ii) the cooperative transport of cargo by motor teams, and (iii) the traffic of many motor-propelled cargoes. At the level (i) of a single motor, most progress has been made in understand-

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ing the structure and the stepping mechanism of kinesin-1, in particular, the coordination of its two heads [19, 21, 42]. The level (iii) of cell wide cargo trafficking has received mainly theoretical attention. On the cellular length scale, each cargo is viewed in a course-grained manner as a particle with the ability to bind and actively move along cytoskeletal filaments, while diffusing randomly in the surrounding solution. This description allows one to calculate quantities such as the densities and fluxes of cargoes throughout the cell [43–46], which are important to characterize the intracellular traffic. For drug delivery, gene therapy, or virus trafficking, another important quantity is the mean first passage time from the point of entry into the cell to the intracellular target [47, 48].

16.3.2 Motor Teams The level (ii) of cooperative transport by several molecular motors will be the focus of this chapter. Many cellular cargoes are attached to more than one motor, and to more than one type of motor [29, 30, 38, 39]. The number of motors in a team is small, typically between 1 and 10 [27]. Cargo transport by small teams of molecular motors is a topic of current research both experimentally [27, 49–54] and theoretically [11, 13, 14, 50, 55, 56]. In the following, we will review our recent theoretical studies of cargo transport by small teams of molecular motors [11–14]. After introducing our model for a single motor in the next section, we will first examine the transport by one cooperating team of molecular motors that all belong to the same species. We will then discuss transport by two antagonistic teams of molecular motors that walk into opposite directions, and finally transport by two teams of motors that can walk on different types of filaments.

16.3.3 Transport Properties of a Single Motor 16.3.3.1 Different Subprocesses of Motor Transport Since we are interested in cargo motion on time scales ranging from seconds to minutes and length scales from a few to hundreds of micrometers, we can describe the motors in a coarse-grained manner. We neglect the details of their protein structure and stepping cycle but take their finite binding time to the track into account. The motors exhibit three basic subprocesses: they walk along the filament with velocity v, unbind from this filament with rate ε, and rebind to it with rate π; see Figure 16.3A. If the motor experiences no load force, these parameters are of the order of π0 ∼ 1/s, vf ∼ 1 μm/s, and ε0 ∼ 1/s, respectively; see Table 16.1. If the motor has to work against a load force F, both the motor velocity v and its unbinding rate ε strongly depend on this force; see Figure 16.3B,C. This force dependence has been investigated experimentally for some motors using optical traps or tweezers. An optical trap uses a focused laser beam, the radiation pressure of which is able to exert forces in the piconewton-range on dielectric objects of nanometer to micrometer size. If a

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v(F)

π0

ε(F)

Vf

vf(1–F/Fs)

0.5 Fs 0 0

(A)

vb(1–F/Fs)

(B)

2 4 6 8 10 12 load force F [pN]

unbind. rate ε [1/s]

F

velocity v [μm/s]

TEAMWORK OF MOLECULAR MOTORS

(C)

8

ε0exp[F/Fd]

6 4

ε0

2 0

297

0

2 4 6 load force F [pN]

Figure 16.3. Transport properties of a single motor. (A) A single motor walks along the filament with velocity v, unbinds from the filament with rate ε, and rebinds to it with rate π = π0. If the cargo experiences a load force F, both the velocity v and the unbinding rate ε depend on F. (B) Motor velocity v as a function of load force F as parameterized in Equations 16.2 and 16.3. The velocity decreases with increasing F until it vanishes at the stall force Fs. For superstall forces F > Fs, the motor steps backwards with a relatively small velocity. (C) Motor unbinding rate ε as a function of F as in Equation 16.4. The unbinding rate increases exponentially with increasing F. (See color insert.) TABLE 16.1 Single Motor Parameters for Conventional Kinesin or Kinesin-1 and Cytoplasmic Dyneina Parameter Stall force Detachment force Unbinding rate Binding rate Forward velocity Backward velocity

Symbol

Kinesin

Dynein

Fs Fd ε0 π0 vf vb

6 pN [25, 7, 58] 3 pN [59] 1/s [59] 5/s [10, 52] 1 μm/s [25, 7, 58] 6 nm/s [25, 7]

7 pN [26] 3 pN ? 0.25/s [60] 1.5/s [60] 1 μm/s [26, 60] 6 nm/s ?

a

These are abbreviated here as “kinesin” and “dynein,” respectively. The experimentally determined values for the stall force Fs of kinesin [25, 7, 58] vary between 5.5 pN and 7.6 pN. For dynein, the parameters with a question mark are currently not available from experimental studies; in the latter case, we used the same parameters for dynein as for kinesin.

single motor is bound to such an object, the movement of this motor can be studied under a constant load force F. 16.3.3.2 Single Motor Parameters The results of optical trapping experiments imply that the motor velocity v decreases approximately linearly with force F and is well described by the functional form v ( F ) = vf (1 − F Fs ) for 0 ≤ F ≤ Fs

(16.2)

which vanishes at the “stall force” F = Fs [25, 59]. For higher load forces F > Fs, the motor walks backwards very slowly [25, 57]; we use the parametrization [13] v ( F ) = vb(1 − F Fs ) for Fs ≤ F

(16.3)

with vb << vf.

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The unbinding rate of the motor from the filament increases exponentially with applied force F according to [11] ε ( F ) = ε 0 exp ( F Fd )

(16.4)

with the detachment force Fd as obtained, for positive F, from the measurements of the walking distance of a single motor as a function of load [59] in agreement with Kramers’ rate theory [61]. Such an exponential form has also been used in the context of receptor-ligand binding at membranes [62]. The binding rate of the motor to the microtubule is taken to be independent of the load force and given by [11] π (F ) = π0

for all F .

(16.5)

In total, a single motor of a certain motor species is described by six parameters as explained in Figure 16.3. For the plus motor kinesin-1 and the cytoplasmic minus motor dynein, most of these single motor parameters have been determined experimentally; the corresponding parameter values are summarized in Table 16.1 [13]. This theoretical description for a single motor incorporates all results of single molecule experiments that are relevant for large-scale cargo transport. 16.3.4 One Team of Identical Motors In this section, we consider cargo transport by one team of motors, that is, N motors of the same type are attached to one cargo as shown in Figure 16.4. Since each motor binds to and unbinds from the filament in a stochastic manner, the number n of motors that crosslink the cargo to the filament and thus can actively pull on it varies with time between n = 0 and n = N as illustrated in Figure 16.4. The motors are taken to act independently. If n motors are bound to the filament in the absence of load, the unbinding of one of these

Figure 16.4. Transport by one team of identical motors. Each motor is characterized by a long stalk (grey), by which it is firmly attached to the cargo particle (red), and by two motor heads (blue), which can bind to the filament and then actively pull on the cargo. In this example, the total number of motors is N = 3 but the actual number n of crosslinking motors fluctuates between n = 0 and n = 3 because of thermally induced motor unbinding and rebinding. (See color insert.)

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(A)

65 μm

100 10

1

311 μm

Velocity [μm/s]

Run length Δxb[μm]

1000

14 μm 3.5 μm 1 μm

1 0

1 2 3 4 Motor number N

5

6 (B)

0.8 N=10

0.6

N=

0.4 0.2 0 0

5

N=1 10 20 30 40 50 Stokes friction φfr [pNs/μm]

Figure 16.5. Transport by one team of identical motors. (A) The cargo’s run length Δxb increases exponentially with the total number N of motors attached to the cargo. (B) The average cargo velocity decreases with increasing friction coefficient φfr, but less so for larger motor number N. The single motor parameters correspond to kinesin-1 as given in Table 16.1.

bound motors is governed by the unbinding rate nε0, and the rebinding of one of the N − n unbound motors back to the filament takes place with rate (N − n)π0 [11]. 16.3.4.1 Processivity Enhancement A major problem of cargo transport by a single motor is the motor’s finite run length Δxb ∼ 1 μm, which is too small to cover typical cellular distances of tens of micrometers. Cargo transport by several molecular motors leads to a dramatic increase in the cargo’s run length; see Figure 16.5A. Intuitively, if one motor unbinds from the filament, the cargo remains attached to the filament as long as it is still crosslinked by another motor, giving the unbound motor a chance to rebind; compare Figure 16.4. An explicit calculation shows that the average run length Δxb is given by [11] Δxb =

vf ⎡(1 + π 0 ε 0 )N − 1⎤⎦ Nπ 0 ⎣

(16.6)

that is, it increases exponentially with the motor number N for large N. If the cargo is transported by kinesin-1 motors, only three such kinesins are required to cross a cell of 10-μm diameter; see Figure 16.5A. The increase of run length with increasing motor number has been observed in vitro [63, 64], but it has been difficult to determine the number of motors pulling the cargo. In recent experiments, the motor number was determined by force measurements [54] or dynamic light scattering [10]. In the latter case, the theory described here was used to describe the data quantitatively. 16.3.4.2 Frictional Forces Another feature of cargo transport by several motors is the ability to sustain large forces arising, for example, from hydrodynamic friction. For a cargo particle of radius R = 1 μm in water with

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dynamical viscosity η = 1 mPa·s, the friction coefficient is φfr ∼ 10−2 pN · s/μm, which does not significantly change the velocity of a cargo particle that is transported by a single motor; see the curve for N = 1 in Figure 16.5B. However, as discussed in Section 16.2.2, particle diffusion in the crowded cytoplasm is reduced by a factor of 10–1000 [3], depending on the size of the particle. Since the friction coefficient in Equation 16.1 is inversely proportional to the diffusion constant, the cytoplasmic friction coefficient is 10–1000 times larger than the one in water and leads to a significant reduction of the cargo velocity if the cargo particle is pulled by only one motor; see Figure 16.5B. When the cargo is pulled by N > 1 motors, these N motors can share the force, so that the average cargo velocity remains relatively high even for large friction coefficient φfr; see Figure 16.5B. 16.3.5 Two Teams of Antagonistic Motors As discussed in Section 16.2.2, many cellular cargoes undergo fast transport both toward the plus end and toward the minus end of the microtubules. This bidirectional transport implies that these cargo particles must be attached to both plus-end and minus-end directed motors. 16.3.5.1 Tug-of-War Between Motors We now consider a cargo particle that is attached to N+ plus and N− minus motors. Because each motor unbinds from and rebinds to the filament in a stochastic manner, both the number of active plus motors and the number of active minus motors now fluctuate; see Figure 16.6. Three situations are possible: in the (+) states, only plus motors are active, which can pull the cargo into the plus direction without any opposition from the minus motors; in the (−) states, only minus motors are active and the cargo exhibits fast minus motion; and in the (0) states both types of motors are bound to the filament. In the latter case, the motors pull the cargo into opposite directions, leading to slow cargo movement. Since many bidirectional cargoes are observed to exhibit long periods of fast unidirectional motion, the motors must “cooperate” in some way to avoid the (0) states. Two mechanisms for bidirectional transport have been proposed [29, 30]: (1) coordination by a putative protein complex, which ensures that only one type of motors is active at any given time; and (2) tug-of-war between the motors that pull on each other until the stronger team wins and determines the direction of motion for a certain period of time. Previously, it was thought that a tug-of-war would lead to a prevalence of (0) states with slow cargo motion and thus would be inconsistent with the experimental observation of fast cargo motion [29, 30]. However, we have developed a realistic tug-of-war model, which is based on the single motor description as presented in Section 16.3.3 as well as on the assumptions that (1) each motor team exerts load forces on the other team and (2) motors of one team share the load force generated by the antagonistic motor team [12, 13]. Such a tug-of-war, which

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(–)

(0)

(0)

(–)

(0)

(0)

(0)

(+)

(+)

301

Figure 16.6. Transport by two antagonistic teams of motors. The motors with blue heads are plus motors that pull to the right, the ones with yellow heads are minus motors that pull to the left. Both types of motors are firmly attached to the cargo particle (grey) via their long stalks. The total number of blue plus motors and yellow minus motors is N+ = 2 and N− = 2, respectively. The cargo particle exhibits (N+ + 1)(N− + 1) = 9 different states and undergoes transitions between these states arising from the unbinding and rebinding of single motors. (See color insert.)

does not require any coordination complex, is consistent with all experimental observations [13, 65]. 16.3.5.2 Patterns of Movements As shown in Refs. 12 and 13, a cargo transported by two teams of motors can exhibit different patterns of motility (or “motility states”) depending on the single motor parameters. For the single motor parameters given in Table 16.1, the cargo transport exhibits a dynamic instability, which effectively makes the (0) states very unlikely: if both types of motors are active simultaneously, they exert forces on each other and tend to pull each other off the filament. If, for example, a minus motor unbinds, each of the remaining minus motors has to sustain a larger force arising from the opposing team. Since the unbinding rate in Equation 16.4 increases exponentially with the force, the remaining minus motors now have a high unbinding probability, and undergo an unbinding cascade until all minus motors are detached. A similar unbinding cascade can happen for the plus motors. Because of these unbinding cascades, the cargo is unlikely to stay in a (0) state, in which both types of motors are attached to the filament, but is likely to attain a (+) or (−) state, in which only plus or only minus motors are bound to filament. Therefore the spatial displacement (or trajectory or kymograph) of a cargo pulled by several kinesin-1 and several dynein motors exhibits

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alternating periods of fast plus and fast minus motion at speeds of micrometers/ second (μm/s); see Figure 16.7A. Switching between the two directions happens when a series of motor binding and unbinding events leads to the predominance of the opposing team, which is then stabilized by another unbinding cascade. 16.3.5.3 Advantages for Cargo Transport The tug-of-war performed by two motor species has several advantages for the cell. First, fast bidirectional motion is possible without any coordination complex. Second, cargo transport can easily be regulated, since a change in a single motor parameter affects the competition of the two teams and can lead to net plus or net minus motion by increasing or decreasing the corresponding run lengths, as found experimentally [33–36]. Third, the transport by teams of motors again leads to larger force generation and to enhanced processivity [66]. In particular, the binding time of a cargo increases exponentially with the numbers N+ and N− of attached plus and minus motors, respectively. The tug-of-war model described here leads to the binding time Δtb as given by Δtb( N +, N − ) ≅

(1 + π 0 + ε 0 + ) N + + (1 + π 0 − ε 0 − ) N − − 2

(16.7)

N+ π0+ + N− π0−

where the indices “+” and “−” label plus and minus motor parameters, respectively. Fourth, the stochastic switching of bidirectional transport leads to enhanced diffusion on long time and large length scales; see Figure 16.7B. On short time scales, the cargo particle moves ballistically, that is, the mean square discplacement increases quadratically with time, while it increases linearly on long time scales, with a diffusion coefficient of about 1 μm2/s. This diffusion constant is similar to the diffusion constant of a 1-μm sized cargo particle in

10

MSD [μm2]

Distance [μm]

15 5 0 −5 0 (A)

10

20 30 Time t [s]

40

104 102 10−2 10−4 0.01

(B)

~t

1 ~t2 1 103 Time t [s]

104

Figure 16.7. Transport by two antagonistic teams of motors. (A) Spatial displacement (or trajectory or kymograph) of cargo particle as a function of time. The cargo is pulled by 4 dyneins and 3 kinesins with single motor parameters as in Table 16.1 and exhibits fast transport in both the plus direction and minus direction. (B) Mean square displacement (MSD) of cargo as a function of time. The cargo is pulled by N+ = 4 plus and N− = 4 minus motors, which are both characterized by kinesin parameters as in Table 16.1. The cargo’s MSD grows quadratically and linearly with time for short and long times, respectively.

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water, and about 100–1000 times larger than the diffusion constant of such a particle in the cytoplasm. This enhanced diffusion should be especially useful for cargoes in search of their destination, or for cargo particles such as mitochondria or pigment granules that must be distributed over the whole cell [29, 36, 67]. Finally, one would intuitively expect that bidirectional transport helps to reduce jams in the dense traffic of cargo particles as found in eukaryotic cells. Indeed, if a jam builds up in one direction, for example, because of an obstacle, the cargo particles at the very end of the jam may then move in the opposite direction and, in this way, start to dissolve the jam. 16.3.6 Two Teams of Motors Working on Different Tracks As discussed in Section 16.2.2, tracks for the long- and short-ranged transport of intracellular cargo particles are provided by microtubules and actin filaments, respectively. In order to switch between these two filament systems without interruptions, many cargo particles are attached to both microtubuleand actin-based motors [38, 39]. The probability of switching between the two filaments depends on the number and types of motors, on the cargo, as well as on cellular regulation [36, 40, 68]. 16.3.6.1 Processivity Enhancement During transport on one type of filament, both types of motors may be crosslinked to this filament. Indeed, myosin V, which walks along actin filaments, can also bind to microtubules and diffuse randomly on these latter filaments, whereas kinesin-1, which walks along microtubules, exhibits a weak affinity for actin filaments as well [49, 69]. This type of transport is illustrated in Figure 16.8A for a cargo particle that is

Cargo Myosin

Kinesin

-

Distance [μm]

6

+

4

2

0 0

(A)

Microtubule

(B)

3

6

9

12

Time t [s]

Figure 16.8. Cargo transport by one active and one “passive” motor. (A) A cargo particle (grey) is crosslinked to a microtubule by one kinesin-1 (blue) and one myosin V (red). The kinesin-1 motor is actively stepping whereas the myosin V motor is passively diffusing along the filament. (B) The trajectory of such a cargo exhibits fast plus motion interrupted by diffusive events. For this example, the total binding time was about 11 s, much longer than in the absence of the “passive” myosin V motor [14]. (See color insert.)

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crosslinked to a microtubule by both one kinesin-1 motor, which actively walks along the microtubule, and one myosin V, which passively diffuses along this filament. Such a cargo particle exhibits fast plus motion interrupted by diffusive events; see Figure 16.8B. It is then necessary to distinguish the run length, which the cargo particle exhibits during its plus motion, from the binding length of the composite cargo motion. The binding length of a cargo particle that is transported by one kinesin-1 motor and one myosin V motor is more than two times larger than that of a cargo particle transported by only one kinesin motor [49]. This processivity enhancement can be understood from the single motor properties of kinesin and myosin (compare Section 16.3.3), provided the myosin motor is characterized by its bound diffusion constant rather than by its velocity [14]. The increase in run length arises in a similar way as for cargo transport by a team of identical motors; see Section 16.3.4. If the kinesins unbind, the myosins still act as crosslinkers between cargo and microtubules, thereby preventing the cargo from diffusing away from the filament and giving the kinesins a chance to rebind. Furthermore, in contrast to the competition between kinesin and dynein as described in Section 16.3.5, the simultaneous crosslinking by kinesin-1 and myosin V does not lead to a tug-of-war: because of the relatively weak affinity of myosin V to microtubules, myosin V motors can easily be dragged along by kinesin motors toward the plus end of the microtubule [14].

16.4

CONCLUSION AND DISCUSSION

Cellular cargoes, including endosomes, RNAs, protein complexes, and filaments as well as whole organelles, travel through the cell with the help of molecular motors that pull them along cytoskeletal filaments: kinesin and dynein motors mediate long-ranged transport on microtubule filaments, while local delivery is accomplished by myosin motors walking on a meshwork of actin filaments. 16.4.1 Team Work of Motors As described in this chapter, molecular motors work in teams in order to meet the challenges of intracellular transport. Three types of teamwork have been identified and elucidated. First, one team of identical motors as shown in Figure 16.4 is able to transport large cargoes through the viscous cytoplasm at considerable speeds of micrometers/second over distances of many micrometers; see Figure 16.5. Second, two teams of antagonistic motors that walk into opposite directions, as in Figure 16.6, accomplish bidirectional transport along an isopolar cytoskeletal network of motor tracks as illustrated in Figure 16.7A. Third, two teams of motors that can walk on different filaments (see

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Figure 16.8) allow a smooth transit from the long-range microtubule traffic to the short-range delivery on actin filaments and vice versa. The presence of several motor teams on one cargo allows versatile transport and fast reaction to cellular regulation. A regulation cascade that targets the motor proteins can change the transport properties of the cargo “on the fly,” making it travel on a microtubule to the minus instead of the plus direction by downregulating kinesin, or switching to actin filaments by upregulating myosin. 16.4.2 Virus, Gene, and Drug Delivery Many viruses hijack the intracellular transport systems and use them for their own purposes. These viruses recruit molecular motors from the host’s cytoplasm and, in addition, “know” how to activate these motors to ensure transport to the desired destination. Copying this viral strategy would dramatically increase the efficiency of gene therapy and drug delivery, because transport through the crowded and viscous cytoplasm would be strongly enhanced. Virus-based expression vectors should include all factors that allow the virus to use the host’s transport machinery during entry and transcription, but not those involved in virus replication and egress. Nonviral vectors and drug carriers should be able to recruit motor proteins from the cytoplasm, for example, by coating them with receptor proteins for molecular motors. Although many of the motor–cargo interactions still remain to be elucidated, an increasing catalog of receptor proteins is emerging [8]. Ideally, the appropriate receptors should be selected from such a catalog in order to bind the correct number and types of motors from the cytoplasm to the gene or drug carrier. The choice of motors should reflect the cargo destination, and possible responses to intracellular or externally applied regulatory stimuli. 16.4.3 Outlook: Open Questions and Possible Applications In order to construct such carrier and delivery systems, a good understanding of molecular motor traffic is necessary. Although much progress has been made during the last couple of years, many open questions remain. These include the details of the molecular stepping mechanisms of the different motors, the understanding of motor cooperativity, as well as the understanding of overall cellular traffic. A particularly challenging topic is the coupling of motor transport to cellular regulation. Such an understanding of molecular motor traffic would also be useful for other types of applications in medicine and nanotechnology. One example is provided by antiviral therapies: efficiency of virus infection would be greatly reduced if the viruses were prevented from hijacking the cellular transport machinery in an appropriate way. Likewise, understanding of molecular motor traffic could be useful in order to treat diseases in which improper intracellular transport plays an important role, such as Alzheimer’s disease or lissencephaly

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[70, 71]. Finally, molecular motors are possible building blocks for nanotechnological applications. One example is provided by molecular motors that carry cargoes on lab-on-a-chip devices [72, 73].

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56. Kunwar, A., Vershinin, M., Xu, J., and Gross, S. P. Stepping, strain gating, and an unexpected force-velocity curve for multiple-motor-based transport. Curr. Biol. 18: 1–11 (2008). 57. Nishiyama, M., Higuchi, H., and Yanagida, T. Chemomechanical coupling of the forward and backward steps of single kinesin molecules. Nat. Cell Biol. 4: 790–797 (2002). 58. Visscher, K., Schnitzer, M. J., and Block, S. M. Single kinesin molecules studied with a molecular force clamp. Nature 400: 184–189 (1999). 59. Schnitzer, M. J., Visscher, K., and Block, S. M. Force production by single kinesin motors. Nat. Cell Biol. 2: 718–723 (2000). 60. King, S. J. and Schroer, T. A. Dynactin increases the processivity of the cytoplasmic dynein motor. Nat. Cell Biol. 2: 20–24 (2000). 61. Kramers, H. A. Brownian motion in a field of force and the diffusion model of chemical reactions. Physica 7: 284–304 (1940). 62. Bell, G. I. Models for the specific adhesion of cells to cells. Science 200: 618–627 (1978). 63. Block, S. M., Goldstein, L. S. B., and Schnapp, B. J. Bead movement by single kinesin molecules studed with optical tweezers. Nature 348: 345–352 (1990). 64. Coy, D. L., Wagenbach, M., and Howard, J. Kinesin takes one 8-nm step for each ATP that it hydrolyzes. J. Biol. Chem. 274: 3667–3671 (1999). 65. Welte, M. A. and Gross, S. P. Molecular motors: a trafifc cop within? HFSP J. 2: 178–182 (2008). 66. Müller, M. J. I., Klumpp, S., and Lipowsky, R. Dynamics of bi-directional transport by molecular motors. Biophys. J. 98: 2610–2618 (2010). 67. De Vos, K. J., Sable, J., Miller, K. E., and Sheetz, M. P. Expression of phosphatidylinositol(4,5) bisphosphate-specific pleckstrin homology domains alters direction but not the level of axonal transport of mitochondria. Mol. Biol. Cell 14: 3636–3649 (2003). 68. Ross, J. L., Ali, M. Y., and Warshaw, D. M. Cargo transport: molecular motors navigate a complex cytoskeleton. Curr. Opin. Cell Biol. 20: 41–47 (2008). 69. Ali, M. Y., Krementsova, E. B., Kennedy, G. G., Mahafy, R., Pollard, T. D., Trybus, K. M., and Warshaw, D. M. Myosin Va maneuvers through actin intersections and diffuses along microtubules. Proc. Natl. Acad. Sci. U.S.A. 104: 4332–4336 (2007). 70. Aridor, M. and Hannan, L. A. Traffic jams II: an update of diseases of intracellular transport. Traffic 3: 781–790 (2002). 71. Duncan, J. E. and Goldstein, L. S. The genetics of axonal transport and axonal transport disorders. PLoS Genet. 2: e124 (2006). 72. Hess, H., Bachand, G. D., and Vogel, V. Powering nanodevices with biomolecular motors. Chem. Eur. J. 10: 2110–2116 (2004). 73. van den Heuvel, M. G. L. and Dekker, C. Motor proteins at work for nanotechnology. Science 317: 333–336 (2007).

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CHAPTER 17

The Potential of Photochemical Internalization (PCI) for the Cytosolic Delivery of Nanomedicines KRISTIAN BERG and ANETTE WEYERGANG Department of Radiation Biology, Institute for Cancer Research, Norwegian Radium Hospital, Oslo University Hospital, Oslo, Norway

ANDERS HØGSET PCI Biotech AS, Oslo, Norway

PÅL KRISTIAN SELBO Department of Radiation Biology, Institute for Cancer Research, Norwegian Radium Hospital, Oslo University Hospital, Oslo, Norway

17.1

INTRODUCTION

Utilization of macromolecules in the therapy of cancer and other diseases is becoming increasingly important. Recent advances in molecular biology and biotechnology have made it possible to improve targeting and design of cytotoxic agents or DNA complexes for clinical applications. Macromolecules with therapeutic potential include proteins such as ribosome-inactivating protein toxins for treatment of cancer and other indications, antibodies and growth factors for cell surface targeting, peptides and mRNA for vaccination, DNA utilizing nonviral and viral vectors for gene therapy, and oligonucleotides (antisense oligonucleotides, ribozymes, peptide nucleic acids (PNAs), and siRNA for gene silencing) and nanoparticles for drug delivery [1]. There are many extracellular and intracellular barriers for these molecules to overcome before they can arrive at the target cells, enter the cell, and reach intracellular therapeutic targets. Degradation by serum enzymes and elimination by cells of the reticuloendothelial system (RES), penetration into the target tissues Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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through the endothelial lining, as well as transport limitations within the tissue are important hurdles to obtain sufficient biological effect of these macromolecules [2]. New delivery systems have improved the cellular uptake of macromolecules, but tissue penetration, cellular uptake, and efficient transfer of the molecules into the cytosol of the target cells through the plasma membrane or the membranes of the endocytic vesicles are still fundamental obstacles. In many cases the targets of macromolecular therapeutics are intracellular. However, the limited release from and degradation of macromolecules in endocytic vesicles after uptake by endocytosis are still major intracellular barriers for the therapeutic application of macromolecules having intracellular targets of action. These limitations may also in many cases cause suboptimal therapeutic effect of nanoparticle-based therapeutics. Photochemical internalization (PCI) is a novel technology for release of endocytosed macromolecules into the cytosol [3]. The technology is based on the use of photosensitizers located in endocytic vesicles that upon activation by light induce a release of macromolecules from their compartmentalization in endocytic vesicles. PCI has been shown to potentiate the biological activity of a large variety of macromolecules and other molecules that do not readily penetrate cellular membranes. PCI is a technology derived from photodynamic therapy (PDT). Thus the basis for PDT will be described as well as the basic principles and utilization of PCI for therapeutic purposes.

17.2

PHOTODYNAMIC THERAPY

PDT is a treatment modality that takes advantage of the phototoxic effects induced by a photosensitizer and light in the presence of oxygen [4]. PDT is approved for several cancer indications, as well as for age-related macular degeneration. In addition, fluorescence diagnosis (FD) based on the fluorescing properties of a PDT-type photosensitizer has recently been approved for detection of bladder dysplasia and cancer [5]. 17.2.1 The Photosensitizer A photosensitizer is defined as a chemical entity that, upon absorption of light, induces a chemical or physical alteration of another chemical entity. Most photosensitizers and all clinically approved photosensitizers (with the exception of methylene blue) used in photodynamic therapy (PDT) are based on or related to the tetrapyrrole macrocycle [6]. Porphyrins consist of four pyrrole subunits linked together by four methine bridges as shown in Figure 17.1. Porphyrins and porphyrin-related dyes used in PDT have substituents in the peripheral positions of the pyrrole rings, on the four methine carbons (mesopositions) and/or coordinated metals. These derivatives are synthesized to influence the water/lipid solubility, amphiphilicity, pKa, and stability of the compounds [7]. These parameters determine the biodistribution of the compounds, that is, the intracellular localization, tissue distribution, and

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N N

Porphin

N N

Chlorin

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Bacteriochlorin

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Q-bands Phthalocyanines Bacteriochlorins

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N N

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O

O

N

N N

Phthalocyanine

IV III II I

N N N N

O

313

400

Chlorins

500 600 700 800 Wavelength (nm)

Naphthalocyanine

Figure 17.1. Basic structure (left side) and absorption spectra (right side) of some photosensitizers. A full spectrum is shown for a porphyrin-type photosensitizer and additionally the locations of the main peaks for photosensitizers developed for PDT are shown on the figure (not to scale).

pharmacokinetics. The photosensitizer must absorb the wavelengths emitted by the light source in order to induce a treatment effect. Clinically, 6–800-nm light is mainly used due to the optimal tissue penetration properties at these wavelengths. Porphyrins have low absorption of light in this wavelength region. Thus the porphyrin structures have been modified in order to enhance the absorption properties of the photosensitizers (Figure 17.1). Necrotic depth after PDT of solid tumors is typically 2–10 mm depending on the photosensitizer in use. 17.2.2 Mechanisms of PDT The photosensitizer is usually administered systemically by means of intravenous injection. There is substantial evidence for a preferential retention of the photosensitizers in neoplastic lesions, usually with a tumour : (normal surrounding tissue) ratio of 2–3 : 1 [8]. At some time interval after photosensitizer administration, the target tissue is illuminated with light of an appropriate wavelength. The light source is often a laser (e.g., a diode laser), but noncollimated light sources may also be used. The photosensitizer must absorb the wavelengths emitted by the light source in order to induce a treatment effect (Figure 17.1). The cytotoxic effect is mediated mainly through the formation of singlet oxygen (1O2). This reactive intermediate has a very short lifetime in cells (<0.04 μs) [9]. Thus the primary cytotoxic effect of PDT is executed during light exposure and very close to the sites of formation of 1O2 (<20 nm). Singlet oxygen reacts with and oxidizes proteins (histidine, tryptophan, methionine, cysteine, tyrosine), DNA (guanine), unsaturated fatty acids, and cholesterol. The therapeutic effect of PDT may be exerted through a direct effect on the target cells, via vascular damage or by an immune stimulatory effect, and often a combination of these effects.

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THE POTENTIAL OF PHOTOCHEMICAL INTERNALIZATION (PCI)

PHOTOCHEMICAL INTERNALIZATION (PCI)

It is well documented that a number of photosensitizers, including di- and tetrasulfonated aluminum phthalocyanine (AlPcSn) and sulfonated tetraphenylporphines (TPPSn) (Figure 17.2), are partly or mainly located in endosomes and lysosomes of cells in culture [10]. The preferential localization of sulfonated tetraphenylporphines and AlPcs in endocytic vesicles is as described in Figure 17.3A, that is, amphiphilic disulfonated compounds are mainly located in the vesicular membranes, while the tetrasulfonated compounds are mainly located in the lumen of the endocytic vesicles [11–13]. The preferential localization of the amphiphilic photosensitizers in the membranes of the endocytic vesicles and the short range of action of singlet oxygen lead to damage mainly to the membranes of these vesicles upon exposure to light, while the contents in the lumen is less affected [11]. These properties of photosensitizing dyes are used in PCI to release endocytosed molecules in a functionally intact form from endosomes and lysosomes as described in Figure 17.3B. Although the exact structure of the damaged vesicles has not been revealed, results with dextran and viral particles (see below) indicate that relatively large particles can escape the vesicles after PCI. PCI of therapeutic macromolecules has been shown to increase the necrotic depth of the treatment and to increase the fraction of complete remissions in animal tumor models. In addition, it has recently been shown that the tumor border is less sensitive to PDT, while PCI has been shown to overcome this limitation of PDT [14, 15]. 17.3.1 Photochemical Delivery of Macromolecules Macromolecules for therapeutic purposes may be divided into three main groups: proteins and peptides, nucleic acids, and synthetic polymers for drug delivery and protection. The present status on the use of PCI for activation of various macromolecular structures is reviewed below. In addition, PCI has

AIPcS2a

TPPS2a SO3−

N N

N

N

N

N

SO3−

N N

AI

N N

N

N SO3

−

SO3−

Figure 17.2. Structure of photosensitizers used in PCI.

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Endosomes/lysosomes A N

~0.5-1 μm

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SO3– SO3– N

~10 nm AlPcS2a

SO3– SO3–

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AlPcS4 SO3–

SO3–

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cytosol

•Amino acids •Unsat. fatty acids •Cholesterol 1O

N N N N NAlN N N –

SO3 Matrix of endocytic vesicles

SO3–

2

lifetime: 0.01-0.04 us diffusion length:10-20 nm

Macromolecules

Figure 17.3. Proposed localization and photochemical effect of sulfonated photosensitizers on endocytic vesicles. (A) A schematic drawing of an endocytic vesicle with approximate dimensions indicated and schematic drawings of the main location of a tetrasulfonated and an amphiphilic disulfonated photosensitizer (not to scale). (B) Schematic illustration of the photochemical effects leading to release of macromolecules from the endocytic vesicles into the cytosol.

been shown to enhance the therapeutic efficacy of some chemotherapeutic agents, including bleomycin [16], which will be utilized in a recently approved clinical Phase I study. 17.3.1.1 PCI for Gene Delivery Gene therapy is a novel therapeutic modality receiving great attention and being generally recognized as having the potential to constitute treatment for a wide range of diseases [17]. However, although there are some encouraging clinical trials, gene therapy has largely given quite disappointing results [18]. An important reason for this is that methods for efficient and specific delivery of therapeutic genes in vivo are lacking. With most gene delivery systems the therapeutic gene is taken into the cell by endocytosis, and for many of these systems, especially nonvirusbased, the lack of efficient mechanisms for translocating the gene out of the endocytic vesicles constitutes a major hindrance for realization of the therapeutic potential of the gene.

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In gene therapy, a tool, named a vector, is usually needed to assist the DNA into the target cells [19]. These tools may be divided into three groups: physical (e.g., electroporation, gene gun), chemical (e.g., polycations like polyethylene imine and polylysine and cationic lipids), and biological vectors (e.g., adenovirus, adeno-associated virus, and retrovirus). The chemical and biological vectors will also protect the DNA from degradation. Both chemical and biological vectors require, with few exceptions, that the DNA–carrier complexes are endocytosed. For efficient gene delivery the therapeutic genes carried by such vectors have to escape from endocytic vesicles so that the genes can be further translocated to the nucleus. Since endosomal escape is often an inefficient process, release of the transgene from endosomes represents one of the most important barriers for gene transfer. Photochemical internalization has been studied as a gene delivery technology (reviewed in Ref. 20) both with several nonviral [21–23] and with adenoviral vectors [24] as well as adeno-associated virus [25], mainly by using reporter genes such as genes encoding EGFP (enhanced green fluorescent protein) or ß-galactosidase. However, PCI-mediated gene delivery has also been shown to induce the delivery of therapeutic genes, such as the genes encoding HSV-tk (herpes simplex virus thymidine kinase) [26], p53 [27], PTEN [28], TRAIL [29], and IL-12 (interleukin-12) (unpublished results). The influence of PCI on transgene expression is illustrated in Figure 17.4. PCI has been shown to increase polycation transfection in some cases 100-fold, cationic lipid-based transfection fivefold, and adenoviral transduction 30-fold [21, 23, 24]. Adenovirus vectors are known to be taken into cells by endocytosis and to be released from endosomes by a regulated process. This endosomal release is usually regarded as a very efficient process [30]. It is therefore somewhat surprising that PCI is able to increase the number of adenoviral transduced cells by up to 30-fold. However, the adenovirus activated by means of PCI seems to follow the same cellular pathways as for conventional adenovirus infection, while PCI increases the number of nucleus-located viral DNA molDNA

+

DNA complexing agent

endocytosis

nucleus

protein

mRNA

Figure 17.4. Schematic illustration of how PCI may improve nonviral vector-based gene transfection.

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317

ecules as measured by real-time PCR and fluorescence in situ hybridization (FISH) [31]. The results so far indicate that the main cause of the photochemical activation of adenovirus transgene expression is related to enhanced release of the viral particles from the endocytic vesicles into the cytosol. 17.3.1.2 Photochemical Delivery of Proteins Ribosome-inactivating protein (RIP) toxins have been evaluated extensively, especially as part of immunotoxins, for use in cancer therapy as well as in graft versus host disease, bone marrow purging, autoimmune diseases, and HIV infection [32]. These protein toxins kill cells by inhibiting protein synthesis. Type I RIP toxins are internalized by fluid-phase endocytosis and transported to lysosomes where these toxins are degraded, while type II RIP toxins can be transported retrogradely from endosomes through the Golgi apparatus and endoplasmic reticulum to cytosol, where these toxins exert their cytotoxic action. PCI has been evaluated by introducing three different type I RIP toxins—gelonin, agrostin, and saporin—into more than 20 different cell lines using TPPS2a or AlPcS2a in combination with light [33, 34]. In all cases a substantial and synergistic reduction in the rate of protein synthesis has been observed when PCI is combined with one of the RIP toxins. These results have also been confirmed in vivo [34–36]. 17.3.1.3 PCI of Dendrimer-Based Drug Delivery Dendrimers are synthetic, highly branched, uniform monodisperse macromolecules of nanometer size with great potential as drug delivery vehicles [37]. They may be used for imaging and drug delivery and may include a targeting moiety. In cancer treatment, due to their size, dendrimers may also benefit from the passive drug delivery through its characteristic leaky vasculature and reduced lymphatic drainage described as the enhanced permeation and retention (EPR) effect. PCI has recently been shown to enhance the therapeutic effect or potential of dendrimers linked to doxorubicin [38, 39], camptothecin [40], saporin [41], and DNA [42–45]. Photosensitizers may also be incorporated into the dendrimers apparently without causing any limiting photooxidation of the drug to be delivered. Interestingly, photosensitizers linked to dendrimers seem to be able to induce the photochemical drug delivery at light doses not causing any phototoxicity [43]. Photochemical treatment as in PDT usually causes skin photosensitivity as its main side effect and dendrimer-delivered photosensitizer may reduce or eliminate this drawback. PCI may also be utilized for treatment of diseases where photocytotoxicity is not acceptable, such as in gene therapy for genetic diseases. Dendrimer-linked photosensitizers for PCIbased drug delivery should also be considered for such purposes. 17.3.2 Targeting and Specificity The photosensitizers accumulate preferentially in neoplastic lesions, which makes these compounds attractive for cancer therapy. One to three days after systemic administration, the concentration of the photosensitizers is usually

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Macromolecule therapy e.g.

Proteins PCI Peptides mRNA DNA Antisense, ribozymes, PNA, siRNA

• Targeting ligands • Specific promoters • Tissue specific replication

PDT • Photosensitizer • Light

• Site-specific light delivery • Preferential retention of PS in cancerous tissues • PS targeting

Improved therapeutic effect and specificity

Figure 17.5. Illustration of how PCI can improve specificity and efficacy of the macromolecular and photodynamic therapies.

two to threefold higher in the neoplastic than in the normal surrounding tissues. Additionally, a cytotoxic reaction will be induced only in the lightexposed areas. PDT is therefore a relatively site-specific treatment modality. A further improved treatment specificity may, however, be established by cell-specific targeting of the photosensitizer and/or the therapeutic macromolecule (Figure 17.5). Several targeting methods are under development, such as the use of antibodies and derivatives thereof, receptor ligands, peptides, tissue-specific promoters, and replication. Some targeting principles have been evaluated in combinations with PCI, such as immunotoxins, transferrin- and epidermal growth factor (EGF)-conjugated vectors, and RGD sequences for integrin targeting [46–51]. These results indicate that PCI can be used in combination with other targeting principles to further improve therapeutic specificity.

17.4

CONCLUSION

PCI is highly efficient in improving internalization of macromolecules. Substantial site direction of the treatment can be obtained by the need for light activation and, in some diseases, such as in solid cancers, the preferential retention of the photosensitizer in the diseased areas. This dual selectivity will reduce damage of the normal surrounding tissue. Most technologies developed so far are useful for internalization of only one group of compounds, while PCI has been shown to act efficiently in internalization of a large variety

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of macromolecules, including nanoparticles, and may be useful for internalization of different macromolecules simultaneously. The method can be combined with most other means for generating site and tissue specificity.

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30. Greber, U. F., Willetts, M., Webster, P., and Helenius, A. Stepwise dismantling of adenovirus 2 during entry into cells. Cell 75: 477–486 (1993). 31. Engesæter, B. O., Tveito, S., Bonsted, A., Engebråten, O., Berg, K., and Mælandsmo, G. M. Photochemical treatment with endosomally localized photosensitizers enhances the number of adenoviruses in the nucleus. J. Gene Med. 8: 707–718 (2006). 32. Thrush, G. R., Lark, L. R., Clinchy, B. C., and Vitetta, E. S. Immunotoxins: an update. Annu. Rev. Immunol. 14: 49–71 (1996). 33. Berg, K., Selbo, P. K., Prasmickaite, L., Tjelle, T., Sandvig, K., Moan, J., Gaudernack, G., Fodstad, O., Kj¢lsrud, S., Anholt, H., Rodal, G. H., Rodal, S. K., and Høgset, A. Photochemical internalization. A novel technology for delivery of macromolecules into cytosol. Cancer Res. 59: 1180–1183 (1999). 34. Selbo, P. K., Høgset, A., Prasmickaite, L., and Berg, K. Photochemical internalisation: a novel drug delivery system. Tumor Biol. 23: 103–112 (2002). 35. Selbo, P. K., Sivam, G., Fodstad, Ø., Sandvig, K., and Berg, K. In vivo documentation of photochemical internalization, a novel approach for site specific cancer therapy. Int. J. Cancer 92: 761–766 (2001). 36. Dietze, A., Peng, Q., Selbo, P. K., Muller, C., Bown, S., and Berg, K. Enhanced photodynamic destruction of a transplantable fibrosarcoma using photochemical internalisation (PCI) of gelonin. Br. J. Cancer 92: 2004–2009 (2005). 37. Patri, A. K., Majoros, I. J., and Baker, J. R. Dendritic polymer macromolecular carriers for drug delivery. Curr. Opin. Chem. Biol. 6: 466–471 (2002). 38. Lai, P. S., Lou, P. J., Peng, C. L., Pai, C. L., Yen, W. N., Huang, M. Y., Young, T. H., and Shieh, M. J. Doxorubicin delivery by polyamidoamine dendrimer conjugation and photochemical internalization for cancer therapy. J. Control. Release 122: 39–46 (2007). 39. Lou, P. J., Lai, P. S., Shieh, M. J., Macrobert, A. J., Berg, K., and Bown, S. G. Reversal of doxorubicin resistance in breast cancer cells by photochemical internalization. Int. J. Cancer 119: 2692–2698 (2006). 40. Cabral, H., Nakanishi, M., Kumagai, M., Jang, W. D., Nishiyama, N., and Kataoka, K. A photo-activated targeting chemotherapy using glutathione sensitive camptothecin-loaded polymeric micelles. Pharm. Res. 26: 82–92 (2009). 41. Lai, P. S., Pai, C. L., Peng, C. L., Shieh, M. J., Berg, K., and Lou, P. J. Enhanced cytotoxicity of saporin by polyamidoamine dendrimer conjugation and photochemical internalization. J. Biomed. Mater. Res. A 87: 147–155 (2008). 42. Shieh, M. J., Peng, C. L., Lou, P. J., Chiu, C. H., Tsai, T. Y., Hsu, C. Y., Yeh, C. Y., and Lai, P. S. Non-toxic phototriggered gene transfection by PAMAM-porphyrin conjugates. J. Control. Release 129: 200–206 (2008). 43. Nishiyama, N., Iriyama, A., Jang, W. D., Miyata, K., Itaka, K., Inoue, Y., Takahashi, H., Yanagi, Y., Tamaki, Y., Koyama, H., and Kataoka, K. Light-induced gene transfer from packaged DNA enveloped in a dendrimeric photosensitizer. Nat. Mater. 4(2): 934–941 (2005). 44. Nishiyama, N., Arnida, Jang, W. D., Date, K., Miyata, K., and Kataoka, K. Photochemical enhancement of transgene expression by polymeric micelles incorporating plasmid DNA and dendrimer-based photosensitizer. J. Drug Target. 14: 413–424 (2006).

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45. Nishiyama, N., Morimoto, Y., Jang, W. D., and Kataoka, K. Design and development of dendrimer photosensitizer-incorporated polymeric micelles for enhanced photodynamic therapy. Adv. Drug Deliv. Rev. 61: 327–338 (2009). 46. Selbo, P. K., Sivam, G., Fodstad, Ø., Sandvig, K., and Berg, K. Photochemical internalisation increases the cytotoxic effect of the immunotoxin MOC31-gelonin. Int. J. Cancer 87: 853–859 (2000). 47. Prasmickaite, L., Høgset, A., Tjelle, T. E., Sandvig, K., and Berg, K. The role of endosomes in gene transfection mediated by photochemical internalisation. J. Gene Med. 2: 477–488 (2000). 48. Weyergang, A., Selbo, P. K., and Berg, K. Photochemically stimulated drug delivery increases the cytotoxicity and specificity of EGF-saporin. J. Control. Release 111: 165–173 (2006). 49. Bonsted, A., Engesæter, B. Ø., Høgset, A., Mælandsmo, G. M., Prasmickaite, L., and Berg, K. Photochemically enhanced transduction of polymer-complexed adenovirus targeted to the epidermal growth factor receptor. J. Gene Med. 8: 286–297 (2006). 50. Kloeckner, J., Boeckle, S., Persson, D., Roedl, W., Ogris, M., Berg, K., and Wagner, E. DNA polyplexes based on the degradable oligoethyleneimine-derivatives: combination with EGF receptor targeting and endosomal release functions. J. Control. Release 116: 115–122 (2006). 51. Yip, W. L., Weyergang, A., Berg, K., Tønnesen, H. H., and Selbo, P. K. Targeted delivery and enhanced cytotoxicity of cetuximab-saporin by photochemical internalization in epidermal growth factor-positive cancer cells. Mol. Pharm. 4: 241–251 (2007).

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CHAPTER 18

Peptide-Based Nanocarriers for Intracellular Delivery of Biologically Active Proteins SEONG LOONG LO Institute of Bioengineering and Nanotechnology, Singapore

SHU WANG Institute of Bioengineering and Nanotechnology, and Department of Biological Sciences, National University of Singapore, Singapore

18.1

INTRODUCTION

Over the past few decades, synthetic peptides have emerged as one of the highly attractive biomaterials and have shown significant potential for biomedical applications. Peptide-based biomaterials were originally developed based on the biochemical understanding that the active sites of protein molecules, such as enzymes, receptor ligands, and antibodies, usually involve only 5 to 20 amino acid residues [1]. With rapid advances in structural biology and high-throughput genomics and proteomics, the identification of peptide motifs associated with biological functions has been drastically accelerated [2]. Thus peptide-based biomaterials offer a highly attractive feature of incorporating various natural or synthetic sequences with biological activities, for example, cell targeting domain or organelle-specific targeting domain. Peptides are relatively easy to synthesize on a large scale and can be characterized with wellestablished chemistry and instrumental operation. Peptides are also generally less toxic and have low immunogenicity compared to high molecular weight polymers [3], and undergo degradation in the body to naturally occurring compounds. Different nanometric structures can arise from peptide–cargo interactions of electrostatic, hydrophobic, or aromatic nature. A good example Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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is nanoparticles formed by peptides and nucleic acids. With increasing evidence showing that peptides are promising nanocarriers for nucleus delivery of plasmid DNA, the concept of using peptides as a vector has been extended to intracellular delivery of proteins, especially into the cytoplasm. Traditionally, delivery of genes encoding proteins through gene transfection or viral transduction is used as an indirect method to introduce protein into cells. The duration and level of protein expression resulting from such indirect ways are usually difficult to control in a precise manner due to the complex cellular machinery for gene expression and possible random chromosomal integration of delivered genes. Therefore direct intracellular delivery of proteins seems to be an attractive alternative. Translocation of proteins across the cell membrane is usually limited by the permeability and selectivity of the membrane for macromolecules. Different strategies that have been employed to improve internalization of proteins include physical methods, modification of proteins, and noncovalent encapsulation with lipids, polymers, or peptides. Physical methods such as electroporation and microinjection are the most direct methods to introduce foreign substances into cells. Although these methods were first used to introduce foreign DNA into cells [4, 5], several studies demonstrated the potential of these methods for protein delivery into cultured cells in vitro [6–14]. The use of physical methods is usually limited due to the high rate of cell mortality caused by high-voltage pulses during electroporation and unavailability of specialized equipment for microinjection. A widely used approach for protein delivery is to modify the cargo proteins using a peptide sequence with membrane penetration ability, termed “cell-penetrating peptide” or “protein transduction domain” [15]. The modification involves the expression and purification of a recombinant fusion protein from an expression vector containing DNA sequence of the cellpenetrating peptide and the cargo protein. One of the cell-penetrating peptides, Tat derived from human immunodeficiency virus [16], has been successful in delivering different therapeutic fusion proteins in vitro and in vivo [17]. Generation of induced pluripotent stem cells using oligoarginine fused transcription factors instead of virus has also been reported recently [18]. However, modification with a cell-penetrating sequence may result in loss of protein biological functions due to possible interference to conformational folding and urea denaturation that is required to increase the accessibility of the internally buried cell-penetrating sequence. Hence noncovalent encapsulation of proteins with lipids and polymers would be a feasible alternative for intracellular protein delivery. Lipids and polymers are known to be effective transfection reagents for plasmid DNA. By optimizing the lipid formulation and polymer composition, these reagents can also be applied for protein delivery [19–25]. Some lipid-based products are commercially available for protein delivery, for example, BioPORTER® (Genlatis), PULSin™ (Polyplus Transfection), and Pro-Ject™ (Pierce). Despite the potential usage for in vivo delivery, allergictype immune reactions triggered by lipid complexes [26, 27] remain as issues to be addressed. In this chapter, we will focus on published studies using

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peptide biomaterials as nanocarriers to encapsulate and deliver biologically active protein cargos into the cytoplasm.

18.2

AMPHIPATHIC PEPTIDE-BASED NANOCARRIERS

Several of the strategies developed for nucleic acids delivery provide good starting points for designing new peptide-based nanocarriers for proteins/ peptides delivery. Depending on their design, peptide-based nanocarriers that have been tested for protein delivery can be categorized into two groups— amphipathic peptides and cell-penetrating peptides (Table 18.1). An amphipathic peptide is a peptide that possesses both hydrophobic (nonpolar) and hydrophilic (polar) properties. The amphipathicity characteristic usually originates from a peptide sequence that simply links a hydrophobic domain and a hydrophilic domain together. The most well-characterized amphipathic peptide-based carrier is the Pep-1 peptide [28], consisting of three domains: (1) a hydrophobic tryptophan-rich domain that promotes hydrophobic interaction with cargoes (KETWWETWWTEW); (2) a spacer with proline residue that improves peptide flexibility (SQP); and (3) a hydrophilic lysinerich domain derived from nuclear localization sequence of simian virus 40 (SV40) large T-antigen (KKKRKV). Both N and C termini were modified by an acetyl group and a cysteamide group, respectively, for improved peptide

TABLE 18.1

Peptides Involved in Intracellular Delivery of Proteins/Peptidesa

Amphipathic peptides Pep-1 Wr-T Wr-Tm1 Wr-Tm2 WR-T Rath P10C 10K-VTW 16K-VTW Cell-penetrating peptides M918 Penetratin Transportan 10 YTA2 pVEC Tat SR9 HGH6

Ac-KETWWETWWTEW-SQP-KKKRKV-Cys KETWWETWWTEWWTEWSQ-GPG-rrrrrrrrr KETWWETWWTEWSQ-GPG-rrrrrrrrr KETWWETWWTEWWTEWSQ-GPG-PKKKRKV KETWWETWWTEWWTEWSQ-GPG-RRRRRRRRR TPWWRLWTKW-HHRRDLPRKPE Ac-VGAlAvVvWlWlWlW(βA)-GSGP-KKKRKVC-Am VTWTPQAWFQWV-GGGS-K10 K16-SIPVKFNKP-VTWTPQAWFQWV MVTLFRRLRIRRACGPPRVRV-Am RQIKIWFQNRRMKWKK-Am AGYLLGKINKALAALAKIL-Am Ac-YTAIAWVKAFIRKLRK-Am LLIILRRRIRKQAHAHSK-Am GRKKRRQRRRPPQ-Am RRRRRRRRR CG(RHGH)5RGC

Ac = acetyl group; Cys = cysteamide group; lowercase letter = d-amino acid; (ßA) = ß-alanine; and Am = amide group. a

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stability and ability to cross the cell membrane. FITC-labeled Pep-1 peptide was found to localize rapidly to the nucleus of different mammalian cell lines at 37 °C and 4 °C, suggesting that the cellular uptake pathway of the Pep-1 peptide is independent of the endocytotic pathway that requires energy [28]. This was further confirmed by unaffected cellular uptake in the presence of inhibitors of endocytotic pathway (nystatin, cytochalasin D, and amiloride) [29]. The amphipathic Pep-1 peptide was capable of forming stable complexes through noncovalent hydrophobic interaction with peptide or protein cargoes in solution, as confirmed by size exclusion chromatography [28]. The resulting Pep-1/cargo complexes could be efficiently internalized intracellularly after incubation with cells for 30 minutes, either in serum-free or serum-containing medium. More importantly, even though the Pep-1 peptide rapidly localized to the nucleus, cargoes like the 119-kDa subunit of β-galactosidase (β-gal), green fluorescent protein (GFP), cell cycle inhibitor protein p27Kip1, and monoclonal antibody anti-β-actin retained their respective intracellular localizations and biological functions after a “de-caging” process that dissociates the interaction between the Pep-1 peptide and cargo [28, 29]. To achieve maximum transduction efficiency, molar ratios of the Pep-1/cargo complexes have been extensively explored and a molar ratio between 15 : 1 and 20 : 1 was found to be optimal. Higher molar ratio resulted in aggregation and precipitation of the Pep-1/cargo complex that significantly reduced delivery efficiency [30]. Based on various characterization studies using physical and spectroscopic approaches, a cellular uptake mechanism underlying Pep-1-mediated protein delivery was proposed [31–33]. The hydrophobic domains of several Pep-1 molecules first interact with cargos through noncovalent hydrophobic interaction. The Pep-1/cargo complexes are formed with “a peptide shell” surrounding cargos. During incubation with the cells, the positively charged nuclear localization sequence of the Pep-1 peptide will interact with the cell membrane through an electrostatic interaction. After the Pep-1/cargo complexes are anchored on the cell membrane, the α-helical folded hydrophobic domains of the peptides form a transmembrane pore to allow translocation of the complexes into the cytoplasm. Following internalization, the “de-caging” process will allow the cargoes to be released from the complexes. However, some studies did not detect pore formation [34–38] but observed perturbation of model lipid membrane in the presence of the Pep-1 peptide [34, 39]. Despite the controversy surrounding the mechanism, the Pep-1 peptide efficiently delivered various peptide or protein cargoes into immortalized mammalian cell lines [29] and primary cells such as hepatocytes [40, 41], pancreatic cells [42, 43], neurons [44], neural retina cells [45], mouse Müller glia cells [46], human stem cells [47], and macrophages [48]. Moreover, the Pep-1 peptide mediated in vivo delivery of functional proteins like the apoptosis related caspase-3 and the signaling molecule protein kinase A in the lung of mice to produce alveolar wall apoptosis and repair a defective step in the signaling pathway, respectively [49, 50]. Interestingly, when the Pep-1 peptide sequence

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was included in proteins like Cu/Zn-superoxide dismutase and heat shock protein 27, the resulting fusion proteins were able to penetrate neuronal cells and provided protection in vitro and in vivo against reactive oxygen species related damage [51–53]. The Pep-1 peptide is currently manufactured and commercialized under the name of Chariot™ by Active Motif Inc. Hence the Pep-1 peptide represents a potent strategy with great potential in basic research and possibly even therapeutic applications. Inspired by the Pep-1 peptide, a derivative, namely, Wr-T, was designed to deliver antitumor peptides to suppress growth of leukemia/lymphomas, glioblastoma cell lines, and tumor xenograft [54, 55]. Similar to the structure of the Pep-1 peptide, the Wr-T peptide consists of a hydrophobic domain, a spacer, and a hydrophilic domain. The hydrophobic domain of the Wr-T peptide has an expanded tryptophan-rich motif that was predicted to be more effective in interacting with cargo. In addition, the original SV40 nuclear localization sequence in the Pep-1 peptide was replaced by oligo-d-arginine in the Wr-T peptide for enhanced cell permeability [56]. Using β-galactosidase as a model protein for delivery, X-gal staining revealed that the Wr-T peptide was significantly more efficient than the Pep-1 peptide and the mutants of Wr-T peptide with either truncated hydrophobic domain or SV40 nuclear localization sequence (Wr-Tm1 and Wr-Tm2) [55]. Cellular uptake of FITClabeled antitumor peptide mediated by the Wr-T peptide was also 25-fold and twofold higher than that mediated by the Pep-1 peptide and the WR-T peptide (a counterpart with oligo-l-arginine), respectively [54]. Comparing the efficiency of inhibition by antitumor peptide p16-MIS delivered by Wr-T peptide or the Pep-1 peptide, growth of lymphoma cells treated with the Wr-T/p16MIS complex was suppressed to a greater extent (73%) than that resulting from the Pep-1 peptide (23%) [54]. The intravenous delivery of antitumor peptides (p16-MIS and p21-S154A) mediated by Wr-T was also confirmed by prolonged overall survival of glioblastoma bearing nude mice compared to the nontreated group [55]. It is noteworthy that the optimal molar ratio of the Wr-T/peptide-cargo complex was 2.5 : 1, which was much lower than that offered by the Pep-1 peptide. Hence the observed enhanced cellular uptake and delivery of peptide cargo could be attributed to the improved packaging and cell penetration associated with expanded hydrophobic domain and oligod-arginine in the Wr-T peptide sequence. Amphipathic peptides can also be designed based on natural proteins. A novel peptide, Rath, derived from VP5 capsid protein of infectious bursal disease virus, was identified as an amphipathic cell-penetrating peptide with dominant β-sheet conformation [57]. FITC-labeled Rath localized mainly in cell nucleus at both 37 °C and 4 °C within 30 minutes and successfully delivered FITC-labeled IgG antibody into monkey kidney epithelial cells (Vero) and chicken embryonic fibroblast (CEF) primary cells [57]. However, extra washing steps with trypsin were not performed before fixation to remove the extracellularly bound complex that might cause artifacts commonly associated with translocation of cell-penetrating peptide [58]. In addition, the optimal molar

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ratio of the Rath/cargo complex was between 285 : 1 and 570 : 1, which is about 15–30 times more than that of the Pep-1/cargo complex. Although the Rath peptide was not cytotoxic at high concentration up to 75 μM, it is not cost effective for cargo delivery and the size of peptide/cargo complex might actually be unfavorable for internalization. Another example is a cysteinecontaining peptide, P10C, that is derived from hydrophobic membrane active sequence of an antibiotic gramicidin A [59]. Sequence of the P10C peptide adopted similar design as the Pep-1 peptide, having a tryptophan-rich hydrophobic domain linked to SV40 nuclear localization sequence with a spacer. The P10C peptide was shown to be five fold more effective in delivering β-gal than the analog without cysteine residue, possibly due to oxidative dimerization through disulfide bond formation that led to more efficient cargo packaging and membrane interaction. It was also demonstrated that the delivery efficiency of the P10C peptide was not affected when the positions of cysteine residue, hydrophobic domain, and hydrophilic domain were shuffled between N and C termini. This suggested that the amphipathicity rather than the relative position was the determining factor of amphipathic peptide-based delivery. The P10C/β-gal complex formation at a molar ratio of 50 : 1 was confirmed with polyacrylamide gel electrophoresis under nondenaturing conditions. The β-gal delivery efficiency of the P10C peptide was also compared with that of the Pep-1 peptide in HeLa cells. In contrast to the previous finding, the β-gal activity mediated by the Pep-1/β-gal complex was negligible at the optimal molar ratio of 20 : 1 [28]. Even when the molar ratio of Pep-1/β-gal complex was increased to 400 : 1, the β-gal activity was about twofold lower than that offered by P10C/β-gal complex formed at a molar ratio of 50 : 1. The difference in delivery efficiency of the Pep-1 peptide might arise from the β-galactosidase used in the experiments. While the previous study had used a 119-kDa subunit of β-gal as the model protein, a homotetramer of β-gal was used in the current study. This discrepancy suggested that the molecular weight of cargo protein might affect the packaging ability and delivery activity of the Pep-1 peptide. Besides natural proteins, peptides identified from a phage display library are another attractive source of amphipathic peptides. In a particular study, a glioblastoma targeting peptide sequence VTW, selected from a 12-mer phage display library, displayed efficient and specific binding to several human glioma cell lines but relatively little binding to other mammalian cell lines [60]. BLAST analysis revealed that the VTW sequence was similar to a 7-residue human interleukin-11 receptor α-chain (IL-11RA) that binds to gp130 found to be overexpressed on glioma cell surface, as assessed by DNA microarray analysis. To test whether the VTW peptide can be employed for targeted gene delivery, the 10K-VTW peptide was synthesized by adding oligolysine to the C terminus of the VTW peptide for DNA binding. However, the 10K-VTW peptide was not effective in gene delivery, probably due to the inability to condense DNA efficiently. Coincidentally, similar to the hydrophobic domain of the Pep-1 peptide, the VTW sequence is rich in tryptophan residues. Hence

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the 10K-VTW peptide is an amphipathic peptide that might be useful for intracellular delivery of proteins. When the 10K-VTW peptide was complexed with a 119-kDa subunit of β-gal at a weight/weight ratio of 1 : 1 (corresponding to a molar ratio of 40 : 1), 60% of the complex-treated human malignant glioma cells (U87MG) were positively stained. When another amphipathic peptide, 16K-VTW, was designed based on a previous publication [61] and used for β-gal delivery at a weight/weight ratio of 1 : 1 (corresponding to a molar ratio of 25 : 1), up to 90% of U87MG cells were positively stained. A dose-dependent increase in staining intensity in U87MG cells was also observed when the weight/weight ratio was doubled. Both 10K-VTW and 16K-VTW did not show obvious protein delivery activity in immortalized human astroglial cells (SVGp12). To verify that the staining was not due to fixation artifact associated with extracellularly bound 16K-VTW/β-gal complex, the β-gal activity was compared between trypsinized and PBS-treated U87MG cells after delivery. No significant difference in β-gal enzyme activity was detected between the two groups, suggesting that the successful delivery of β-gal was mediated by the 16K-VTW peptide. To our best knowledge, this is the first demonstration of using a peptide-based nanocarrier for intracellular delivery of protein in a tumor cell-specific manner. With further optimization, like replacement of oligolysine with a cell-penetrating sequence to enhance cellular uptake, the VTW-based amphipathic peptide could be a promising nanocarrier for targeted therapy.

18.3

CELL-PENETRATING PEPTIDE-BASED NANOCARRIERS

Cell-penetrating peptides (CPP) are usually rich in arginine (R) and lysine (K) that are highly positively charged. Some CPPs, like penetratin [62] and Tat [16], are directly derived from natural protein while others are a designed sequence, such as oligoarginine [63] and transportan [64]. Many different cargoes have been successfully delivered into cells by CPPs through covalent conjugation or noncovalent formation of nanocarrier, demonstrating that a CPP is a highly versatile delivery agent [65]. Indeed, several studies have successfully demonstrated the delivery of proteins using CPP-based nanocarriers, although the exact mechanism underlying complex formation between a CPP and protein cargo remains unclear. One of the natural protein-derived CPPs is the M918 peptide that has a sequence of amino acids 1–22 of tumor suppressor protein p14ARF (positions 3–8 inverted) [66]. Similar to the other two CPPs, penetratin and transportan 10 (TP10), the internalization of the M918 peptide involved endocytosis, as confirmed by vesicular distribution and decreased internalization in the presence of inhibitors (4 °C incubation, sucrose or chloroquine). However, the cellular uptake of the M918 peptide was significantly higher than penetratin and TP10 in human cervical cancer cells (HeLa) and Chinese hamster ovary

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(CHO) cells. All these three CPPs also promote delivery of FITC-labeled avidin and streptavidin at a molar ratio of 25 : 1.Compared to penetratin and TP10, the M918 peptide provided up to 2.5-fold increase in cellular uptake of FITC-labeled cargoes. A synthetic CPP, YTA2, was initially designed to conjugate cytostatic agent methotrexate for drug delivery [67]. It was later discovered that the YTA2 peptide also has the capability to deliver proteins into cells without conjugation with the cargo [68]. The cellular uptake and toxicity of the YTA2 peptide was compared with other CPPs, including pVEC, penetratin, TP10, and Tat, in a human breast cancer cell line (MDA-MB-231) and a human melanoma cell line (Bowes). The amount of internalized the pVEC peptide was the highest, followed by YTA2, TP10, penetratin, and Tat. However, the YTA2 peptide resulted in significantly higher lactate dehydrogenase leakage than the other CPPs and the Pep-1 peptide, indicating that the YTA2 peptide disrupted the cell membrane and could be cytotoxic to a certain extent. Nonetheless, the YTA2 peptide was as efficient as the Pep-1 peptide in delivering the 119kDa subunit of β-gal into Bowes cells. When the cargo was changed to 66-kDa TRITC-labeled streptavidin, the YTA2 peptide appeared to be half as effective as the Pep-1 peptide, suggesting that the protein delivery activity of the YTA2 peptide might be related to protein property. Another synthetic CPP, nona-arginine (SR9), has also been demonstrated to facilitate in vitro delivery of different proteins such as red fluorescent protein (RFP), green fluorescent protein (GFP), and β-gal [69]. In vivo transdermal delivery of GFP with the SR9 peptide to the backs of mice exhibited strong green fluorescence rapidly at different skin layers, indicating SR9 peptide-mediated penetration of protein cargo. To facilitate reversible association with protein cargoes, cysteine residues can be added to N and C termini of a peptide to form interpeptide disulfide bonds that will be cleaved by reductive gluthatione species in the cellular environment. Based on this concept, a cysteine-flanked CPP, HGH6, was tested for the capacity to deliver proteins [70]. The HGH6 peptide formed a complex with 465-kDa β-gal tetramer reversibly, between a peptide/protein molar ratio of 50 : 1 and 100 : 1. X-gal staining after treatment with trypsin confirmed that the HGH6/β-gal complex (molar ratio of 50 : 1) was internalized by various mammalian cell lines. Reduced cellular uptake was observed with higher molar ratios, probably due to excessive interpeptide disulfide bond formation that affected the dissociation process. Protein delivery activity of the HGH6 peptide was not affected by 4 °C incubation and ATP depletion, suggesting an energy-free internalization pathway. Similar to the Pep-1 peptide, the HGH6 peptide was capable of delivering proteins with different molecular weights, including caspase-3 (12- and 17-kDa heterodimer), placental alkaline phosphatase (116-kDa dimer), and immunoglobulin (150-kDa IgG). In vivo studies also showed that apoptosis of tumor cells was induced after multiple intratumor administrations of HGH6/caspase-3 complexes to subcutaneous tumor formed by HeLa cells in nude mice.

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OUTLOOK AND FUTURE PERSPECTIVES

18.4

331

OUTLOOK AND FUTURE PERSPECTIVES

The completion of the Human Genome Project has advanced and accelerated the discovery and development of protein-based therapeutics. The identification of genes encoding for various proteins can allow the production of therapeutic proteins by recombinant DNA technology. Lack of an efficient delivery system appears to be the bottleneck for making protein-based therapeutics useful for clinical applications. As the half-life of proteins might be short in blood circulation, noncovalent encapsulation with nanocarriers can potentially protect proteins from rapid degradation. Functioning as delivery systems, peptides appear to be superior to many other types of materials in several interesting aspects. During peptide design, different functional domains can be included to overcome extra- and intracellular barriers to bioactive molecule delivery. This offers the flexibility of functioning peptide vectors for tissue or cell type-specific delivery. With their relatively short sequences, peptides can be produced without complicated instrumental operation or chemistry. Peptide backbones can be chemically modified during synthesis to reduce degradation and improve bioavailability. The fact that the molecular structure and purity of peptides can be accurately determined makes the quality control of peptide materials somewhat easy. Attractively, as a biocompatible and biodegradable material, peptides can degrade in the body to naturally occurring molecules and produce relatively low toxicity. Owing to their low molecular weight, peptides are less immunogenic, thus offering the possibility for repeated administration to achieve long-term therapeutic effects when needed. Thus rationally designed peptidebased nanocarriers hold great potential for protein delivery. Indeed, one of the peptides, Pep-1, has been demonstrated to deliver various biologically active proteins in vitro and in vivo without affecting their respective cellular localizations and functions. The challenge in the field of peptide-based protein delivery arises from the unique and variable properties of different types of protein cargoes. Given the wide variations in sequence, structure, chemistry, and function that proteins can have, available delivery systems are unlikely to be universally suitable, necessitating that several factors be taken into account for the design of peptide-based delivery systems. First, the potential use of in silico threedimensional modeling and prediction of peptide–protein interaction should be explored when moving toward rational design of peptide sequence. Second, the interaction between peptide and protein, such as possible strong electrostatic interaction or disulfide bond formation by highly positively charged or cysteine-containing peptide sequence, should be assessed carefully to account for any adverse effects on the biological activities of proteins. Third, the molar ratio to form the peptide/protein complex will depend on the size and properties of the protein. Optimization of this molar ratio is necessary for delivery of different proteins. Fourth, the cellular uptake mechanism of the peptide/ protein complex should be investigated. When the endocytotic pathway is

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involved, the endosomal escape of the complex will be a limiting factor for efficient delivery. To avoid use of endosomolytic agents, it is possible to incorporate histidine residues into the peptide design to favor escape of the complex from endosomes [71–73]. Fifth, immunogenicity of peptides, proteins, and complexes should be investigated carefully when moving toward in vivo application. A cell type-specific targeting motif, like the VTW peptide sequence [60], will reduce the possibility of stimulating the immune system and also lower the dosage to be used for therapeutic purposes. In conclusion, recent progress provides increasing evidence that peptides are promising nanocarriers for protein delivery. Inputs from researchers in different disciplines will undoubtedly aid in a better understanding of protein functions, peptide chemistry, biophysical properties of the peptide/protein complex, pharmacodynamics, and pharmacokinetics, thereby accelerating the development of peptide-based nanocarriers for protein delivery.

ACKNOWLEDGMENTS We would like to thank Ms. Yukti Choudhury for proofreading and discussion.

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46. Wang, M. H., Frishman, L. J., and Otteson, D. C. Intracellular delivery of proteins into mouse Muller glia cells in vitro and in vivo using Pep-1 transfection reagent. J. Neurosci. Methods 177(2): 403–419 (2009). 47. Chan, S. A., et al. Adrenal chromaffin cells exhibit impaired granule trafficking in NCAM knockout mice. J. Neurophysiol. 94(2): 1037–1047 (2005). 48. Garnon, J., et al. Fragile X-related protein FXR1P regulates proinflammatory cytokine tumor necrosis factor expression at the post-transcriptional level. J. Biol. Chem. 280(7): 5750–5763 (2005). 49. Aoshiba, K., Yokohori, N., and Nagai, A. Alveolar wall apoptosis causes lung destruction and emphysematous changes. Am. J. Respir. Cell Mol. Biol. 28(5): 555–562 (2003). 50. Maron, M. B., et al. PKA delivery to the distal lung air spaces increases alveolar liquid clearance after isoproterenol-induced alveolar epithelial PKA desensitization. Am. J. Physiol. Lung Cell Mol. Physiol. 289(2): L349–L354 (2005). 51. Choi, H. S., et al. PEP-1-SOD fusion protein efficiently protects against paraquatinduced dopaminergic neuron damage in a Parkinson disease mouse model. Free Radic. Biol. Med. 41(7): 1058–1068 (2006). 52. An, J. J., et al. Transduced human PEP-1-heat shock protein 27 efficiently protects against brain ischemic insult. FEBS J. 275(6): 1296–1308 (2008). 53. Cho, J. H., et al. Effective delivery of Pep-1-cargo protein into ischemic neurons and long-term neuroprotection of Pep-1-SOD1 against ischemic injury in the gerbil hippocampus. Neurochem. Int. 52(4-5): 659–668 (2008). 54. Kondo, E., et al. Highly efficient delivery of p16 antitumor peptide into aggressive leukemia/lymphoma cells using a novel transporter system. Mol. Cancer Ther. 3(12): 1623–1630 (2004). 55. Kondo, E., et al. Potent synergy of dual antitumor peptides for growth suppression of human glioblastoma cell lines. Mol. Cancer Ther. 7(6): 1461–1471 (2008). 56. Wender, P. A., et al. The design, synthesis, and evaluation of molecules that enable or enhance cellular uptake: peptoid molecular transporters. Proc. Natl. Acad. Sci. U.S.A. 97(24): 13003–13008 (2000). 57. Bais, M. V., et al. Novel Rath peptide for intracellular delivery of protein and nucleic acids. Biochem. Biophys. Res. Commun. 370(1): 27–32 (2008). 58. Richard, J. P., et al. Cell-penetrating peptides. A reevaluation of the mechanism of cellular uptake. J. Biol. Chem. 278(1): 585–590 (2003). 59. Stoilova, T. B., et al. Gramicidin A-based peptide vector for intracellular protein delivery. Biochim. Biophys. Acta 1778(10): 2026–2031 (2008). 60. Wu, C., et al. A peptide-based carrier for intracellular delivery of proteins into malignant glial cells in vitro. J. Control. Release 130(2): 140–145 (2008). 61. Ho, I. A., Lam, P. Y., and Hui, K. M. Identification and characterization of novel human glioma-specific peptides to potentiate tumor-specific gene delivery. Hum. Gene Ther. 15(8): 719–732 (2004). 62. Derossi, D., et al. The third helix of the Antennapedia homeodomain translocates through biological membranes. J. Biol. Chem. 269(14): 10444–10450 (1994). 63. Mitchell, D. J., et al. Polyarginine enters cells more efficiently than other polycationic homopolymers. J. Pept. Res. 56(5): 318–325 (2000).

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CHAPTER 19

Organelle-Specific Pharmaceutical Nanotechnology: Active Cellular Transport of Submicro- and Nanoscale Particles GALYA ORR Chemical and Materials Sciences Division, Pacific Northwest National Laboratory, Richland, Washington

19.1

VIRUSES INSPIRE THE DESIGN OF SYNTHETIC CARRIERS

Living cells have evolved highly efficient transport systems that support the delivery of vesicles and molecules within and between the intracellular and extracellular environment. These systems have been hijacked and manipulated by viruses and other microorganisms to support their cellular and nuclear invasion and trafficking [reviewed in Ref. 1]. Inspired by the success of viruses to efficiently deliver their genes and shuttle their proteins, artificial viruses or synthetic carriers are now being designed for gene and drug delivery [reviewed in Refs. 2–4]. Unlike viruses, synthetic particles can be designed with a larger carrying capacity and cell-specific targeting mechanisms, while being safer for clinical applications [reviewed in Ref. 2]. In this chapter, the transport systems that are exploited by viruses and synthetic particles to support their trafficking will be briefly reviewed, and a special emphasis will be given to recent findings about the retrograde recruitment system, which propels particles along filopodia and microvilli toward the cell body. Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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19.2 MOTOR PROTEINS POWER THE TRANSPORT OF VIRUSES AND SYNTHETIC PARTICLES ALONG CYTOSKELETAL FILAMENTS The transport systems consist of dynamic cytoskeletal filaments and their associated motor proteins. Cargo can be transported along microtubules, powered by dynein or kinesin motors. Dyneins power most minus-end directed transport, such as the transport of vesicles or viruses toward the nucleus or the microtubule organizing center [5–10], while kinesins power most plus-end directed transport, such as the delivery of newly synthesized viruses from the site of assembly to the plasma membrane [11–15]. The active transport of nonviral engineered nanoparticles along microtubules has been observed in neuronal and nonneuronal cells [reviewed in Refs 16 and 17]. The cationic polyethylene imine (PEI) nanocarrier is thought to be one of the most efficient synthetic vectors for DNA delivery. The PEI carrier is able to protect DNA from degradation and facilitate endonsomal escape [18–21]. The active transport of individual PEI/DNA nanocomplexes to the perinuclear region has been quantified in cultured COS-7 cells, showing striking similarities to the transport pattern of viral particles designed for DNA delivery [22]. PEI/DNA complexes have also been shown to undergo a retrograde transport from the axon to the neuronal soma in vivo and in vitro [23, 24]. However, most nonviral carriers are actively transported within endocytic vesicles, which become acidic and degrade their content over time. Synthetic carriers that could escape endosomes early in the process and take advantage of the active transport, while being free of endosomes, would greatly increase the delivery efficiency of the carrier. Inspired by the ability of viruses to directly interact with motor proteins and be transported along microtubules, synthetic nanoparticles that carry dynein-binding peptides at their surface are currently being designed [25–27]. Actin filaments, which are dynamically assembled and disassembled under the control of multiple proteins [reviewed in Ref. 28], provide tracks for transporting cargo, powered by myosin motors [29]. Certain mysoins, such as myosin I or V, carry cargo toward the plus end of the filaments, while other myosins, such as myosin VI, carry cargo toward the minus end [30]. Several studies show that the integrity of the actin network dynamics is critical for the infection process of many viruses [31–33], especially when entering through the endocytic pathway [34–36]. The actin network dynamics beneath the plasma membrane plays important roles in the endocytic process [37–40]. Myosin VI, which is colocalized with endocytic vesicles, is thought to power the transport of the vesicles from the plasma membrane through the cortical actin toward early endosomes [41]. Actin filaments in the nucleus are also involved in active transport of progeny capsids to the nuclear membrane [42], or the export of viral genome to the cytosol [43, 44].

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ACTIN FLOW UNDERLIES THE SURFING OF VIRUSES

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19.3 ACTIVE TRANSPORT WITHIN AND BETWEEN CELLS CAN BE POWERED BY ACTIN POLYMERIZATION Active transport can be generated by the sheer polymerization of actin filaments [reviewed in Ref. 45]. Actin polymerization, which underlies the generation of cytoplasmic extensions at the leading edge of motile cells [reviewed in Ref. 28], has been exploited by certain bacteria and vaccinia virus to propel their way within or between cells [reviewed in Refs. 46 and 47]. By recruiting host actin and cytoskeletal proteins to their surface, these microorganisms activate the assembly of an actin tail, resembling a comet tail. These organisms have evolved distinct proteins that recruit the actin assembly machinery to one of their poles, leading to the continuous assembly of the tail that propels the organism within the cytoplasm and into neighboring cells. By coating polystyrene beads or anionic plasmid carriers with ActA, the Listeria protein responsible for initiating comet-tail formation, it was possible to induce the formation of the tail and the directional propulsion of the particles in an actin reach cytoplasmic extract [48, 49].

19.4 ACTIN FLOW UNDERLIES THE SURFING OF VIRUSES ALONG FILOPODIA OR MICROVILLI TOWARD THE CELL BODY Actin filaments at the leading edge and within cellular extensions, such as filopodia and microvilli, undergo a continuous cycle of assembly at their distal barbed end and disassembly at their proximal end. This actin treadmilling generates the flow of actin from the distal end toward the cell center, with or without the help of myosins [50, 51]. The retrograde flow of actin plays an important role in cell migration or axonal growth cone guidance, among other cellular processes [reviewed in Ref. 52]. The flow of actin has also been shown to underlie the retrograde motion of membrane receptors and their ligands, including the epidermal growth factor receptor along filopodia [53] and the nerve growth factor receptor at the axonal growth cone [54], as well as ricin and ferritin along microvilli [55, 56]. The efficient transport from the periphery to the cell body has been hijacked by viruses to propel their way along filopodia or microvilli toward the cell body, enhancing their infection of polarized and sparse cells in culture [57, 58]. Receptor-dependent surfing of fluorescently labeled viral particles along filopodia and microvilli has been observed prior to their fusion with the membrane or their entry within membrane vesicles at the cell body [57–59]. Moreover, certain viruses can induce dynamic changes in the actin network, leading to the formation of filopodia that enable their further recruitment from the extracellular environment to the cell body [60].

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19.5 SUBMICRO- AND NANOSCALE INORGANIC PARTICLES CAN BE PROPELLED ALONG FILOPODIA AND MICROVILLI Polymeric and inorganic particles, like viruses, have been shown to travel along filopodia and microvilli toward the cell body [61–63], pointing to a mechanism by which synthetic carriers can be actively recruited from the extracellular environment in cells that form filopodia or microvilli. Inhaled particles ranging from 5 nm to 1 μm that enter the respiratory tract are likely to reach the alveolar region [64, 65], where alveolar type II epithelial cells, which carry apical microvilli, are sparsely found. When grown in culture, these cells form microvilli that are shorter and crowded at the apical surface (Figure 19.1A) and are longer and less crowded at the basolateral surface (Figure 19.1B), where they can grow over the edge of the cell onto the bottom surface of the dish. Amorphous silica particles, which have been explored for drug delivery and medical imaging and sensing [66–69], can become engaged in a robust retrograde motion toward the cell body, once they land on these elongated microvilli [62]. Figure 19.2 demonstrates the retrograde motion of (A)

(B)

Figure 19.1. Scanning electron micrographs of alveolar type II epithelial cells grown in culture, showing the formation of microvilli that are shorter and crowded at the top of the cell (A) and are longer and sparser toward the periphery, where they can grow over the edge of the cell onto the glass surface of the dish (B). (Adapted with permission from Ref. 62.)

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Figure 19.2. A sequence of DIC images, selected from a time series [62, Movie 1], demonstrates the directed retrograde motion of positively charged 500-nm amorphous silica particles along the elongated microvilli of alveolar type II epithelial cells in culture. The particles are indicated by color-coded arrows as they appear along the time series, and the cumulative time relative to the first image in the series is shown in the lower right corner of each image. Particles are tracked from one image to the next in the series and their trajectories are plotted in the last frame, where the cell body from which the elongated microvilli originate is outlined with the dashed line. (Adapted with permission from Ref. 62.) (See color insert.)

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(A)

(B)

Figure 19.3. (A) An individual amorphous silica nanoparticle (≤100 nm), positively charged with amino groups and tagged with an average of 3 fluorescent molecules, is identified and tracked using single-molecule fluorescence techniques. The fluorescence images, selected from a time series [62, Movie 2], demonstrate the motion of the particle, indicated by the arrow, toward the cell body, which is outlined by the dashed line. (B) The trajectory of the nanoparticle is overlaid on the DIC image, showing that the particle travels toward the cell body along the elongated microvillus in an irregular pattern that includes both retrograde and anterograde motions. The origin of the trace is indicated by the arrow, and the area shown in the fluorescence images is marked by the dashed lines. (Adapted with permission from Ref. 62.)

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RETROGRADE MOTION IS MEDIATED BY TRANSMEMBRANE MOLECULES

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amine-modified positively charged 500-nm particles, indicated by color-coded arrows as they land and travel along the microvilli. Particles can dwell in their landing spot for a while or become engaged in a directed motion immediately after landing [62, Movie 1], eventually reaching the cell body where they can be internalized. Positively charged 100-nm amorphous silica particles can also become engaged in a retrograde motion. By tagging the nanoparticles with a small number of fluorescent molecules and using single-molecule fluorescence techniques [70] to determine the presence of single particles or quantify the number of particles within a cluster [62], it was possible to track individual nanoparticles over time (Figure 19.3A). Although clearly directed toward the cell body, the motion of the nanoparticles was found to include both rearward and forward motions [62, Movie 2], as shown in Figure 19.3B, where the time trajectory is overlaid on the DIC image. The distinct motion patterns of 500nm particles and 100-nm particles suggest that particle surface area and/or particle mass determine the robustness of the motion. Interestingly, unmodified bare amorphous silica particles, which carry negative surface charge, cannot become engaged in the retrograde motion [62]. This observation indicates that the motion of the particles is dependent on electrostatic interactions between negatively charged molecules in the plasma membrane and the positively charged surface of the particles.

19.6 RETROGRADE MOTION IS MEDIATED BY TRANSMEMBRANE MOLECULES THAT INTERACT DIRECTLY OR INDIRECTLY WITH ACTIN FILAMENTS Treating alveolar type II epithelial cells with cytochalasin D, a drug that caps the barbed end of the filaments and prevents actin polymerization, strongly inhibits the retrograde motion of the particles [62]. Cells that are trasfected with GFP-actin show fluorescent filopodia and microvilli, where individual fluorescent clusters are occasionally identified. These clusters can undergo a retrograde motion, which is also inhibited by cytochalasin D (Figure 19.4A). The inhibition of the motion correlates with the appearance of depolymerized and fragmented filaments (Figure 19.4A) [62, Movie 6b], which is in agreement with the actin treadmilling being the mechanism underlying the motion. By measuring the velocity of the particles and actin clusters along filopodia and microvilli it was found that both particles and clusters move at the same rate, about 12 nm/s [62], further supporting the role of actin flow in propelling the particles. The concerted motion of the particles and actin clusters is demonstrated in Figure 19.4B. Together with the observation that only positively charged particles can be propelled along filopodia and microvilli, it became clear that the interactions of the particles at the cell surface are mediated by a negatively charged transmembrane molecule that interacts either directly or indirectly with actin filaments.

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(A)

(B)

Figure 19.4. (A) Alveolar type II epithelial cells that are transfected with GFP-actin form fluorescent filopodia and microvilli that often contain small fluorescent clusters, which are distinguished from the diffused signal. Following the clusters over time reveals their retrograde motion toward the cell body, as demonstrated by the cluster that is indicated by the arrow in the first two images, which are selected from a time series [62, Movie 6b]. Treating the cells with cytochalasin D leads to fragmented filaments and the arrest of the retrograde flow, as demonstrated in the third and fourth images where the cumulative time relative to the time of applying the drug is shown in the lower right corner. (B) Images of a 500-nm fluorescent particle (red, white arrow) and a GFP-actin cluster (green, yellow arrow) demonstrate that the particle and the cluster travel toward the cell body at the same rate. (Adapted with permission from Ref. 62.) (See color insert.)

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HEPARAN SULFATE PROTEOGLYCANS PLAY A CRITICAL ROLE

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19.7 HEPARAN SULFATE PROTEOGLYCANS PLAY A CRITICAL ROLE IN THE ATTACHMENT AND INTERNALIZATION OF POSITIVELY CHARGED INORGANIC PARTICLES One of the most negatively charged family of molecules at the cell surface is the sulfated proteoglycan family. Sulfated proteogylycans consist of a core protein that is covalently linked to one or two types of glycosaminoglcans, which are long linear polysaccharide chains that bear negative charges [71]. The sulfated proteoglycans are a highly diverse family of molecules that are abundant in the extracellular matrix of all tissues and are also expressed at the cell surface of most cells. However, only a few sulfated proteoglycans are transmembrane molecules that potentially could couple the particles with acin filaments across the cell membrane. Among them are certain members of the heparan sulfate proteoglycan (HSPG) subfamily [71]. HSPGs, as a group, have been shown to mediate the cellular infection of certain viruses, such as herpes simplex virus [72], murine cytomegalovirus [73], HIV-1 [74], and human papilloma virus [75]. They have also been shown to mediate the internalization of polylysine– DNA and lipid–DNA complexes [76], or polycationic macromolecules and peptides designed for delivery of proteins and other molecules [77]. To determine whether hapran sulfate chains mediate the cellular interactions of positively charged inorganic particles, alveolar type II epithelial cells were preincubated with heparinase I and II, which cleave the heparan sulfate chains into the basic disaccharide repeats. Untreated and treated cells were exposed to positively charged 500-nm fluorescent amorphous silica particles and the degree of particle association with the cell was measured by flowcytometry [78]. Untreated cells that were incubated with the particles appeared as two distinct populations (Figure 19.5A, filled bars). The first population overlapped with cells that were not exposed to particles at all [shown in Ref. 78], indicating that only a subpopulation, showing at higher fluorescence intensity values, became associated with the particles. Trypan blue, which is a membrane impermeable dye, can quench the fluorescent particles at the cell surface but leaves intact the fluorescent particles inside the cell. Incubating the cells with trypan blue (Figure 19.5A, open bars) shifted the second population to lower values, indicating that about half of the particles [calculations in Ref. 78] were already internalized and protected from quenching within the intracellular environment. However, heparinase treatment decreased dramatically the degree of particle association with the cells (Figure 19.5B, open bars), when compared with untreated cells (Figure 19.5B, filled bars), and almost completely inhibited the internalization of the particles, as indicated by the disappearance of the second population after incubation with trypan blue (Figure 19.5C, filled bars). Treating the cells with chondroitinase ABC, which cleaves the chondroitin sulfate chains that are carried by certain transmembrane proteoglycans, also decreased the degree of particle association with the cells (Figure 19.5D, open bars) when compared with untreated cells (Figure 19.5D, filled bars), although not nearly as drastically as the treatment with heparinase. About half of the particles that became associated with

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Figure 19.5. (A) The flow cytometry histogram of cells that are exposed to positively charged fluorescent 500-nm particles (filled bars) shows two distinct cell populations, where the first population overlaps with cell that were not exposed to the particles (not shown here). The second population, at higher fluorescence intensity values, consists of cells that become associated with the fluorescent particles. Using trypan blue (open bars) to distinguish between particles at the cell surface and those that are protected within the cytoplasm it is found that about half of the particles are internalized, as indicated by the shift of the second population [calculations in Ref. 78]. (B) Treating the cells with heparinase (open bars) leads to a dramatic decrease in the degree of particle association with the cells, when compared with untreated cells (filled bars), as indicated by the significant degrease in the second population. (C) Incubating heparinase-treated cells (open bars) with trypan blue (filled bars) leads to a near complete disappearance of the second peak, indicating minimal particle internalization under heparinase treatment. (D) Treating the cells with chondroitinase (open bars) leads to a smaller but significant decrease in the degree of particle association with the cell, when compared with untreated cells (filled bars). (Adapted with permission from Ref. 78.)

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chondroitinase-treated cells were internalized, as determined using trypan blue [78]. These observations revealed a critical role for heparan sulfate chains and a smaller but clear role for chondroitin sulfate chains in capturing and mediating the internalization of positively charged synthetic amorphous silica particles in alveolar epithelial cells bearing microvilli. Among the HSPGs are the syndecans, a small group of four transmembrane proteoglycans [79, 80] that carry mostly heparan sulfate chains but also chondroitin sulfate chains at their extracellular domain [81–83], which could mediate the interactions and internalization of the particles.

19.8 SYNDECAN-1 MEDIATES THE COUPLING OF POSITIVELY CHARGED INORGANIC PARTICLES WITH ACTIN FILAMENTS ACROSS THE CELL MEMBRANE Syndecan-1, which is expressed predominantly in epithelial cells and carries both heparan sulfate and chondroitin sulfate chains at the extracellular domain [81–83], interacts indirectly with actin filaments [84–86]. These properties have made syndecan-1 a likely candidate for mediating the interactions of the particles and their coupling with the filaments across the cell membrane. Using a fluorescent antibody specific to the extracellular domain of the core protein of syndecan-1 and applying time-lapse multicolor fluorescence imaging, it was possible to follow syndecan molecules and particles simultaneously over time. Using this approach, it was found that the particles and syndecan molecules become engaged in a directed motion along microvilli once they are tightly associated with each other [78, Movies 1 and 2]. Figure 19.6A demonstrates the engagement of the particles and syndecan molecules in a directed motion as they overlap on the microvillus, which is indicated by the appearance of the yellow color (also see Figure 5 in Ref. 78). Since the directed motion is powered by actin flow, the engagement of the particles and syndecan molecules in the directed motion unravels their coupling with actin filaments across the cell membrane, as illustrated in Figure 19.6B. The four syndecans have an identical PDZ binding motif at their cytoplasmic C terminus [80], which allows them to bind PDZ proteins, such as syntenin [87–89] or CASK [90, 91]. Syntenin and CASK, in turn, bind actin via actin binding proteins, such as merlin [92, 93] or protein 4.1 [90], respectively. The association of syndecan molecules with actin requires their clustering or crosslinking [84–86]. The clear engagement of syndecan molecules in a directed motion following their interaction with the particles [78, Figure 5 and Movie 2] suggests that the interaction with the particles facilitates the clustering or crosslinking of the molecules and their subsequent association with actin. This interpretation is supported by the observations that the interactions of syndecan molecules with extracellular matrix molecules induce their clustering and association with actin, which has been suggested to anchor the cytoskeleton to the extracellular matrix [84, 86, 94].

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(A) 2

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Figure 19.6. (A) A DIC image of the elongated microvillus and a sequence of fluorescence images, selected from a time series [72, Movie 1], demonstrate the engagement of the particle (red), indicated by the arrow, and syndecan-1 molecules (green) in the motion along the microvillus as they become tightly associated (yellow) with each other. (B) The directed motion along the microvillus unravels the coupling of syndecan-1 and the particle with actin filaments across the cell membrane via docking and actin binding proteins. See discussion in the text. (Adapted with permission from Ref. 78.) (See color insert.)

19.9 POSITIVELY CHARGED INORGANIC PARTICLES CAN INDUCE SYNDECAN-1 CLUSTERING AND SUBSEQUENT ACTIN COUPLING AND INTERNALIZATION The clustering of syndecan molecules is required for both their association with actin [84–86] and their internalization [95, 96]. The internalization of syndecans and their cargo can occur via macropinocytosis [97, 98], a fluid-

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phase endocytic process associated with a dynamic rearrangement of actin filaments and the induction of membrane ruffling or blebbing that collapse and fuse with the plasma membrane to form membrane vesicles [reviewed in Refs. 38 and 99]. The increase in actin dynamics and the formation of membrane protrusions are initiated by the activation of growth factor receptors and other external stimuli, including certain bacteria and viruses [reviewed in Refs. 38 and 99]. Vaccinia virus, for example, initiates the massive formation of blebs that, in turn, mediate the internalization of the virus by macropinocytosis [59]. Positively charged 500-nm amorphous silica particles also enter alveolar type II epithelial cells by macropincytosis [78]. The majority of the particles were found colocalized with 70-kDα dextran, a marker for macropinocytosis, and their internalization was completely inhibited by dimethyl amiloride hydrochloride, an inhibitor of macropincocytosis [78, Figures 7 and 8]. Importantly, syndecan-1 and the majority of the particles were found colocalized within small membrane vesicles, indicating that syndecan and the particles are cointernalized [78, Figure 6]. As no internalization was observed when the cells were treated with heparinase, these observations support a role for syndecan-1 molecules in mediating the internalization of the particles via their own macropinocytotic pathway. Together, the observations identify a role for syndecan-1 in mediating the interactions of the particles at the cell surface, their coupling with the intracellular environment, and their subsequent internalization via macropinocytosis. Since the coupling of syndecan-1 molecules to actin as well as their internalization require their clustering or crosslinking [84–86, 96, 97], these observations also support the idea that the particles initiate their own internalization by facilitating the clustering or crosslinking of syndecan-1 molecules around them and their subsequent actin coupling and internalization. Syndecan-1 was found to mediate the internalization of Opa expressing Neisseria gonorrhoeae [100] and certain types of human papillomavirus [101, 102], as well as certain cationic liposome–DNA complexes [103]. Whether these pathogens and liposomes also induce the clustering of syndecan molecules and their subsequent actin coupling and internalization is unclear. The new understanding of the mechanism by which syndecan-1 mediates the retrograde recruitment of positively charged inorganic particles from the extracellular environment and their internalization at the cell body [62, 78] provides new clues for designing synthetic carriers for efficient delivery to cells bearing apical microvilli.

19.10 CONCLUSION The ability of viruses and other pathogens to exploit the cellular transport systems has inspired the design of artificial viruses or synthetic carriers for effective drug and gene delivery. New synthetic particles that can surf toward the cell body, penetrate the cortical actin filament barrier, and ride on microtubules and actin filaments within and between the cytoplasm and the nucleus

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could increase significantly the success and efficiency of their drug or gene delivery. A better understanding of the molecules and mechanisms that underlie the transport systems will enable the design of synthetic particles that are capable of assimilating into the cellular environment with minimal adverse effects while being equipped to use the endogenous machinery.

ACKNOWLEDGMENT This work was supported by the EPA STAR grant RD833338 to GO, the Air Force Research Laboratory grant FA8650-05-1-5041 to ONAMI-SNNI, and the National Institute of Environmental Health Sciences (NIEHS) grant RC2ES018786-02 to GO. The research was performed using EMSL, a national scientific user facility sponsored by the DOE’s Office of Biological and Environmental Research and located at Pacific Northwest National Laboratory.

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90. Cohen, A. R., Woods, D. F., Marfatia, S. M., Walther, Z., Chishti, A. H., and Anderson, J. M. Human CASK/LIN-2 binds syndecan-2 and protein 4.1 and localizes to the basolateral membrane of epithelial cells. J. Cell Biol. 142: 129–138 (1998). 91. Hsueh, Y. P., Yang, F. C., Kharazia, V., Naisbitt, S., Cohen, A. R., Weinberg, R. J., and Sheng, M. Direct interaction of CASK/LIN-2 and syndecan heparan sulfate proteoglycan and their overlapping distribution in neuronal synapses. J. Cell Biol. 142: 139–151 (1998). 92. Kang, B. S., Cooper, D. R., Jelen, F., Devedjiev, Y., Derewenda, U., Dauter, Z., Otlewski, J., and Derewenda, Z. S. PDZ tandem of human syntenin: crystal structure and functional properties. Structure 11: 459–468 (2003). 93. Scoles, D. R. The merlin interacting proteins reveal multiple targets for NF2 therapy. Biochim. Biophys. Acta. 1785: 32–54 (2008). 94. Bernfield, M. and Sanderson, R. D. Syndecan, a developmentally regulated cell surface proteoglycan that binds extracellular matrix and growth factors. Philos. Trans. R. Soc. Lond. B Biol. Sci. 327: 171–186 (1990). 95. Fuki, I. V., Meyer, M. E., and Williams, K. J. Transmembrane and cytoplasmic domains of syndecan mediate a multi-step endocytic pathway involving detergentinsoluble membrane rafts. Biochem. J. 351: 607–612 (2000). 96. Fuki, I. V., Kuhn, K. M., Lomazov, I. R., Rothman, V. L., Tuszynski, G. P., Iozzo, R. V., Swenson, T. L., Fisher, E. A., and Williams, K. J. The syndecan family of proteoglycans. Novel receptors mediating internalization of atherogenic lipoproteins in vitro. J. Clin. Invest. 100: 1611–1622 (1997). 97. Tkachenko, E., Lutgens, E., Stan, R. V., and Simons, M. Fibroblast growth factor 2 endocytosis in endothelial cells proceed via syndecan-4-dependent activation of Rac1 and a Cdc42-dependent macropinocytic pathway. J. Cell Sci. 117: 3189–3199 (2004). 98. Nakase, I., Tadokoro, A., Kawabata, N., Takeuchi, T., Katoh, H., Hiramoto, K., Negishi, M., Nomizu, M., Sugiura, Y., and Futaki, S. Interaction of arginine-rich peptides with membrane-associated proteoglycans is crucial for induction of actin organization and macropinocytosis. Biochemistry 46: 492–501 (2007). 99. Mercer, J. and Helenius, A. Virus entry by macropinocytosis. Nat. Cell Biol. 11(5): 510–520 (2009). 100. Freissler, E., Meyer auf der Heyde, A., David, G., Meyer, T. F., and Dehio, C. Syndecan-1 and syndecan-4 can mediate the invasion of OpaHSPG-expressing Neisseria gonorrhoeae into epithelial cells. Cell Microbiol. 2: 69–82 (2000). 101. Shafti-Keramat, S., Handisurya, A., Kriehuber, E., Meneguzzi, G., Slupetzky, K., and Kirnbauer, R. Different heparan sulfate proteoglycans serve as cellular receptors for human papillomaviruses. J. Virol. 77: 13125–13135 (2003). 102. de Witte, L., Zoughlami, Y., Aengeneyndt, B., David, G., van Kooyk, Y., Gissmann, L., and Geijtenbeek, T. B. Binding of human papilloma virus L1 virus-like particles to dendritic cells is mediated through heparan sulfates and induces immune activation. Immunobiology 212: 679–691 (2007). 103. Mounkes, L. C., Zhong, W., Cipres-Palacin, G., Heath, T. D., and Debs, R. J. Proteoglycans mediate cationic liposome-DNA complex-based gene delivery in vitro and in vivo. J. Biol. Chem. 273: 26164–26170 (1998).

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CHAPTER 20

Subcellular Targeting of VirusEnvelope-Coated Nanoparticles JIA WANG, MOHAMMAD F. SAEED, ANDREY A. KOLOKOLTSOV, and ROBERT A. DAVEY Department of Microbiology and Immunology, University of Texas Medical Branch, Galveston, Texas

20.1

INTRODUCTION

A major problem in drug delivery is the compromise that must be made in having a drug freely soluble in the circulation, while still being able to cross the hydrophobic barrier of the cell membrane (unless acting on a receptor). A drug must balance these factors to be effective. Furthermore, permeability barriers between different tissues can further restrict the usefulness of a drug. This is a particular problem with the blood–brain barrier, where compounds must be lipophilic to gain access to the brain [1]. The ideal delivery scenario is to have a drug encapsulated, minimizing unwanted interactions with cells until it accesses the target tissue and cell type. Microencapsulation of drugs at the nanoscale provides a way to engineer a new generation of drugs that remain inert until interacting with the target cell. Functionalized capsules will also allow specific targeting of drugs to cells, which means the overall dosage of drug can be reduced while increasing the therapeutic index. Encapsulation can also promote increased circulation times without increased toxicity by avoiding metabolic clearance by the liver and kidneys. To achieve these goals capsulation methods need to be developed that coat drugs with biocompatible materials. These materials should be biodegradable, tolerated by the body, enhance targeting to cells, and, importantly, deliver the drug into the cell cytosol. Viruses are nanoscale mechanisms that have been perfected through coevolution with their hosts to have long circulation times while protecting their Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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core containing the virus genome from nuclease and protease degradation. They efficiently seek out cells that can support their replication by interacting with specific receptors and then inject the virus genome into the cell cytoplasm while minimizing toxicity or triggering innate or adaptive immune responses. For enveloped viruses, targeting and delivery are mediated by the envelope glycoproteins (eGPs), which not only bind cell receptors but sense the environment and, when triggered, catalyze the fusion of the cell and virus lipid bilayers to release the capsid cargo into the cytoplasm. These systems are so finely tuned that they can target specific steps of the cellular endocytic pathway so that the virus genome is only released in ideal subcellular locations suitable for virus replication. In this chapter we will discuss the potential of eGPs for drug delivery and discuss approaches to harness mammalian virus eGPs as a highly efficient way of targeting cells and subcellular compartments and delivering lipid-coated nanoparticles into the cell cytoplasm.

20.2

VIRUS CLASSIFICATION

Mammalian viruses are classically divided by morphological, biochemical, and genetic criteria. When examined closely by electron microscopy, viruses appear spherical or filamentous (Figure 20.1). Careful examination reveals that spherical viruses usually have an icosahedral structure while filamentous

Figure 20.1. Electron micrographs of enveloped viruses. Enveloped viruses are diverse in structure and range from spherical particles to fibers. Examples are shown for murine leukemia virus, a spherical retrovirus, vesicular stomatitis virus, which has a “bullet” shape, and Ebola virus, which is filamentous and forms rings (as shown) and rods. Transmission electron microscopy was used for the murine leukemia virus and Ebola virus images and cryoelectron microscopy was used for the image of vesicular stomatitis virus. The scale bar is 100 μm for each image. The capsid is indicated by the asterisk and the lipid bilayer (best seen in the cryoelectron microscopy image) is indicated by the arrowhead. In each case the eGPs project from the lipid bilayer but are poorly visible.

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viruses are rod-like. For spherical viruses, size varies from 30 nm for the parvo and picornaviruses, up to 300–450 nm for the poxviruses of which smallpox is best known. Filamentous viruses can be much larger in length, extending up to 1 μm with cross-section diameters of around 100 nm. The appropriately named filoviruses (filo, derived from the Latin for filament) are good examples of this virus type and are represented by Ebola virus. Between each of these extremes is an extensive set of virus families that have intermediate sizes and shapes averaging 100–200 nm. Further divisions are made on the basis of genetic structure. All viruses contain a genome made from RNA or DNA or a mix of both. The genetic material can be single or double stranded and present on a single or multiple segments of nucleic acid. Biochemical composition is a third criterion used to classify viruses and is the most relevant to later sections of this chapter. All viruses contain protein, either encoded by the virus genome or taken from infected cells, but a major distinction is the presence of a bounding lipid bilayer that is also derived from the cell membrane when virus buds from the cell. Viruses that have lipid membranes are termed enveloped viruses and will be the main subject of this chapter. They are special, as embedded in the membrane are the viral eGPs. Often called “spike” proteins, the eGPs project from the virus membrane and are responsible for the initial interaction with the cell as well as catalyzing fusion of the virus and cell membrane. Since the lipid bilayer encapsulates the virus core, fusion of virus and cell membranes results in the seamless merging of virus and cell membranes to release the core into the cell cytoplasm. The ability of the eGP to induce membrane fusion of lipid bilayers, and the fact that the majority of eGPs are structurally distinct functional units that operate independently from the other virus proteins that underlie the membrane, offers great advantages over nonenveloped virus proteins since the size and composition of the cargo are not restricted and simply need to be bound in a lipid bilayer.

20.3 ENVELOPE GLYCOPROTEIN STRUCTURE, FUNCTION, AND CLASSIFICATION eGPs are multifunctional proteins that bind to specific surface-exposed receptors on cells and allow the virus to penetrate into the cell cytoplasm. Both these properties are highly desirable aspects of a nanocargo delivery mechanism. eGPs comprise three structural classes and have been discussed extensively in other review articles [2, 3], so only a brief summary is given with greater emphasis on details relating to nanoparticle production and cell delivery. Class 1 eGPs are type 1 membrane proteins with N termini that extend outside the virus particle, have one alpha-helical transmembrane domain that spans the viral membrane, and a C terminus inside the virus lumen. A surface domain, comprising the receptor binding motif, is connected to the transmembrane domain by noncovalent (hydrogen bonding) as well as disulfide

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bonds between cysteine residues [4]. The transmembrane domain contains a fusion peptide at its tip, which is normally buried under the surface domain. When triggered, the surface domain moves away to reveal the fusion peptide, which in turn moves up and away from the virus membrane [5]. Dimers or trimers of eGP monomers come together to form distinct functional units that can be seen by electron microscopy as spike proteins. Class 1 eGPs are well suited for coating of nanoparticles (as discussed later) as each eGP oligomer can be separated from other eGP oligomers and still remain active. While Class 2 eGPs are also connected to the virus membrane by alpha-helical transmembrane segments, they differ from Class 1 proteins by having an interlocking structure in which eGP dimers or trimers interact through extensive hydrogen bonding networks to cover the surface of the virus particle [6]. The fusion peptide is present as a loop between eGP domains and is moved toward the cell membrane by a scissoring action. Flavivirus [7] and togavirus [8] eGPs are examples of this form. The interlocking nature of Class 2 eGPs makes them difficult to use in a recombinant system as they cannot be easily separated from neighboring eGPs and are geometrically constrained, limiting the size of particle that can be produced. Class 3 eGPs are a recently recognized group of eGPs that are also embedded in the virus membrane by a transmembrane alpha-helix but are more similar to Class 1 proteins with eGP monomers forming trimers that behave as distinct functional units. They differ from Class 1 proteins in having a distinct structural conformation where the membrane fusion domains are formed from multiple fusion peptides that come together when triggered. The eGPs of vesicular stomatitis virus are representative of this class and are unique in having the ability to undergo a reversible conformation change that alternates between pre- and postmembrane penetration states [9].

20.4

VIRUSES AS MEMBRANE PENETRATION DEVICES

The plasma membrane of all cells is composed of a stable lipid bilayer that cannot be easily broken [10]. For virus infection to occur this bilayer must be breached in a way that does not kill the cell. This is a significant hurdle and is not easily achieved in most systems used today. All viruses perform this task efficiently by altering the physicochemical properties of the virus surface proteins from hydrophilic to hydrophobic and altering the structure of the lipid bilayer of the cell. Nonenveloped viruses, that lack a membrane of their own, break the cell membrane by pore formation through interaction of surfaceexposed capsid proteins with the cell membrane. This often occurs from within an endosome and involves the concerted action of proteins over the capsid surface to open a pore in the cell membrane [11]. Harnessing this type of mechanism for cargo delivery into cells is difficult as each protein is structurally constrained by its neighbors and the mechanism is not well understood. In contrast, the fusion process of enveloped viruses is well understood (Figure 20.2) and often used as a paradigm for how membrane fusion is induced in

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Figure 20.2. Virus anatomy and the three stages of eGP-mediated delivery into cells. Virus particles and eGP-coated nanoparticles each interact with cells identically. The top image indicates the parts of a virus and show interaction with cells through eGP binding to specific receptors. eGPs are shown as trimers, which is typical for most Class 1 and 3 viruses. Actin is often involved in recruiting receptor. The particle is then taken into endosomes (for pH-dependent viruses), where the pH of the vesicle is made progressively more acidic as it moves through the cell, maturing from an early to late endosome. The eGP functions as an environmental sensor and when triggered undergoes a large conformational change to expose previously buried fusion peptides. These embed in the cell membrane and catalyze membrane fusion of the virus and endosomal membrane. Since this occurs at a specific pH, the core is delivered to a specific subcellular location within the cell. For pH-independent viruses, like HIV, membrane fusion may occur at the cell surface but this has recently been challenged and it too may use endosomes but is triggered by other stimuli [50]. (See color insert.)

everyday biological processes [12]. Enveloped viruses breach the cell membrane by inducing membrane fusion between the cell and virus membranes. In this case hydrophobic fusion peptides are made to penetrate into the cell membrane, initially causing the outer leaflets of each membrane to break and mix, after which a pore forms where each leaflet of the membrane merges.

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The pore then widens and the virus core moves into the cell cytoplasm. The eGP acts as a mechanism to bring the membranes together by undergoing large conformational changes as well as a catalyst to promote lipid mixing [12]. Using this method of entry, enveloped viruses avoid widespread disruption of cell membranes and potential release of endosomal contents into the cell cytoplasm. Together, eGPs from enveloped viruses are well suited for cargo delivery into cells. 20.4.1 Receptor Interaction Virus binding to cells is through receptor recognition, where surface-exposed portions of the eGP serve as a ligand. Binding affinities range from picomolar to nanomolar. However, the interaction of multiple eGPs within oligomers and multiple oligomers with multiple receptors on cells results in a very high avidity binding. This ensures that once an eGP oligomer binds to a cell with the appropriate receptor density, the particle remains bound. A diverse range of receptor types are bound by eGPs; however, each virus has high selectivity for a single or set of receptors. The types of receptors used by viruses range from carbohydrate modifications on proteins [13], GPI anchored proteins [14], single span membrane proteins [15], multipass membrane proteins [16], and lipids themselves [17]. A nonexhaustive list of some of the receptors bound by different virus eGPs is given in Table 20.1. No particular receptor form appears to dominate and it is unclear if the physical properties of each membrane protein play a key role in the entry process. Indeed, in some cases receptor function can be transferred into nonreceptor proteins by genetic engineering, but this is not possible in many circumstances [33]. However, it is clear that the eGP– receptor interaction defines the host range of virus particles and this is mediated through a specific eGP receptor binding domain. Receptor choice is also dictated by the infection route favored by the virus. The abundance of sialic acid in the airways of the lungs is likely an important factor for its use in infection by influenza viruses [34]. Similarly, the choice of CCR5 as a coreceptor for HIV is important for its infection of mucosal macrophages which express this receptor [15]. Class 1 and 2 eGPs have distinct receptor interaction domains that have been identified through mutagenesis and high-resolution structural analysis. In each case, the receptor binding domain resides at or close to the top of the surface domain of the eGP where it can interact with receptors. Many of the receptor binding domains have immunoglobulin-like folds [35, 36], indicating that they are highly stable structures resistant to protease degradation. Like immunoglobulins, the portions interacting with the cell receptor are presented in loops and helices that line the surface of the fold [35]. Due to the modular construction of the eGPs it is possible to generate recombinant receptor binding forms that retain receptor binding function [37]. In these situations the receptor specificity of the virus can be transferred to a nanoparticle cargo

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TABLE 20.1

Examples of eGP Defined Receptor Specificity and Cell Tropism for Enveloped Viruses Virus

Receptor Useda

Cell Tropismb

Alpha v beta 3 integrin Laminin receptor CD4 and CXCR4/CCR5

Coronavirus

West Nile virus Sindbis virus Human immunodeficiency virus Ecotropic murine leukemia virus Ecotropic murine leukemia virus SARS virus

Filovirus Rhabdovirus Bunyavirus

Ebola virus Vesicular stomatitis virus Hantaan virus

Broad Broad T cells and macrophages Mouse cells, excluding liver Now infects EpoR+ cells Lung, intestine, kidney, liver Dendritic cells Broad Broad

Orthomyxovirus

Influenza A virus

Paramyxovirus

Measles virus

Arenavirus Hepadnavirus

Lassa fever virus Junin virus Hepatitis B virus

Herpesvirus

Herpes simplex 1 virus

Virus Family Flavivirus Togavirus Retrovirus

a

mCAT-1, a 14 spanning transmembrane protein Modified to bind EpoR ACE-2 Folate receptor, Axl Lipid Alpha v beta 3 integrin receptor Sialic acid modifications on proteins SLAM or CD46 α-Dystroglycan Tranferrin receptor CD81, SR-B1, occludin, and claudin-1 HVEM, Nectin-1

Routes of Infection

Reference

Through skin Through skin Mucosa

18 19 15, 20

Germ line or blood

21

Blood

22

Lung and oral

23, 24

Through skin Through skin Lung

25, 26 17 27

Broad

Lung

13

Broad

28

Broad Broad Liver cells

Lung and then other tissues Mucosa Through skin blood

29 30 31

Neuronal

Through skin

32

For some viruses multiple receptors have been identified. It is unclear if receptor usage varies by cell type or if a receptor complex is needed to infect cells.

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for targeting purposes, just like any other peptide ligand. Making artificial ligands based on virus proteins has become popular with the development of recombinant single chain antibody–virus eGP fusions. These constructs have allowed an extensive characterization of the receptor interactions of several virus eGPs [38]. The advantage of such “immunoadhesins” is that they behave like antibodies, being divalent and maintain binding with protein A and G making detection and conjugation straightforward and generic. However, these constructs lack the ability to catalyze membrane fusion. Conversely, some success has been seen in modifying eGP receptor specificity, while maintaining eGP membrane fusion activity. This has the advantage of maintaining the potent delivery activity of the eGP while giving novel receptor specificity. Small ligands that are functional in loops have been the most effective. These include epidermal growth factor [39, 40], erythropoietin [22], the RGD sequence that binds integrins [41], and insulin-like growth factor [42]. Other methods have been developed to incorporate single chain antibodies into eGPs, giving a generic platform in which novel binding specificities can be inserted [43]. Unfortunately, these approaches have not been as simple as expected since it appears in many cases that receptor specificity and activation are closely tied, perhaps due to lack of environmental triggering encountered when using specific receptors or disruption of eGP structure [44]. 20.4.2 eGPs as Environmental Sensors The trigger that stimulates an eGP to undergo the conformational change toward the membrane fusion-mediating state is most often pH. The evidence for the role of pH comes from the use of drugs that block endosomal acidification through inhibition of proton pumps or by buffering the pH gradient established by the action of these pumps. By protonating amino acid side chains of the eGPs, one set of hydrogen bonds is weakened and the profusion state is adopted [45]. For Class 1 and 2 eGPs, this change in conformation occurs with a release of energy, which is harnessed to promote membrane fusion and is irreversible [2]. Class 3 proteins are unusual in undergoing a reversible conformational change but are nevertheless potent membrane fusion inducers. For most pH-sensitive eGPs, exposure to an acidic medium, as occurs in endosomes, is sufficient to induce the profusion state. Brief exposure of these viruses to an acidic medium will trigger membrane fusion to cells or artificial membranes like liposomes [46]. Such eGPs would be useful where a cargo was to be delivered to multiple target types. Exceptions do exist with some avian retroviruses requiring receptor binding to take place before being responsive to a drop in pH [47]. This two-stage trigger mechanism has advantages where the fusion event needs to be controlled more precisely. Another two-stage trigger mechanism that is more commonly employed by multiple virus types is the need for proteolytic activation of the eGP. In this case the eGP is locked in a state unable to undergo the comformational rearrangement needed for the profusion state. However, after interaction with cells the eGP becomes cleaved by cellular protease (often between the surface

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and transmembrane polypeptides) and the eGP becomes responsive to an acidic environment [48]. Viruses that are not sensitive to the actions of endosomal acidification inhibitors are considered to be pH insensitive. This does not preclude the use of endocytosis to infect cells [49, 50], but means that they are responsive to triggers other than acidification. Retroviruses (like HIV) and paramyxoviruses (like measles) are examples of pH-insensitive enveloped viruses. Generally, it is thought that receptor interaction alone is sufficient to trigger the eGP into the profusion state, although little direct evidence for this has been reported [50]. Some reports have suggested response to changes in redox potential at the cell membrane or in endosomes through the action of membrane-bound disulfide isomerase enzyme may be necessary. However, more work is needed to confirm this as a general mechanism of eGP activation [51, 52]. For purposes of harnessing eGPs for nanoparticle cargo delivery into cells, the pH-dependent virus eGPs are more likely to be useful. While pHindependent viruses are thought to share similar mechanisms of membrane fusion, much less is known about how they are triggered and which parts of the eGP are necessary to sense a trigger stimulus. In contrast, the mechanism of action of pH-dependent eGPs is better studied and understood, with highresolution structures available of pre- and postmembrane fusion forms for eGPs of each structural class available [53]. For the pH-dependent eGPs, the structural data also makes possible the design and reengineering of an eGP to have desirable properties (such as specific receptor interactions) without disruption of function. 20.4.3 eGP Targeting to Endocytic Pathways While receptor binding can dictate the cell type that is targeted, the ability of an eGP to give controlled delivery of particles into the cell cytoplasm is equally important. The rationale behind why a virus chooses a particular receptor or set of receptors is complex but is partly defined by the types of cells that can support virus replication. Another factor in receptor choice is access to specific endosomal trafficking pathways that a receptor moves through. Just as receptor choice has been optimized to identify specific cells that can support virus replication, receptors are also chosen that can deliver a virus particle into a specific endosomal environment. The eGP thereby provides a novel mechanism to target not only a cell but also a particular endocytic pathway and subcellular destination. Endocytosis encompasses distinct pathways used for uptake and movement of extracellular materials, such as nutrients, solutes, and ligand–receptor complexes, in lipid-bound vesicles. The process begins with the invagination of the plasma membrane around the cargo to be internalized, which subsequently detaches, internalizing the cargo into a membrane-bound endocytic vesicle. The cargo is then trafficked through various vesicular compartments by successive fusion of the endocytic vesicle membrane to that of the next. Ultimately, the cargo reaches its destination where the contents are released. To date,

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about ten different endocytic routes have been recognized in mammalian cells. They are distinguished from each other by a number of criteria including morphology of vesicles, the type of cargo they carry, and the cellular factors involved in their production and movement [54]. 20.4.4 Endocytic Routes Targeted by eGPs As described above, the eGP defines receptor usage but the choice of receptor also defines the route of endocytic uptake into the cell. Endocytosis can be through passive means when the eGP attaches to receptors that are constitutively taken up into the cell, as in the case of receptors bound by influenza A through sialic acid [13] or through active recruitment and internalization. The role eGPs play in cell signaling to induce receptor endocytosis is just now being appreciated and it is likely that many virus eGPs play some role in virus uptake [55, 56]. Different viruses employ different routes; however, three pathways are relatively better understood in terms of virus entry. They are clathrin-mediated endocytosis (CME), caveolin-mediated endocytosis (CavME), and macropinocytosis [57]. Other routes, even though targeted by virus eGPs, are not well understood and are generally classified as nonclathrin–noncaveolar endocytosis (NCE). It is important to note that a given eGP may target more than one route to enter cells, depending on the host cell type and/or other factors. CME is the oldest recognized endocytic pathway. The internalization of cargo (ligand bound to receptor) occurs in specialized areas of cell membrane called clathrin-coated pits (CCPs). CCPs are formed on the cytoplasmic side of the plasma membrane through sequential assembly of various components including the protein clathrin that forms a cage-like structure lining the pit. With the binding of cargo to its receptor, the growing pit invaginates and is eventually severed from the plasma membrane, by action of the protein dynamin, to form a clathrin-coated vesicle that contains the internalized cargo. The clathrin coat constrains the size of the CCP to 100–300 nm in diameter and so CCPs have limited capacity but many pits form per second. This means that cargoes need to be constrained in size to access this pathway. After moving into the cell the vesicle sheds the clathrin coat, a prerequisite for further trafficking and fusion with other compartments. A number of accessory, adaptor, and signaling molecules partake in this process and provide a tight regulation of the pathway. Some, such as accessory protein-2 (AP2) and Eps15, are specifically associated with CME, while others such as dynamin are involved in other endocytic pathways [54]. CME appears to be the most commonly used route targeted by eGPs. For a variety of viruses, such as influenza A, Semliki Forest, and vesicular stomatitis viruses, the eGP targets CME as the primary route of internalization. Even though CME occurs constitutively in cells, receptors can be induced to access this pathway by a number of ligands including eGPs. Influenza virus particles enter through CCPs that assemble underneath the particles after receptor

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binding [58]. This induction in CME is believed to be due to virus-induced signaling. siRNA screens have identified a variety of cellular kinases that regulate processes important in cytoskeleton rearrangement, cell cycle and cell growth, and endocytic trafficking after virus stimulation [59]. Cargoes internalized through CME are delivered to early endosomes (EEs). These organelles are progressively acidified during transit and reach a pH of 6.0–6.5. The GTPase Rab5 is considered a marker of the early endosome and its GFP-tagged form is effective at visualizing EEs in live cells [60]. From EEs, cargoes are transferred to late endosomes (LEs), which become more acidic. GFP-tagged Rab7, another GTPase that transiently associates with endosomes, can be used as a live marker of LEs [60]. As discussed above, eGPs can sense and react to environmental pH. This simple device thereby allows the eGP to deliver its cargo precisely at a point along the endosomal trafficking chain. Until recently, EEs were considered to be homogeneous. However, it is now clear that subpopulations of EEs exist that serve distinct purposes and can carry cargoes to distinct subcellular destinations. Comparison of influenza virus and Semliki Forest virus (SFV), both of which are internalized through CME, shows that influenza virus-containing EEs rapidly move to the perinuclear region, where membrane fusion occurs [61] whereas SFV membrane fusion occurs after transfer to late endosomes and occurs close to the endoplasmic reticulum [62]. CavME was first observed for endocytosis of SV40, a nonenveloped virus [63]. Few eGPs have been shown to target this pathway for entry; the eGP of murine leukemia viruses [64] seems to be an exception showing good colocalization with markers in this pathway. However, because of its importance in receptor-mediated vesicle uptake, other enveloped viruses are expected to be identified as using this entry route in future work. CavME is distinct from CME, both in terms of internalization process and vesicular transport. CavME occurs through caveolae, which are small flask-like invaginations in the plasma membrane enriched for the protein caveolin and associated with cholesterolrich membrane microdomains such as lipid rafts. They range in size from 80 to 120 nm in diameter, which means that only small cargoes can be taken up. It is believed that lipid rafts serve as organizing centers for CavME, bringing the receptor, coreceptors, and other signaling proteins together, facilitating the induction of processes associated with lipid rafts including CavME. Some cellular factors such as caveolin and Eps15R are specific for CavME, while others such as dynamin are shared. GFP-tagged caveolin can be used to visualize caveolae in live cells and this was used to demonstrate that murine leukemia virus eGP-coated nanoparticles entered cells through caveolae, similar to the eGP donor virus [65]. Other cellular factors including phosphoinistide-3 kinase, Rho family GTPases, and actin have also been implicated in CavME function [55]. Unlike CME, which can be constitutive or induced, CavME is induced by ligand (eGP)–receptor engagement [66] and a number of cellular kinases were found to be involved in the CavME of SV40. Interestingly, there was very little overlap between these and the kinases involved in CME-dependent entry of

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vesicular stomatitis virus [59]. This demonstrates the capacity of virus proteins to induce alternate signaling pathways that can further mediate particle uptake into cells by distinct routes. The destination of cargoes internalized through CavME differs considerably from CME. As opposed to CME where the virus is delivered to low-pH EEs, caveolar endocytosis leads to neutral pH vesicles. These vesicles, referred to as caveosomes, are morphologically and biochemically distinct from EEs. They are devoid of the CME markers EEA1 and Rab5 and are enriched in cholesterol and caveolin. From the caveosome cargo travels along the microtubules and is delivered to the endoplasmic reticulum [66] or to recycling endosomes as seen with mouse polyomavirus [67]. Macropinocytosis is a third endocytic uptake pathway. Until recently, macropinocytosis was thought to be involved in the bulk and nonselective uptake of extracellular fluid. However, it is becoming increasingly apparent that it is a morphologically and mechanistically distinct, highly regulated form of endocytosis. A growing number of viruses have been shown to exploit this pathway for entry, and investigations involving these viruses have significantly added to our understanding of the process [57]. While the current knowledge of the pathway is incomplete, morphological and regulatory characteristics that distinguish macropinocytosis from other endocytic processes have begun to emerge. Macropinocytosis occurs in outward extensions of the plasma membrane, termed ruffles, formed by localized assembly of actin filaments underneath the membrane [68]. Formation and recession of ruffles is a normal physiological process. A ruffle can fold back upon itself forming a cavity, which if followed by membrane fusion can trap fluid and solutes. Detachment from the cell membrane results in formation of a relatively large-sized vesicle called a macropinosome that ranges in size from 0.2 to 10 μm. This means they can take up large cargoes into the cell. As with CME, macropinocytosis occurs constitutively at low levels in all cells but can be induced following activation of receptors such as EGFR. This ligand-mediated receptor activation leads to a global increase in actin polymerization near the cell surface, inducing more ruffling of the membrane with a subsequent increase in macropinocytosis [69]. Little is known about trafficking following macropinosome formation; however, evidence points toward usage of a conventional endolysomal pathway involving early and late endosomes [69]. eGPs that target the macropinosome uptake pathway offer a major opportunity for cell delivery of cargoes. The overall large capacity of the macropinosome, which exceeds CME or CavME by an order of magnitude, offers an optimal path to deliver drugs into cells without requiring multiple uptake events. At the present, it is unclear how many viruses and which eGPs target this pathway but it is likely that large viruses like paramyxoviruses, filoviruses, and poxviruses require some form of macropinocytosis as they would be precluded from other uptake pathways based on size alone. The eGPs of these viruses are then likely to have specificity for receptors that are taken up by and induce macropinocytosis but will need to be confirmed experimentally. A few viruses use the so-called NCE pathway, which represents several distinct forms of endocytosis that are poorly characterized but do not involve

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clathrin or caveolin, and are not morphologically related to macropinocytosis. They require cholesterol, suggesting a role for lipid rafts or similar membrane microdomains, but involve conventional endolysosomal trafficking through EEs and avoid caveosomes. Lymphocytic choreomeningitis virus (LCMV), an old-world arenavirus, likely uses such a pathway [70] and is delivered into late endosomes from which membrane fusion is triggered. The pathways discussed above are those that are best characterized to date. The viruses and their eGPs that target each of these pathways have coevolved with the host to optimally deliver the virus core into the cell cytoplasm at a place conducive for replication. The endocytic uptake pathway is dictated by the eGP–receptor interaction. The ability of an eGP to target different endocytic pathways also enables tailoring of delivery route to the cargo size, chemical properties, and desired subcellular delivery site. Methods that can extract functional eGPs from virus particles or purify them from recombinant sources therefore offer an important source of material to target and deliver cargoes into these endocytic pathways and then to the cell cytoplasm. 20.4.5 Delivery Systems Using Virus Envelope Glycoproteins The coupling of eGPs to most virus cores is not dependent on specific amino acid sequences. This is best demonstrated in the production of virus pseudotypes, where the eGP from one virus can supplant the native eGP of another virus and deliver its core into cells. The core then adopts the receptor specificity, membrane fusion mechanism, and endocytic uptake pathway of the eGP donor virus. Many different pseudotyped viruses have been reported and span all families of the enveloped viruses [71–79]. This observation makes the delivery of artificial virus cores by eGPs realistic. The only common feature between the various virus cores that are good eGP acceptors is that the outer coat protein is often post-translationally modified with a fatty acid group (myristoylated) and is slightly acidic in amino acid composition [80]. In designing an artificial cargo (Figure 20.3), it is likely that a similar composition should be mimicked although the role of each of these factors for particle production and delivery still needs to be experimentally assessed. 20.4.6 Inactivated Viruses Most work in using eGPs as a delivery mechanism has used crude virus material, made by mechanical disruption of virus or by making virosomes from detergent solubilized eGPs. Disrupted viruses have proved effective in enhancing gene and protein delivery into cells. Sendai virus, also called hemagglutinating virus of Japan, is one example where UV-inactivated virus could enhance plasmid DNA transfection into cells [81]. Delivery efficiency was further enhanced by combination with biocompatible polymers such as protamine sulfate or cationized gelatin, which promote DNA and virus material association [82]. Such mixtures could also deliver plasmid DNA, oligonucleotide, or protein-coated magnetic particles. In further work, 26-nm magnetic

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Figure 20.3. Assembly and delivery of viruses and eGP-coated nanoparticles. At top: (1) Virus cores assemble from simple protein subunits called capsomers. Assembly is usually spontaneous and is enhanced by the presence of nucleic acid encoding the virus genome. Assembly can occur in the cytoplasm or in association with lipid membranes. (2) For enveloped viruses, the capsid interacts with the cell membrane in which the eGPs are present. (3) Capsid then buds through the membrane becoming coated with lipid and the embedded eGPs to form the infectious particle. (4) Infection of a new host can occur by multiple routes and targets specific cell types at the site of inoculation and delivers the virus into the cell cytoplasm by membrane fusion. The tropism of the virus is dictated by the eGP. The production of nanoparticles coated in eGPs mimics virus particle assembly. At bottom: (1) The nanoparticles are assembled from small molecule precursors and are made to polymerize in the presence of a compound of interest, for example, a drug or fluorescent marker. (2) eGPs extracted from viruses by mechanical disruption or detergent lysis are made to interact with the particle. (3) Coating occurs by electrostatic interaction and can be enhanced by mechanical means like sonication or extrusion. The coated nanoparticle now adopts the cell tropism of the eGP donor virus.

particles were coated with DNA and used to transfect cells. Mixing these particles with inactivated Sendai virus enhanced transfection of cells in culture and mouse liver [83]. This work demonstrates that eGPs can enhance delivery of material into cells. However, characterization of the process was complicated by the nature of the material.

20.5

VIROSOMES

Virosomes are composed of lipid vesicles (liposomes) with embedded eGPs. These were made from different virus eGPs including influenza virus, Sendai virus, and vesicular stomatits virus. Here the eGPs are purified away from other virus components either partially or to homogeneity. Virosome production usually involves dissolving the virus in detergent, removal of capsid protein through single or multiple purification steps, and then reconstitution

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of the eGPs into artificial lipid membranes after detergent removal. Virosomes were initially studied as potential drug carriers because they encapsulated drugs and protected them from degradation. However, without an underlying core, virosomes had disadvantages in low encapsulation efficiency, rapid leakage of water-soluble drugs in the blood, and poor storage stability [84]. Nevertheless, the methods used for virosome production are the same that would be used for production of eGP-lipid coated nanoparticles. 20.5.1 Preparation of Virus eGPs All viruses are relatively pure sources from which eGPs can be extracted. Virus particles are usually composed of three to four proteins of which the proteins making up the virus core (capsid and tegument/matrix) are predominant species and the eGPs are the next most common. Of these major proteins, only the eGPs contain hydrophobic transmembrane domains and are embedded in the viral lipid bilayer [2]. Nonionic detergents have been shown best for extraction of the eGPs while preserving receptor binding and membrane fusion functions. The detergents C12E8 and β-d-octylglucoside were used for producing virosomes of influenza A [85], Sendai [86], and vesicular stomatitis [87, 88] virus. Unfortunately, many eGPs do not survive treatment with these detergents. The newer short-chain phosphatidylcholine-based detergents promise better preservation of membrane protein function while giving good extraction from the membrane [89]. This is because they are chemically similar to lipids and extract proteins by displacing only the excess lipids from around proteins, leaving those closely associated with the protein in place [90]. The residual native lipids aid in shielding highly hydrophobic parts of the protein from the aqueous solvent. DCPC, a member of this detergent class, was shown effective in making influenza A virus virosomes [91]. It remains to be determined if these new detergents will prove useful for extraction of eGPs from other viruses.

20.6

RECONSTITUTION AND LOADING OF VIROSOMES

To make virosomes, detergent solubilized eGPs are reconsitituted into liposomes by removal of excess detergent in the presence of lipid. Removal of detergent is achieved through dialysis, gel filtration [92], direct extraction [93], or adsorption [85]. Each of these techniques relies on the concentration of detergent falling below the critical micelle concentration (CMC) of the detergent. At this point the equilibrium between eGP and lipid containing detergent micelles switches in favor of eGP–lipid complexes and liposomes spontaneously form. Detergent properties therefore need to be carefully chosen to have high CMC values (millimolar) and small micelle sizes. Each of the detergents discussed above have these properties. Nucleic acids and antigens are usually incorporated into virosomes by adsorption onto the surface or capture within the lumen of the liposome.

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Cationic lipids like DODAC and DOTAP (as used in lipid-based transfection reagents) aid capture of nucleic acids through electrostatic interactions. This method was used to enhance delivery of plasmid DNA using influenza A eGP virosomes. Although the plasmid was not encapsulated in the interior of the virosome, inducing membrane fusion by lowering the pH of the medium gave enhanced transfection efficiency of cells. Direct encapsulation of factors into virosomes has mostly relied on fluid-phase entrapment of solutes during formation of liposomes. For this, solutes were mixed with virus membrane extracts with or without adding lipids and detergent removed by adsorption to SM-2 Bio beads through hydrophobic interaction [94]. In order to improve encapsulation efficiency one group made siRNA-cationic lipid (DODAC) complexes first and then added the detergent-solubilized virus membranes prior to reconstitution [95]. This method gave a moderate enhancement of siRNA transfection efficiency. Another study showed that antisence oligodeoxy-nucleoside phosphorothioates were encapsulated into reconstituted influenza virosome containing cationic lipids by sonication [96]. An independent approach to compound delivery involved simply mixing the virosomes with liposomes containing the compound of interest (efficiently loaded by hydration of lipid and compound together). By briefly lowering the pH of the buffer, membrane fusion between the liposomes and virosomes was triggered and the two groups of vesicle merged. Importantly, some of the eGP survived to bind and deliver the cargo into target cells. DNA and peptides were demonstrated to be encapsulated in the virosomes using this method and could be delivered to cells [97, 98].

20.7

VIRUS ENVELOPE GLYCOPROTEIN-COATED POLYMERS

The entrapment of nanoscale particles within eGP–lipid films is a key goal in enhancing cargo capacity of virus-based delivery systems. However, methods need to be developed to drive spontaneous assembly of such particles and need to be tailored to the eGP proteins. Only a few articles have described approaches to achieve this goal. One group made hydrogel nanoparticles of crosslinked polyvinylpyrrolidone (PVPf-nanoparticle) containing fluoresceinated dextran (FITC-Dx) by polymerization. The PVPf-nanoparticles were mixed with detergent solublized Sendai virus membranes containing only the membrane fusion (F) protein. Virosomes were then formed by stepwise removal of detergent. This study showed specific delivery to the cytoplasm of human hepatoblastoma cells in vitro, a target of Sendai virus eGPs. Interestingly, compared with free FITC-Dx loaded virosomes, the eGPdelivered PVPf-nanoparticles mediated slower and more controlled release of the entrapped FITC dextran [99]. However, no particular effort was made to drive formation of the eGP-lipid+particle complex. Another approach, from our group, has been to coat fluorescent polystyrene nanoparticles (NPs) with Moloney murine leukemia virus eGP contain-

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ing membranes. For this, the nanoparticles were chosen to closely mimic a virus core in terms of size (100 nm), charge (carboxylated), and hydrophobicity (polystyrene base). Instead of using detergent to prepare virosomes, the virus membrane itself was removed from the virus particle by osmotic shock and sonication. This membrane was then coated over the nanoparticles by extrusion, a process where nanoparticles and virus membranes were forced through small pores (200 nm) in a polycarbonate filter. This process promoted interaction of particles and membranes that resulted in uniform coating of the particle with the virus membrane. Since fluorescent nanoparticles were used, it was possible to track them binding and entering cells. The coated nanoparticles mimicked native virus by only interacting with cells expressing the virus receptor and were taken up into early endosomes as indicated by colocalization with Rab5 (Figure 20.4).

Figure 20.4. eGP-coated nanoparticles enter cells through endocytosis. The set of images shows ecotropic murine leukemia virus eGP-coated nanoparticles (blue) interacting with cells expressing the virus receptor tagged with mStrawberry (red) and GFP tagged Rab5 (green, a marker of early endosomes). Each image is a plane from a z-stack and was taken using a Zeiss LSM 510 confocal microscope. The z-axis depth is indicated at upper left. A combined projection of all z-axis planes is shown at lower right. Arrowheads indicated nanoparticles interacting with receptor (violet), rab5 (cyan), or receptor and rab5 together (white). (See color insert.)

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Furthermore, we demonstrated cytosolic delivery of a cargo protein (βlactamase) that was coated on the nanoparticles prior to membrane coating. Cytosolic entry was demonstrated by measuring enzyme activity inside the cell [65]. We have also applied this approach to other viruses like vesicular stomatitis virus (unpublished work). While promising, problems still exist with this approach. The virus membranes are not free of other virus components (like capsid) and low level residual infectious virus is still present. Also, the contribution of particle composition to coating efficiency has not been clearly defined. These issues will need to be resolved before further advances can be made using this technique.

20.8 RECOMBINANT SOURCES OF VIRUS ENVELOPE GLYCOPROTEINS To overcome the problems associated with residual virus protein and nucleic acids present in virus extracted material, recombinant sources of eGPs are preferable. Unfortunately, most eGPs rely on a complex pattern of disulfide bonds, glycosylation, and post-translational processing, like protease cleavage, to be active. This rules out the use of bacterial expression systems and means that the more expensive eukaryotic systems need to be used. Some viruses (like hepatitis B virus) can generate noninfectious empty eGP containing particles in recombinant systems. Hepatitis B virus eGPs consist of three proteins called small (S), middle (M), and large (L) [100]. Hepatic cell specificity is determined by the L protein [101]. Kuroda et al. [102] showed that it was possible to produce particles of 23 nm in diameter in yeast expressing a recombinant form of the L protein. The use of the yeast system allowed the production of functional, glycosylated, and membrane-associated eGP proteins. Later work demonstrated that these particles could encapsulate and transfer green fluorescent protein or the fluorescent dye calcein to human hepatocytes in culture, or in a xenograft model [103, 104]. The use of engineered recombinant eGPs should make possible the selfassembly of eGP–nanoparticle complexes that will further enable eGP-mediated cargo delivery. Since the C terminus of the eGP is not essential for function, addition of small ligand binding domains could enable eGPs to bind to particles directly. Tags such as the His6 tag, which binds nickel ions, maltose binding protein, or epitope tags that can be bound by antibodies would be useful. While numerous examples of tagged eGPs exist, using them for selfassembly of a nanoparticle–eGP composite has not been reported. 20.8.1 Challenges in the Use of Virus Envelope Proteins for Cargo Delivery into Cells The coevolution of viruses with the host has meant that eGPs are highly adapted proteins that enable targeting of viruses to specific receptors, uptake

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through specific endosomal pathways, and finally delivery to the cell cytosplasm through membrane fusion. Each of these attributes are key components for an effective cell delivery mechanism for nanoscale particles. The major advantage of eGPs over other ligands is that each component is already present in one protein that can behave as a functional unit outside the native virus. This is particularly true for Class 1 and 3 eGPs, where each eGP trimer can act in isolation and does not interact with other trimer proteins. The fact that eGPs function as enzymes that promote lipid mixing and membrane fusion means that cargo size, which is constrained for nonenveloped viruses, is not a limiting factor in delivery into the cell cytosol. A major issue with using eGPs for delivery into the body is the complex interaction that takes place with the immune response. This is a similar situation to that seen for any virus-based therapy with neutralizing antibody responses being the most problematic. However, the need to evade a strong antiviral immune response has resulted in virus eGPs often being weak immunogens that illicit poor or short-lived humoral and cell-mediated immune responses [105]. The eGPs are glycosylated and sugar groups often mask the strongest epitopes, leaving only weak epitopes or easily varied regions exposed [106]. Importantly, these variable regions differ between related virus strains and so in the event of a strong immune response an eGP from a related strain with identical receptor specificity and entry mechanism could be substituted without having to extensively redesign the system. Similarly, chemical modification of eGPs with PEGs has been shown to shield vesicular stomatits virus eGP from serum inactivation [107]. The greatest question to be faced in using an eGP-based system for cargo delivery is which eGP should be chosen. This is a complex equation where factors of receptor usage, endosomal uptake pathway, fusion mechanism, source of material, and ability to reconstitute the eGP into the delivery vehicle must be balanced. From a practical standpoint the latter two factors are the most important. The ability of a detergent to preserve eGP function is often limiting. The ability to produce the eGP recombinantly is another limiting factor when working with virus is to be avoided. At the present, only a few eGPs have been extensively studied for effectiveness in particle delivery and the parameters for success have not been well defined but it is likely that many eGPs will be suitable as a delivery tool. It is apparent that eGPs enhance delivery of compounds into cells and this is likely due to the receptor binding, cell uptake, and membrane fusion functions of the eGP but the final destinations of the cargoes, where they enter the cell cytoplasm, is not well understood. Most of these studies rely on the development of specific reagents and an in-depth understanding of the cell biology of endocytosis, which is a developing field in itself. eGP-coated nanoparticles may also play an important role in the delineating endocytosis by use as probes. The ability to chemically functionalize cargoes will aid in this work. While additional future work is still needed to fully comprehend the potential of eGPs from different virus families, we expect that these proteins will play important roles

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in developing the next generation of drug delivery vehicles for targeted treatment of disease.

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of alpha-dystroglycan as a receptor for lymphocytic choriomeningitis virus and Lassa fever virus. Science 282: 2079–2081 (1998). Radoshitzky, S. R., Abraham, J., Spiropoulou, C. F., Kuhn, J. H., Nguyen, D., Li, W., Nagel, J., Schmidt, P. J., Nunberg, J. H., Andrews, N. C., Farzan, M., and Choe, H. Transferrin receptor 1 is a cellular receptor for New World haemorrhagic fever arenaviruses. Nature 446: 92–96 (2007). Burlone, M. E. and Budkowska, A. Hepatitis C virus cell entry: role of lipoproteins and cellular receptors. J. Gen. Virol. 90: 1055–1070 (2009). Spear, P. G. Herpes simplex virus: receptors and ligands for cell entry. Cell Microbiol. 6: 401–410 (2004). Pedersen, L., Johann, S. V., van Zeijl, M., Pedersen, F. S., and O’Hara, B. Chimeras of receptors for gibbon ape leukemia virus/feline leukemia virus B and amphotropic murine leukemia virus reveal different modes of receptor recognition by retrovirus. J. Virol. 69: 2401–2405 (1995). Kumlin, U., Olofsson, S., Dimock, K., and Arnberg, N. Sialic acid tissue distribution and influenza virus tropism. Influenza Other Respi. Viruses 2: 147–154 (2008). Fass, D., Davey, R. A., Hamson, C. A., Kim, P. S., Cunningham, J. M., and Berger, J. M. Structure of a murine leukemia virus receptor-binding glycoprotein at 2.0 angstrom resolution. Science 277: 1662–1666 (1997). Szakonyi, G., Klein, M. G., Hannan, J. P., Young, K. A., Ma, R. Z., Asokan, R., Holers, V. M., and Chen, X. S. Structure of the Epstein-Barr virus major envelope glycoprotein. Nat. Struct. Mol. Biol. 13: 996–1001 (2006). Davey, R. A., Zuo, Y., and Cunningham, J. M. Identification of a receptor-binding pocket on the envelope protein of friend murine leukemia virus. J. Virol. 73: 3758–3763 (1999). Adkins, H. B., Brojatsch, J., Naughton, J., Rolls, M. M., Pesola, J. M., and Young, J. A. Identification of a cellular receptor for subgroup E avian leukosis virus. Proc. Natl. Acad. Sci. U. S. A. 94: 11617–11622 (1997). Haynes, C., Erlwein, O., and Schnierle, B. S. Modified envelope glycoproteins to retarget retroviral vectors. Curr. Gene Ther. 3: 405–410 (2003). Verheije, M. H., Lamfers, M. L., Wurdinger, T., Grinwis, G. C., Gerritsen, W. R., van Beusechem, V. W., and Rottier, P. J. Coronavirus genetically redirected to the EGF receptor exhibits effective antitumor activity against a malignant glioblastoma. J. Virol. 83: 7507–7516 (2009). Valsesia-Wittmann, S., Drynda, A., Deleage, G., Aumailley, M., Heard, J. M., Danos, O., Verdier, G., and Cosset, F. L. Modifications in the binding domain of avian retrovirus envelope protein to redirect the host range of retroviral vectors. J. Virol. 68: 4609–4619 (1994). Chadwick, M. P., Morling, F. J., Cosset, F. L., and Russell, S. J. Modification of retroviral tropism by display of IGF-I. J. Mol. Biol. 285: 485–494 (1999). Aires da Silva, F., Costa, M. J., Corte-Real, S., and Goncalves, J. Cell type-specific targeting with sindbis pseudotyped lentiviral vectors displaying anti-CCR5 singlechain antibodies. Hum. Gene Ther. 16: 223–234 (2005). Katane, M., Takao, E., Kubo, Y., Fujita, R., and Amanuma, H. Factors affecting the direct targeting of murine leukemia virus vectors containing peptide ligands in the envelope protein. EMBO Rep. 3: 899–904 (2002).

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CHAPTER 21

Mitochondria-Targeted Pharmaceutical Nanocarriers VOLKMAR WEISSIG Department of Pharmaceutical Sciences, Midwestern University College of Pharmacy, Glendale, Arizona

GERARD G.M. D’SOUZA Department of Pharmaceutical Sciences, Massachusetts College of Pharmacy and Health Sciences, Boston, Massachusetts

21.1

INTRODUCTION

Mitochondrial research has made an enormous leap since mitochondrial DNA mutations were identified as the molecular cause for human diseases in 1988 [1–4] and the organelle’s decisive role in the complex pathway of apoptosis was identified during the 1990s [5–7]. Major molecular components of the mitochondrial machinery involved in the large number of mitochondrial functions have been identified and significant research efforts are increasingly ongoing. Effective therapies for diseases caused by malfunction of this organelle, however, still remain elusive. A major obstacle to manipulating mitochondrial dysfunction is the limited accessibility of mitochondria to direct physical, biochemical, and pharmacological intervention. Mitochondria form a complex tubular network, the shape of which is constantly changing based on permanently ongoing fission and fusion events [8, 9]. Furthermore, it has recently been demonstrated that the mitochondrial network is tightly intertwined with the microtubular meshwork, even ring-shaped mitochondria enclosing microtubules have been observed [10]. For the development of effective mitochondria-targeted therapies it is of high importance to devise strategies for overcoming all intracellular hurdles Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

385

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any molecule once taken up by a cell is facing before it can reach its intracellular target site. This chapter will critically review the state-of-the-art of mitochondria-targeted drug and DNA delivery systems currently under development.

21.2 INTRACELLULAR BARRIERS FOR MITOCHONDRIA-TARGETED DRUG AND DNA DELIVERY The interior of a cell is very different from any aqueous buffer solution, in which small drug molecules can freely diffuse and randomly interact with potential cosolutes. Any movement of low molecular weight compounds inside a cell is strongly affected by the cytoskeletal network, by dispersed organelles, and last but not least by the high amount of dissolved macromolecules. The concentrations of dissolved macromoleucles in the nucleoplasm and cytoplasm of living cells has been determined to be between 50 and 400 g/L [11, 12]. Subsequently, any transport process or diffusion event in such crowded solution has to be expected to differ significantly from those in buffer solutions and efforts are under way aimed at thoroughly understanding cellular material properties such as cytoplasmic viscosity, as recently reviewed by Matthias Weiss [13]. One of the major questions with respect to intracellular diffusion is how the intracellular environment affects the molecular mobility. Generally, intracellular diffusion has been characterized as hindered diffusion; that is, diffusion that is hindered by immobile barriers, by molecular crowding, and by binding interactions with immobile or mobile molecules [14, 15]. Two major groups of factors determine the fate of a low molecular weight drug molecule once it has entered a cell: first, the intracellular milieu and second, the nature of the drug itself. Both these groups of factors are tightly intertwined with each other. Fluid-phase viscosity of the cytoplasm, collisional interactions due to macromolecular crowding, and binding to intracellular components are all factors that prevent the free diffusion of solutes inside a cell [16, 17]. Their impact on the cytoplasmic diffusion of a low molecular weight compound is measurable and can be expressed as the translational diffusion coefficient. For example, using spot photobleaching, it was possible to measure the movement of DNA fragments of different sizes following microinjection into the cytoplasm of HeLa cells [16]. Not surprisingly, the rate of diffusion decreased with increasing size of the DNA. Fragments larger than 3 kb actually did not diffuse at all; the implications of this finding for the area of gene therapy are obvious. To what extent a low molecular weight drug molecule might interact or even bind to subcellular components, like membranes and cell organelles, depends on the physicochemical properties of the drug. Based on the intracellular distribution of a large variety of fluorophores, Richard Horobin has developed a quantitative structure–activity relationship (QSAR) model for predicting cellular uptake and intracellular distribution of

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TABLE 21.1 Localization

387

Molecular QSAR Parameters Determining Intracellular Drug

Parameter

Abbreviation

Charge Amphiphilic index Conjugated bond number Partition coefficient Acidity constant Molecular Weight Size of largest conjugated fragment Ratio largest conjugated fragment to conjugated bond number

Z AI CBN logP pKa MW LCF LCF / CBN

CBN<40 & logP<0

CBV>40 or logP (or AI) >8

CBN<40; 8>logP>0

Mitochondrion Cell membrane

Figure 21.1. Richard Horobin’s QSAR decision rules for predicting membrane permeation of low molecular weight compounds (for abbreviations see Table 21.1). This image was graciously provided by Richard W. Horobin.

low molecular weight compounds [18]. Table 21.1 summarizes the physicochemical parameters being used in Horobin’s QSAR approach. Figures 21.1 and 21.2 illustrate the applications of Horobin’s QSAR decision rules for predicting membrane permeation and intracellular distribution, respectively, of low molecular weight compounds. Figure 21.1 shows the parameters that the molecule needs to meet in order to cross the cell membrane, and Figure 21.2 depicts criteria for selected organelle specificities. Horobin’s QSAR approach was recently applied in depth to characterize potential common chemical features of molecules that are known to

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Lysosome

Endoplasmic reticulum 6>logP>0; Z>0; 6>AI>3.5

Z>0; 0>logP>−5 pKa<10; Z>0; 5>logP>0

For non-specific uptake into biomembranes: 8>logP>5

0>logP>−5; Z>0; pKa>10... Mitochondrion

DNA

Cell membrane

Figure 21.2. Richard Horobin’s QSAR decision rules for predicting the intracellular distribution of low molecular weight compounds (for abbreviations see Table 21.1). This image was graciously provided by Richard W. Horobin.

selectively accumulate at or inside mammalian mitochondria within living cells [19]. The authors generated from the literature a nonbiased sample of more than 100 so-called mitochondriotropic compounds and examined this data set using physicochemical classifications, quantitative structure–activity relationship (QSAR) models, and a Fick–Nernst–Planck physicochemical model. The ability of the latter two approaches to predict mitochondriotropic behavior was assessed, and comparisons were made between methods and with current assumptions. All approaches provided instructive pictures of the nature of mitochondriotropics. Most interestingly, however, although delocalized lipophilic cations have been regarded as the most common structural type of mitochondriotropic molecules [20–22], only a third were such. Much the same proportion were acids, potentially or actually anions. Many mitochondriotropics were electrically neutral compounds. From Table 21.2 it can be seen that selective mitochondrial accumulation involves electric potential, ion-trapping, and complex formation with cardiolipin, while nonspecific accumulation involves membrane partitioning and nonspecific access requires only low lipophilicity. Using QSAR and the Fick– Nernst approaches the authors were able to predict the mitochondriotropic behavior of more than 80% of the selected data set and allowed them to specify in some detail the physicochemistry of mitochondriotropic molecules. Overall, this approach is expected to facilitate guided syntheses and selection

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TABLE 21.2 Physicochemical Features Favoring Targeting or Accumulation of Xenobiotics in Mitochondria within Intact Cells

Accumulation & targeting processes

Selectivity

Cell line applicability

Physicochemical thumbnails, including parameter ranges (see also footnote)

Compounds with access to, but not necessarily accumulating in, mitochondria Membrane No selectivity All cell lines Lipophilic but not permeance superlipophilic: 8 > log P > 0 Compounds accumulating in mitochondria Reversible Low: uptake All cell lines Strongly but not partitioning into into all super-lipophilic: membranes biomembranes 8 > log P > 5 OR Strongly but not super-amphiphilic: 8 > AI > 5 Low All cell lines Weak acids with Ion-trapping of lipophilic less-ionized lipophilic weak species: Z ≤ 0; log acids within Pless ionized > 0; mitochondria pKa = 7 ± 3 [lysosomes & nuclei may also show uptake] High All cell lines Lipophilic to strongly Precipitation by lipophilic cations: cardiolipin & Z > 0; 5 > log P > 0 attraction by electrical potential Cells with higher Moderately hydrophilic Attraction by high High to moderately mitochondrial mitochondrial lipophilic cations: membrane membrane Z > 0; −2 > log P > 2 potential potential (Uptake by normal cell nuclei if log P < 0) Footnote: few pharmaceuticals have conjugated systems large enough, or are sufficiently amphiphilic or lipophilic, to be trapped in the plasmalemma. However, note that all above specifications have implied requirements for non-trapping by the plasmalemma, namely: AI < 8; CBN < 40; log < 8.

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of optimal mitochondriotropic structures. Moreover, the strategy developed by Horobin and Weissig [23] should generally be applicable to the future design of low molecular weight drugs aimed at acting on or inside all other cell organelles and has already proved useful for the modeling of cationic transfection lipids [23].

21.3

MITOCHONDRIA-TARGETED NANOCARRIERS

21.3.1 DQAsomes for Drug and DNA Delivery to Mitochondria Based on their resemblance to an old South American hunting weapon, a group of symmetric amphiphilic molecules, in which two hydrophilic residues are linked by a hydrophobic chain, have become widely known as “bolalipids.” Well-characterized bola-amphiphiles are archaebacterial lipids consisting of two glycerol backbones, which are connected by two hydrophobic chains. The self-assembly behavior of these archaeal lipids has thoroughly been investigated. It could be shown that such archaeal bola-lipids are able to self-associate into mechanically stable monolayer membranes [24]. While screening mitochondriotropic drugs potentially able to interfere with the mitochondrial DNA metabolism in Plasmodium falciparum [25], a serendipitous discovery revealed the tendency of dequalinium chloride, a bolaamphiphile (Figure 21.3A), to self-associate into colloidal structures. Electron microscopy (Figure 21.3C, right panel) as well as photon correlation spectroscopy confirmed the formation of particles with a diameter between about 70 and 700 nm. Freeze fracture electron microscopic images (Figure 21.3C, right panel C) showed both convex and concave fracture faces, thereby demonstrating the liposome-like aggregation of dequalinium. At the time of their discovery, these unusual vesicles were termed DQAsomes (pronounced dequasomes), that is, dequalinium-based liposome-like vesicles [26]. One striking structural difference between dequalinium and archaeal lipids, however, involves the number of hydrophobic chains between both polar head groups. In contrast to archaeal lipids, dequalinium possesses only one alkyl chain connecting both hydrophilic head groups. Therefore, theoretically, two different conformations within a self-assembled layer structure are imaginable. While the stretched conformation would give rise to the formation of a monolayer, assuming the horseshoe conformation would result in the formation of a bilayer (Figure 21.3B). Just like cationic liposomes, which are widely being explored as nucleartargeted nonviral transfection vectors [27], also DQAsomes were found to form complexes with DNA following mixing the positively charged vesicles with the negatively charged polynucleotide acid. This observation that DQAsomes, that is, cationic vesicles entirely composed of mitochondriotropic molecules, are able to efficiently bind DNA [26] and to protect the DNA from nuclease digestion [28], led to the first carrier-based strategy ever proposed

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(a) CH3 NH2

CH3

N – CH2 – CH2 – CH2 – CH2 – CH2 – CH2 – CH2 – CH2 – CH2 – CH2 – N

NH2

Cl

Cl

(b)

OR

(c)

Figure 21.3. (A) Chemical structure of dequalinium chloride, with overlaid gray shades indicating the bola-like nature of the molecule. (B) Theoretical possible conformations of dequalinium chloride, that is, stretched versus horseshoe conformation, leading to either a monolayer or a bilayer membranous structure following the process of selfassembly. (C) Left panel: Monte Carlo computer simulations demonstrate the possible self-assembly of dequalinium chloride into vesicles. The first image represents a transverse section of the second image, which represents a complete spherical vesicle. (D) Right panel: Electron micrographs of vesicles (“DQAsomes”) made from dequalinium chloride; from left to right: negatively stained transmission electron micrograph, rotary shadowed transmission electron micrograph, and freeze fracture scanning electron micrograph. (Reproduced with permission from Ref. 67).

for direct mitochondrial gene therapy [20–22]. This approach requires the transport of a DNA-mitochondrial leader sequence (MLS) peptide conjugate to mitochondria using DQAsomes, the liberation of this conjugate from the cationic vector upon contact with the mitochondrial outer membrane, followed by DNA uptake via the mitochondrial protein import machinery. DQAsome/DNA complexes (“DQAplexes”) were shown to release their DNA cargo only upon contact with mitochondrial membranes [29, 30]. Furthermore, utilizing a novel protocol for selectively staining free pDNA inside the cytosol, it was demonstrated that DQAsomes selectively deliver pDNA to and release the pDNA exclusively at the site of mitochondria in living mammalian cells [31]. Finally, utilizing confocal fluorescence

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microscopy and using DNA–MLS peptide conjugates, it was shown that DQAsomes not only deliver oligonucleotides but are also able to deliver plasmid-sized DNA into mitochondria within living mammalian cells [32]. In addition to having been established as the first mitochondria-specific DNA delivery system, DQAsomes have also been explored as a mitochondriatargeted nanocarrier system for small drug molecules, in particular, for anticancer drugs known to trigger apoptosis via direct action on mitochondria [33, 34]. For these studies, paclitaxel has been chosen as a model drug. It is a wellknown antitumor agent used in the treatment of several cancers. Clinically, the therapeutic potential of paclitaxel is limited due to a very narrow span between the maximal tolerated dose and intolerable toxic level. In addition, its poor aqueous solubility requires the use of emulsion formulations containing Cremophor EL, a surfactant of considerable toxicity in itself [35]. Paclitaxel is largely believed to exert its action by stabilizing the microtubules of cells, but the precise mechanism of paclitaxel-induced apoptosis remains unclear [36, 37]. It has also become evident that paclitaxel has other targets inside the cell, most notable of which is the mitochondrial network. For example, it was shown that paclitaxel could act directly on isolated mitochondria from human neuroblastoma cells to induce the permeability transition pore (PTP)-dependent release of mitochondrial cytochrome c [38]. Very interestingly from the perspective of subcellular drug targeting, a 24-h delay has been observed between the paclitaxel-triggered release of cytochrome c in intact cells versus a cell-free system [39]. This delay has been attributed to the existence of several drug targets inside the cell, making only a subset of the drug molecules available for mitochondria [39]. Consequently, in order to increase the apoptotic activity of paclitaxel, its subcellular (i.e., mitochondrial) bioavailability would have to be increased. Hence paclitaxel appears to be a molecule whose action may be significantly improved by specific subcellular delivery to the mitochondrion. This premise has been perfectly confirmed. First, it was shown that DQAsomal encapsulation of paclitaxel changes the subcellular distribution of a labeled derivative of the drug. Confocal fluorescence microscopic images demonstrate that, in contrast to the free paclitaxel, the DQAsomal encapsulated drug at least partially colocalizes with mitochondria [34]. Subsequently, the metabolic cytotoxicity was compared with the apoptotic activity. As anticipated, it was found that encapsulation of the drug into DQAsomes did not significantly alter the dosedependent metabolic toxicity of paclitaxel, but it clearly improved the proapoptotic action [34]. Figure 21.4A shows fluorescence micrographs of the nuclear morphology of colo205 colon cancer cells treated with different preparations. The quantitative analysis of apoptotic nuclei detected by this assay shown in Figure 21.4B revealed a significant increase (p < 0.05) in apoptotic nuclei in cells treated with DQAsomal paclitaxel over free drug or over paclitaxel mixed with empty DQAsomes. The results of the nuclear morphology assay were further confirmed by a DNA fragmentation assay, which suggested that 10-nM DQAsomal encapsu-

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c

(B) b

d

% Apoptotic Nuclei

a

393

25 20 15 10 5 0 20 nm DQA

20 nM DQA + 10 20 nm DQA w/10 nm paclitaxel nm encapsulated (mixed) paclitaxel

Figure 21.4. Nuclear morphology assay for determining apoptosis. (A) Representative fluorescence micrographs showing nuclear morphology of Hoechst-stained, colo205 colon cancer cells treated with (a) negative control, (b) 20 nM DQAsomes, (c) 10 nM free paclitaxel + 20 nM DQAsomes, and (d) DQAsomal-encapsulated paclitaxel (10 nM free paclitaxel + 20 nM DQAsomes) for 20 h. (B) Quantitative estimation of apoptotic nuclei based on 400 cells counted for each group. Inset: Representative image showing normal (open arrow) and apoptotic nuclei (solid arrow). (Reproduced with permission from Ref 34).

lated paclitaxel was comparable to 50-nM free paclitaxel in inducing DNA fragmentation characteristic of apoptosis: that is, DQAsomal encapsulation increased the apoptotic activity of paclitaxel approximately five times [34]. Three years earlier the same group had tested paclitaxel-loaded DQAsomes for their ability to inhibit the growth of human colon cancer tumors in nude mice and their data suggested that encapsulation of paclitaxel in DQAsomes leads to improved in vivo efficacy [33]. Taken together, these data demonstrate that DQAsomes are able to change the intracellular distribution of drugs and that the specific targeting of a drug to an appropriate subcellular target can potentiate the desired drug activity. The antitumor efficiency of DQAsomal encapsulated paclitaxel was most recently further enhanced by modifying the DQAsomal surface with folic acid (FA) [40]. The folate receptor is a folate high-affinity membrane binding protein, which is overexpressed in a large variety of human tumors [41–43]. FA conjugates have been shown to be internalized in a tumor cell specific manner by receptor-mediated endocytosis, resulting in an increased toxicity of the corresponding drug [44–46]. Vaidya and colleagues [40] have incubated EDC activated FA-PEG-COOH with preformed paclitaxel-loaded DQAsomes at different molar ratios resulting in DQAsomes in which up to 5.3% of all dequalinium molecules were conjugated to FA-PEG-COOH. Cell cytotoxicity studies using folate receptor expressing HeLa cells demonstrated that folic acid conjugated DQAsomes possess better antitumor activity as compared to plain paclitaxel-loaded DQAsomes, folic acid conjugated paclitaxel-loaded liposomes, and the free drug. The authors concluded from their data that folic

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acid conjugated DQAsomes delivered the drug not only to the cytosol but also to mitochondria, whereas folic acid conjugated liposomes delivered the drug into the cytosol only [40]. 21.3.2 Mitochondria-Targeted Polyethylene Imine A mitochondrial leader peptide (MLP), derived from the cytosolically expressed but mitochondrially localized ornithin transcarbamylase, was recently used to render polyethylene imine (PEI) mitochondriotropic [47]. PEI had been developed in the mid-1990s as a versatile vector for gene and oligonucleotide transfer into cells in culture and in vivo [48, 49]. Lee et al. [47] conjugated the mitochondrial leader peptide to PEI via a disulfide bond and confirmed the complex formation of PEI-MLP with DNA by a gel retardation assay. In vitro delivery tests into living cells performed with rhodamin-labeled DNA demonstrated that PEI-MLP/DNA complexes were localized at mitochondrial sites in contrast to controls carried out with PEI/DNA complexes lacking MLP. The author’s data suggest that PEI-MLP can deliver DNA to the mitochondrial sites and may be useful for the development of “direct mitochondrial gene therapy,” a strategy for the cure of mitochondrial DNA diseases originally proposed by Seibel et al. [50] and by Weissig and Torchilin [20–22] as an alternative to allotopic expression. 21.3.3 Mitochondriotropic Liposomes To render liposomes mitochondria specific, Weissig’s group has modified the liposomal surface with triphenyl phosphonium (TPP) cations [51]. Methyltriphenylphosphonium (MTPP) cations were shown 40 years ago to be rapidly taken up by mitochondria in living cells [52] and Murphy’s group has extensively explored the marked mitochondriotropism of MTPP for the delivery of biological active molecules to and into mitochondria [53–56]. Weissig and co-workers replaced the methyl group in MTPP with a stearyl residue in order to moor mitochondriotropic and hydrophilic TPP cations to the liposomal surface. The intracellular distribution of such TPP surface-coated liposomes (stearyl triphenylphosphonium liposomes, STPP liposomes) was analyzed by confocal fluorescence microscopy using rhodamine-labeled phosphatidylethanolamine (Rh-PE) as a liposomal fluorescence marker [57]. For control, liposomes with incorporated positively charged yet nonmitochondriotropic dioleoyl trimethyl ammonium propane (DOTAP) were used. Both preparations (i.e., the control liposomes and STPP liposomes) displayed the same positive surface charge. Not unexpectedly, both types of surface-linked cations were found to enhance the association of liposomes with cells in an almost identical manner, as was shown by flow cytometric analysis. However, the intracellular distributions of STPP liposomes and DOTAP liposomes were quite different. While confocal fluorescence microscopy revealed an almost complete association of STPP liposomes with mitochondria, DOTAP liposomes displayed significantly less

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mitochondrial association. The authors concluded from their study that a positive surface charge of liposomes enhances cell association per se, while for directing liposomes to specific intracellular target sites an appropriate organelle-specific ligand on the surface of liposomes is essential. Following the demonstration that STPP liposomes deliver a lipophilic fluorescence marker (i.e., Rh-PE) almost exclusively to mitochondria, the authors then investigated whether increased drug concentrations at the site of mitochondria would also result in increased drug efficiency. To this end, ceramide was incorporated into STPP liposomes. Ceramide mediates a large variety of intracellular biological responses to extracellular stimuli [58–60]. It has also been found that anticancer drugs can cause an increase of ceramide concentrations in the vicinity of mitochondria [61] and it has been hypothesized that such increase enables the formation of ceramide channels in the mitochondrial outer membrane [62, 63]. Boddapati et al. [57] incorporated ceramide into STPP liposomes and assessed the apoptotic activity of such formulations in vitro and in vivo. It was found that at the low drug concentration used only the mitochondria-targeted ceramide was able to elicit a strong apoptotic response. Subsequent in vivo studies were carried out with polyethylene glycol bearing STPP liposomes. Interestingly, it was revealed that the cationic TPP ligand did not significantly change the biodistribution of STPP-PEG5000 liposomes in comparison to conventional charge-neutral PEG liposomes. Even more important, the tumor accumulation of STPP-PEG liposomes was almost identical to their noncharged counterparts. Mice were inoculated subcutaneously with mouse mammary carcinoma cells and upon formation of palpable tumors, the animals were divided into three groups for treatment with either buffer or empty or ceramide-loaded STPP-PEG liposomes. Figure 21.5A shows the tumor volumes measured during the tumor growth inhibition study. In both control groups, half of the animals developed necrotic morbidity after 12 days and had to be euthanized. Strikingly, none of the animals treated with mitochondria-targeted ceramide showed any morbidity even after 18 days. Statistical analysis of tumor growth rate at the 12-day time point (n = 6) showed that the treatment with ceramide in STPP-PEG liposomes significantly inhibited tumor growth rate compared to sham treatment (Figure 21.5B). It should be emphasized that in Weissig’s study a significant tumor growth inhibition was achieved with ceramide doses as low as 6 mg/kg. For comparison, administering ceramide formulated in conventional (i.e., nontargeted) liposomes, Stover et al. [64] were able to observe tumor growth reduction only at ceramide doses at least six times that high (i.e., equal or above 36 mg/kg). 21.3.4 The MITO-Porter Concept: Mitochondrial Delivery Via Membrane Fusion A unique liposome-based nanocarrier system able to deliver its cargo to the mitochondrial interior via membrane fusion has been developed by Harashima’s group [65, 66]. Using fluorescence resonance energy transfer (FRET) analysis,

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(A)

Tumor Volume (mm3)

1000 800 600 400 200

Da y − Da 2 y Da 0 y Da 2 y Da 4 y Da 6 y Da 8 y Da 10 y Da 12 y Da 14 y Da 16 y 18

0

Tumor Volume Increase (mm3/day)

(B)

70 60 50 40 30 *

20 10 0 Sham

Empty Nanocarrier

Ceramide in Nanocarrier

Figure 21.5. Tumor growth inhibition. (A) Tumor volume (mm3) measured over time period of treatment in BALB/c mice bearing murine 4T1 mammary carcinoma tumors (n = 6); after treatment with buffer (䊏), empty STPP nanocarrier (䉱), and ceramide in STPP nanocarrier (䉬). (B) Tumor growth (in mm3/day) at day 12. (n = 6, error bars denote standard deviation); (asterisk indicates a Student t test p-value of <0.05). (Reproduced with permission from Ref. 57).

the authors screened liposomes composed of a variety of lipids for their fusion activity with isolated rat liver mitochondria and found liposomes with surfacelinked octaarginine and containing either phosphatidic acid or dioleoyl phosphatidylethanolamine to display the highest fusion potential. Following encapsulation of green fluorescence protein (GFP) into such fusogenic vesicles termed “MITO-Porter,” their ability to selectively deliver their cargo into mitochondria in living cells was demonstrated based on confocal fluorescence microscopy.

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CONCLUSION

Following the introduction of DQAsomes as the prototype of mitochondriatargeted pharmaceutical nanocarriers 10 years ago, a variety of nanotechnology-based strategies for the delivery of macromolecules and nucleic acids to and into mitochondria in living mammalian cells have been designed. It is hoped that the ongoing merger of nanoscience with mitochondrial research and medicine will eventually lead to new and efficient therapies for the large number of human diseases caused by mitochondrial disorders.

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65. Yamada, Y., Akita, H., Kamiya, H., Kogure, K., Yamamoto, T., Shinohara, Y., Yamashita, K., Kobayashi, H., Kikuchi, H., and Harashima, H. MITO-Porter: a liposome-based carrier system for delivery of macromolecules into mitochondria via membrane fusion. Biochim. Biophys. Acta 1778: 423–432 (2008). 66. Yamada, Y. and Harashima, H. Mitochondrial drug delivery systems for macromolecule and their therapeutic application to mitochondrial diseases. Adv. Drug Deliv. Rev. 60: 1439–1462 (2008). 67. Cheng, S. M., Boddapati, S. V., D’Souza, G. M., and Weissig, V. DQAsomes as mitochondria-targeted nano-carriers for anticancer drugs. In: M. Amiji, ed. Nanotechnology for Cancer Therapeutics. CRC/Taylor & Francis Group, Boca Raton, FL, 2007, pp. 787–802.

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CHAPTER 22

Cell-Penetrating Peptides for Cytosolic Delivery of Biomacromolecules CAMILLA FOGED, XIAONA JING, and HANNE MOERCK NIELSEN Department of Pharmaceutics and Analytical Chemistry, Faculty of Pharmaceutical Sciences, University of Copenhagen, Copenhagen, Denmark

22.1

INTRODUCTION

A major unsolved problem in drug delivery is how to transport therapeutic biomacromolecules (peptides, proteins, and nucleic acids) across membrane barriers. The issue becomes even more pertinent in the future as many new drug candidates are biopharmaceuticals that have to pass one or more membrane lipid double layers to reach their pharmacological target site that is often localized in the cytosol of cells. The challenge to be overcome is that the size and the hydrophilic nature of biomacromolecules are unfavorable for their permeation of biological membranes, resulting in very limited bioavailability. The identification and exploitation of cell-penetrating peptides (CPPs) during the last two decades of research represents an elegant and promising approach to overcome membrane barriers that was inspired by the discovery of naturally occurring CPPs like Tat and penetratin [1–5]. Such CPPs constitute an essential part of shuttling proteins (e.g., from viruses) and appeared to mediate the entry of these proteins into cells. Tat and penetratin, as well as many additional identified CPPs, have been shown to be capable of transporting other macromolecules such as proteins through membranes into cells [6]. These peptides were first called Trojan horses or protein transduction domains (PTDs), but were also named CPPs after a reevaluation of the internalization mechanism, which showed that endocytosis, supplementary to transduction, is a major internalization pathway [7].

Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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Multiple studies have consolidated the high efficiency of CPP-mediated cargo delivery in vitro as well as in vivo; the first in vivo study being the demonstration by Schwarze et al. [8] that fusion of the protein β-galactosidase to the Tat domain led to ubiquitous delivery of the protein upon intraperitoneal administration in mice. CPPs therefore seem to hold great promise as delivery agents for biomacromolecules with the intracellular targets. In this chapter we focus on the use of CPPs as carriers for biomacromolecules with the pharmacological target site of action in the cytosol of cells. In addition, we describe the combination of CPPs with particulate drug delivery systems for cytosolic delivery of therapeutic biomacromolecules. 22.1.1 Cell-Penetrating Peptides as Carriers for Cargos CPPs constitute a very heterogeneous and large group of peptides that is often subdivided into different classes and according to various criteria. Major classes are the polycationic or arginine-rich peptides (e.g., Tat, penetratin, R8 (octaarginine) [9]) and the amphipatic peptides (e.g., transportan [10]). In addition, CPPs may be classified as naturally occurring fragments of proteins (Tat, penetratin), as entirely synthetic peptides (e.g., MPG [11], R8, Pep-1 [12], YTA2 [13]), or as chimeras such as transportan. In general, all CPPs are short peptide sequences of usually 7–16 amino acids. There seems to be no or little homology between primary and secondary structure among CPP classes, but a common feature is that CPPs are often rich in basic residues (arginine (R), lysine (K)) that can interact with negatively charged cell surface-bound molecules [14]. CPPs are associated with remarkably low toxicity at effective concentrations and have therefore been applied to induce translocation of a wide range of cargo types into cells. In general, two strategies can be used for association of cargo molecules to CPPs: (1) covalent attachment of carrier and cargo via various types of linkers [see Ref. 15 for review] and (2) noncovalent self-assembly whereby aggregates or nanoparticles are formed [see Ref. 16 for review]. In most cases reported with successful outcome, the CPP and the cargo are chemically conjugated, but even complexation of the CPP and the cargo or simple coadministration seem in some cases to be efficient. Covalent conjugation is experimentally demanding with time-consuming synthesis and, importantly, purification of conjugates from unconjugated entities. In some cases, physical self-assembly is possible and especially amphipathic peptides have the ability to selfassemble [16]. Several studies have shown membrane translocating effect of noncovalent complexes of CPP and cargo [12, 17, 18]. No matter which type of conjugation strategy is used, in general, it should be stressed that a careful characterization of the complexes is required in order to fully deduce their biological effect. Both when administering conjugated and complexed CPP– cargo construct, these constructs are well defined only at the time of dosing, but after proof-of-concept testing, it will be of utmost value for future systematic design of optimal constructs to correlate the stability and physical characteristics of the conjugated or complexed structures to their effect.

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INTRODUCTION

405

Efficacy assays using end point determination of cargo activity or biological/ pharmacological effect supplementing cellular or tissue localization are necessary to support the vast number of mechanistic assays carried out with fluorophore-labeled CPPs. It is becoming more and more evident that not only the type of CPP [19] but also the type of cargo affects the delivery efficacy of the CPP, which is reflected in the effect on cellular uptake [20] and toxicity [21, 22]. The cargo can affect the cell-penetrating properties of CPPs in various ways such as alteration of the internalization mechanism, changed intracellular trafficking, and modified membrane destabilizing properties, which in the worst case renders the CPP–cargo complexes inactive [reviewed in Ref. 23]. An example is a study performed with transportan 10 that suggests that direct membrane effects may cause membrane translocation of transportan 10 alone and of smaller complexes, whereas these effects do not contribute for larger cargos [24]. Below we describe how CPPs have been applied for transmembrane transport of peptides, proteins, antigens, nucleic acids, and colloidal carrier cargos, and how association of cargos to CPPs affects the cell-penetrating properties of the CPPs. 22.1.1.1 Peptides and Proteins As previously mentioned, the interest in CPPs as delivery vehicles for peptide and protein delivery kicked off with the significant distribution of Tat-fused β-galactosidase to most organs, even the brain, in mice upon intraperitonal injection [8]. Following that, a number of different therapeutic peptides and proteins have been applied together with CPPs, some of which are compiled in Table 22.1 [for a more comprehensive overview, see Refs. 18 and 25–29]. Tat is the most widely tested CPP in combination with an active cargo, but also penetratin, the synthetic polyarginines, and other CPPs have been used for peptide and protein delivery. 22.1.1.2 Antigens For vaccines, delivery of antigen to the cytosol of professional antigen-presenting cells (APCs) might be advantageous for induction of cytotoxic T lymphocyte (CTL) responses, which are essential for elimination of intracellular pathogens. CPPs have therefore been suggested as transporters for antigens to the cytosol of APCs. CTLs exert their most important effector mechanism by antigen-mediated killing of infected cells [30]. Dendritic cells (DCs) are professional APCs that capture antigens, process them into peptide fragments, and present them on major histocompatibility complex (MHC) classes I and II to specific T cells. The majority of the MHC class I-restricted peptides are generated in the cytosol by proteolytic degradation of the proteins into peptides catalyzed by the proteasome [31]. The peptide fragments are transported into the lumen of the endoplasmic reticulum (ER) via the transporters associated with antigen processing (TAP), and the peptide fragments are then loaded onto newly synthesized MHC class I molecules [32] and transported to the cell surface for presentation to CD8+ T cells, which subsequently differentiate into CTLs. Additionally, CD8+ DCs have a unique ability to deliver exogenous antigens to the MHC class I

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406

TABLE 22.1 a

CPP

Examples of Delivery of Peptides and Proteins by CPPs Peptide/Proteina

Application

Assay

Result Evaluation (Effect/Transduction)

Reference 8 20 139 39 40

Tat Tat Tat Tat Tat

β-Galactosidase GFP Insulin Ovalbumin TRP2

Conjugation Conjugation Conjugation Conjugation Conjugation

In vivo, mouse In vitro In vitro In vivo, mouse In vivo, mouse

Tat

TRP2-derived antigenic peptide 5 Amino acid peptide

Conjugation

In vivo, mouse

Conjugation

In vitro

Uptake and effect Uptake Transepithelial transport CD8+ T-cell responses CD8+ T-cell-dependent tumor prevention and treatment CD4+ and CD8+ T-cell-dependent tumor prevention and treatment Uptake

3–4 Amino acid peptides Carbonic anhydrase

Conjugation

In vitro

Uptake

Conjugation

In vitro

Uptake, effect

Insulin

Coadministration

In situ, rat

Transepithelial transport and effect

138

Cyclosporin A Cell cycle inhibitory protein 27kip1 β-Galactosidase GFP β-Galactosidase Streptavidin

Conjugation Complexation

In vivo, mouse In vitro

Effect Effect

141 18

Complexation Coadministration

In vitro In vitro

Uptake and effect Uptake and effect

Tat Penetratin Transportan MAP R7 R7W R8 R16 R6 R8 R10 R7 Pep-1 Pep-1 Pep-1 YTA2 pVEC 7/30/2010 2:19:43 PM

a

GFP, green fluorescent protein; Rn, n-arginine; TRP2, tyrosine-related protein 2.

140 19

136 9

12 17

INTRODUCTION

407

(cross-presentation) pathway. Several models for cross-presentation have been proposed [33]. One model is based on uptake of antigens by phagocytosis and proceeds via vacuolar pathways, whereby peptides derived from exogenous antigens bind to MHC class I molecules within the post-Golgi vacuolar compartment or on the cell surface [34]. Another model suggests that whole organisms or antigens escape from vacuolar compartments after phagocytosis into the cytosol, undergo cytosolic processing, and subsequently bind to MHC class I molecules in the ER [35–38]. Several studies have confirmed that CPPs enable antigen-specific stimulation of CD8+ T-cell responses (Table 22.1). Conjugation of Tat to ovalbumin resulted in efficient ex vivo loading of DCs and induction of potent antigenspecific CD8+ T-cell responses upon inoculation in mice, in contrast to DCs loaded with ovalbumin alone [39]. In a tumor model, vaccination with a tumor antigen fused to Tat induced both tumor prevention and tumor treatment in a CD8+ T-cell-dependent way [40]. Vaccination with Tat-antigen-transduced DCs induced a robust CTL response and durable antitumor immunity. These studies suggested that Tat might facilitate cross-presentation by enhancing endosomal escape of antigen to the cytosol. However, in accordance with more recent studies of the transduction mechanism of CPPs, a study showed that CPPs rather induce cross-presentation in vitro via more efficient endosomal uptake without endosomal escape, since the arginine-containing peptide PolyR induced enhanced cross-presentation of PolyR-conjugated, class I-restricted peptides in DCs [41]. CPPs have also been suggested as vaccine excipients that could enhance transport of exogenous material to protease-containing MHC II peptide loading compartments and prolong antigen presentation [42]. MHC class II molecules acquire peptide cargo that is generated by proteolytic degradation in endosomal compartments. The precursor proteins of these peptides include exogenous material and endogenous components that access the endosomes by autophagy. Vaccination with an antigenic peptide conjugated to a CPP resulted in efficient CD4+ T-cell-dependent tumor regression in a murine model of peptide-based immunotherapy, as compared to the unconjugated antigenic peptide [42]. Enhanced uptake and transport to endosomal compartments might lead to improved antigen processing and MHC class II-peptide loading sustaining the presentation of antigen [42, 43]. 22.1.1.3 Nucleic Acids CPPs are studied with increasing interest as carriers for nucleic acid-based macromolecules like oligonucleotides and plasmid DNA with target sites of action in the cell cytosol or the nucleus for modulation of gene expression [reviewed in Refs 44 and 45]. Oligonucleotide-cargos successfully transported by CPPs include peptide nucleic acid (PNA) and short interfering RNA (siRNA). However, transport efficiency appears to be dependent on the charge of the nucleic acid cargo and the type of CPP association. Table 22.2 shows examples of CPP-mediated nucleic acid delivery referred to in the sections below.

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TABLE 22.2 408

CPPa

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Transportan Penetratin Penetratin Tat Tat Penetratin Transportan R6-penetratin MPG MPG EB1 Transportan Penetratin Tat Penetratin R6-penetratin R3-penetratin R9-penetratin K8 (R-Ahx-R)4 Tat R7 KLA Transportan R7–9 Tat Penetratin NLS Transportan10 Tat Tat-phage Branched Tat a

Examples of Delivery of Nucleic Acids by CPPs Nucleic Acida

Application

Assay

Result Evaluation (Effect/Transduction)

Reference

siRNA

Conjugation

In vitro

Uptake and effect

47

siRNA siRNA siRNA

Conjugation Conjugation Conjugation

In vitro In vitro In vitro

Uptake and effect Uptake and effect Uptake and effect

48 49 50

PNA

Conjugation

In vitro

Effect

51

siRNA siRNA siRNA PNA

Complexation Complexation Complexation Conjugation

In vitro In vitro In vitro In vivo, rat

Uptake and effect Uptake and effect Uptake and effect Uptake and effect

52 53 54 59

PNA

Conjugation

In vitro

Uptake and effect

60

PNA

Conjugation

In vitro

Uptake and effect

57

PNA

Conjugation

In vitro

Uptake and effect

62

Plasmid Plasmid Plasmid Plasmid

Complexation Complexation Conjugation Complexation

In vitro In vitro In vitro In vitro

Uptake and effect Uptake and effect Uptake and effect Uptake and effect

64 65 66 67

K8, octalysine; NLS, nuclear localization signal; PNA, peptide nucleic acid; Rn, n-arginine; siRNA, small interfering RNA.

INTRODUCTION

409

There are very few published examples of successful delivery of negatively charged siRNA into cells by covalent attachment of CPPs [reviewed in Ref. 46]. Muratovska and Eccles [47] conjugated transportan and penetratin with N-terminal cysteins via disulfide-bonds to the 5′ end of the sense strand of siRNA, and observed a reduction of luciferase and GFP (green fluorescent protein) expression in reporter cells by the conjugates. However, a nonspecific “diamide”-assisted oxidation method of a mixture of free thiol-containing siRNA and peptide was applied, and the complexes were added to cells without further purification and characterization, leaving an open question of which chemical entity is causing the silencing effect. Furthermore, no data on toxicity was reported. Davidson et al. [48] linked penetratin to the 5′ end of the sense strand of siRNA, and the construct was delivered efficiently into primary neuronal cells without further purification and caused a downregulation of protein production after 6 hours that preceded mRNA degradation. In a third study, Chiu et al. [49] conjugated Tat to siRNA through a stable thiomaleimide linkage and purified the conjugate by polyacrylamide gel electrophoresis. The conjugate was able to silence gene expression of a reporter gene (GFP) as well as an endogenous gene in vitro. However, none of these three reported studies have been reproduced in the literature, which indicates the complexity of the conjugation procedure and suggests that careful characterization of conjugates is important for a reproducible biological effect. More solid approaches to couple, purify, and characterize covalent complexes of CPPs and oligonucleotides/siRNA have been reported by the group of M. Gait [50, 51]. Disulfide-linked conjugates of CPPs were first synthesized via C-terminal cystein residues to 5′ -thiol-containing sense strand oligonucleotides and then annealed to the antisense siRNA strand containing a 3′ fluorescein residue. Formamide was used as denaturing agent during conjugation and ion exchange HPLC was applied for purification to prevent aggregation and complete separation of excess peptide. This method seems to circumvent the difficulties in coupling a polyanion to a polycation and in annealing sense strand conjugates containing highly cationic peptides to the corresponding negatively charged antisense strand. Some inhibition of P38α MAP kinase mRNA expression was observed with penetratin and Tat peptide–siRNA conjugates. However, very high levels of conjugates were needed (micromolar concentrations), increasing the risk of nonspecific off-target effects. This shows that the translocation efficiency of the conjugates is inhibited by the multiple anionic charges present on the phosphate backbone, and that a single peptide thus appears insufficiently powerful to trigger enough release from vesicular or endosomal compartments into the cytosol. However, combination of covalent oligonucleotide–CPP complexes and other endosome disrupting strategies might enhance endosomal escape, as described in the forthcoming section. Noncovalent complexation of siRNA with certain types of CPPs has been shown to be a feasible delivery method at certain molar and charge ratios, where compact, cationic particles are formed. These peptides include MPG

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[52, 53] and the penetratin analog EB1 [54]. Unfortunately, biological studies are either often done under serum-free conditions, and thus do not address the important issue of serum stability of the complexes, and/or utilize poorly characterized complexes making interpretation of results difficult. Optimizing such noncovalent complexes toward serum stability might be a promising approach for siRNA delivery. Recent studies suggest that chimeras of the CPP R9 and targeting ligands like a rabies virus glycoprotein (RVG)-derived peptide [55] or a CD7-specific single chain antibody [56] can be used as delivery vectors for siRNA in vivo, enabling transvascular delivery of siRNA to the brain and suppression of human immunodeficient virus (HIV) infection in mice, respectively. However, the R9 moiety seems in this case merely to function as a binding site for siRNA, although structural characteristics of the complexes as well as the uptake mechanism are poorly understood. Covalent conjugates of CPPs and the electrically neutral nucleic acid analogs, such as PNA and phosphoroamidate morpholino oligomers (PMOs) that can hybridize to RNA targets with high specificity [57, 58], appear much more promising than the anionic oligonucleotides. Conjugates of PNA and CPPs such as transportan and R6-penetratin have been shown to promote PNA translocation and splice correction of target genes [51, 59–62]. Some peptide sequences with optimal chemical and structural features might thus be capable of escaping from endosomes in the absence of other endosomolytic agents [58, 63] but additional endosomal membrane destabilizing strategies are required for most CPPs [57]. Finally, few studies have reported successful delivery of plasmid DNA with CPPs. There seems to be little or no effect with single peptides [64, 65]. However, branched CPPs or a combination of CPPs with other delivery agents might be useful for plasmid transfection [64, 66, 67]. 22.1.2 Particulate Drug Delivery Systems and CPPs CPPs have also been widely tested for transmembrane delivery of a range of larger-sized/colloidal cargos such as liposomes [68, 69], polymeric nanoparticles [70, 71], polymeric micelles [72], magnetic nanoparticles [73], and quantum dots [74] that have been reviewed recently [26, 75, 76]. Selected examples are shown in Table 22.3. An inherited problem of applying CPPs for delivery purposes, independently of cargo size, is their lack of cell specificity. Most CPPs will associate to membranes of all cell types nonspecifically due to global expression of heparan sulfate proteoglycans [8]. The resulting ubiquitous membrane translocation may be beneficial under some circumstances, but may be undesired in other cases where targeting of specific cell types or tissues is needed. A way to introduce specificity is to associate CPPs to other excipients of drug delivery systems, for example, colloidal carriers, which by themselves can deliver a drug in a specific way, either by passive targeting via the enhanced permeability and retention (EPR) effect upon systemic

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TABLE 22.3 a

CPP

Transportan10 Tat Tat Tat Tat Tat Tat Tat Tat Pep-1 Rn Tat Tat a

Examples of Delivery of Colloidal Carriers by CPPs Colloidal Carriera

Application

Assay

Result Evaluation (Effect/Transduction)

Reference

PEI Phage Liposome Liposome MEND PEG-PEI Polymeric micelles Magnetic nanoparticles Quantum dots

Complexation Conjugation Conjugation Conjugation Conjugation Complexation Conjugation Conjugation Conjugation

In vitro In vitro In vitro In vitro In vitro In vivo, mice In vitro In vivo, mice In vitro

Uptake and effect Uptake and effect Uptake and effect Uptake and effect Uptake and effect Uptake and effect Uptake Uptake Uptake

64 66 68 69 70 71 72 73 74

Liposomes Liposomes Micelles

Conjugation Conjugation

In vivo In vitro

Uptake and effect Uptake

78 79

MEND, multifunctional envelope-type nanodevice; PEG, poly(ethylene glycol); PEI, polyethylene imine; Rn, n-arginine.

411

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CELL-PENETRATING PEPTIDES FOR CYTOSOLIC DELIVERY OF BIOMACROMOLECULES

administration or by active targeting approaches [26]. In this section we discuss recently published approaches that combine the membrane translocating property of CPPs with the specificity of colloidal drug carrier systems in “smart”, stimuli-responsive nanocarriers. pH-responsive, Tat-modified long-circulating liposomes and micelles have been designed by Torchilin et al. [77–79]. These long-circulating poly(ethylene glycol) (PEG)-coated liposomes and micelles are targeted actively to a specific cell type or organ via the attachment of an antibody to PEG-phosphatidylethanolamine (PE) at their surface. PEG-PE is degradable at low pH due to a pH-sensitive bond between PEG and PE. The carriers are furthermore modified with Tat-short PEG-PE derivatives. At normal physiological conditions the longer PEG chains shield or hide Tat on the shorter PEG chains. However, at lower pH, as in tumors, the longer PEG chains are cleaved from the complexes, whereby Tat is exposed, enhancing cellular internalization. In another recent study, polymeric micelles consisting of a hydrophobic core of polylactic acid and a hydrophilic shell of PEG-conjugated Tat were complexed with a pH-sensitive diblock copolymer of poly(methacryloyl sulfadimethoxine) (PSD) and PEG (PSD-b-PEG) [72]. The anionic PSD is complexed with the cationic Tat whereby PSD-b-PEG shields the Tat-containing micelles. At low pH the complexes are deshielded, which exposes Tat leading to translocation into cells in vitro. The same principle has been tested in murine tumor models with doxorubicin-loaded micelles that inhibited tumor growth significantly [80]. Future studies will show if these “smart” long-circulating drug delivery systems are able to introduce specificity in vivo by passive targeting via the EPR effect in tumor tissue (eventually combined with active targeting strategies), where the CPP is exposed and translocates the drug-containing cargo into cells. The above referred studies apply highly complex carrier systems that require thorough characterization to fully understand how individual components of the carrier system contribute to delivery in vivo.

22.2 INTERNALIZATION MECHANISMS AND INTRACELLULAR TRAFFICKING The puzzling feature of how CPPs, which are often highly cationic and thereby charged molecules, can pass nonpolar, lipophilic cell membranes has fueled an intense interest in elucidating their mechanisms of internalization. Despite extensive studies with a number of different sequences, there seems to be no consensus regarding one specific mechanism as a preferred route of CPP uptake [81]. It is becoming increasingly evident that CPP uptake pathways may vary depending on the physicochemical properties of the CPP and the cargo they deliver, the specific cell types, and the experimental conditions [82]. The interaction between CPPs and the membrane is the very first step of the internalization process (Figure 22.1). Nonamphipathic or polycationic

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INTERNALIZATION MECHANISMS AND INTRACELLULAR TRAFFICKING

CPP

413

CPP-cargo covalent attachment noncovalent complexation CPP modified nanoparticle – +

Clathrin- & Caveolin-independent endocytosis Inverted micelle Cytosol

Direct penetration

Golgi apparatus

Early endosome

Caveolin-mediated endocytosis

Late endosome

Clathrin-mediated endocytosis ER Nucleus

Lysosome

Macropinocytosis

Figure 22.1. Mechanisms of CPP-assisted cytoplasmic drug delivery. (See color insert.)

CPPs are short and highly positively charged like Tat [4] and R8 [9]. They have a high affinity toward anionic glycosaminoglycans (GAGs) on cell membranes, only destabilize membranes at high micromolar concentrations, exhibit low toxicity, and may subsequently induce translocation mainly through endocytosis rather than direct translocation. Primary amphipathic CPPs such as MPG [11] and transportan [10] behave similarly to antimicrobial peptides. They can bind to and insert into both neutral and anionic lipid membranes mainly via hydrophobic interactions and might form transmembrane pores and exhibit toxic properties even at low concentrations. Secondary amphipathic CPPs such as penetratin [83] and pVEC [84] undergo conformational changes upon interaction with lipids (or GAGs) converting them into amphipathic molecules that can be inserted superficially into the anionic lipid bilayers. Besides the different physicochemical properties of various CPPs, other factors such as (1) the characteristics of the cargo, (2) the linkage between CPP and the cargo, (3) the cell type, and (4) the peptide to cell ratio and other experimental conditions also possibly lead to multiple entry mechanisms [85]. The direct penetration or reverse micelle mechanisms might be responsible for the translocation of a CPP itself. However, when CPPs are linked to a large cargo, endocytosis would be favored as diffusion of ion pair complexes would be expected to decrease with size. For example, CPPs alone or CPP-conjugated small molecules penetrate cell membranes via electrostatic interactions and hydrogen bonding [86], while Tat-mediated intracellular delivery of large cargoes proceeds via macropinocytosis [87].

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22.2.1 Endocytosis Mechanisms Endocytosis is a regulated, energy-dependent cellular uptake process by which cells internalize macromolecules and particles into transport vesicles derived from the plasma membrane (Figure 22.1). There are three different types of endocytosis: (1) phagocytosis for uptake of large particles, (2) pinocytosis for uptake of fluids and solutes, and (3) receptor-mediated endocytosis [88]. Phagocytosis is limited to APCs such as macrophages and DCs, while pinocytosis is ubiquitous in all cells. Pinocytosis can be further classified into four different pathways: clathrin-mediated, caveolin-mediated, clathrin- and caveolin-independent endocytosis, and macropinocytosis. All of these pathways are related to the internalization of different CPPs with various cargoes [81, 89]. One of the differences between these pathways is the size of endocytic vesicles formed. For instance, macropinosomes normally form vesicles larger than 1 μm in diameter, clathrin-coated vesicles are about 120 nm, vesicles in clathrin- and caveolin-independent endocytosis are approximately 90 nm, and caveolae pits are around 60 nm [88]. Cargoes are transferred to early endosomes where they can be either recycled and exocytosed or transported to organelles like lysosomes, golgi, and mitochondria. The trafficking pathways include receptor-dependent and receptor-independent pathways. Receptor-dependent endocytosis involves assistor proteins. The vesicle formation for the caveolin-1 or clathrin pathway is similar and requires the action of dynamin-2, but the intracellular fates are distinct. In clathrin-dependent uptake, mature endosomes subsequently fuse with lysosomal vesicles resulting in enzymatic destruction of cargoes, whereas in clathrin-independent internalization, early endosomes pass to late endosomes of higher acidity where molecules are hydrolyzed [90]. 22.2.1.1 Endosomal Escape The fate of CPPs and their cargoes after uptake is highly influenced by the acidic pH and digestive enzymes present in the endosomes. Endosomal escape is the major rate-limiting step for CPPmediated delivery of cargoes to the cytoplasm. Agents that can enhance endosomal release include chloroquine, high concentrations of Ca2+, sucrose, and photosensitizers. Besides, fusogenic lipids, membrane-disruptive peptides, and membrane-disruptive polymers have also been exploited for efficient drug delivery systems. These strategies are summarized in Table 22.4. In the discussion of CPPs for intracytosolic drug delivery, membrane-disruptive peptides or fusogenic peptides are of particular interest. Fusogenic peptides usually exploit the natural acidification process occurring in the endocytic pathway. One example is the fusogenic peptide HA2 from influenza virus, which contains several acid residues [91]. The protonation of the acid residues is triggered by the decrease in endosomal pH, and results in a conformational change to an amphipathic helix, which can destabilize the membrane [92, 93]. Some intracellular drug delivery systems have adapted this mechanism that viruses use for the endosomal escape by coupling with

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TABLE 22.4

Endosome-Disrupting Strategies to Improve Cytosolic Delivery

Endosomal Disruptor Chloroquine

Ca2+ Sucrose Photosensitizer Fusogenic lipid

Cargo Cre protein PNA PNA

Coincubation Coincubation Coincubation

DNA PNA PNA PNA Plasmid

Coincubation Coincubation Coincubation Coincubation Liposome incorporation (DOPE/CHEMS) Liposome incorporation (DOPE/CHEMS) Liposome incorporation (DOPE/CHEMS/EPC) Liposome incorporation (EPC/Chol/DOTAP/ PE-HZ-PEG)

siRNA Ovalbumin Plasmid

Fusogenic peptides 10 Histidine-2 cysteine4(3)E HA2

Endosomal Disruptor’s Linkage to Cargoa

CPPs and Their Linkage to Cargo TAT fusion protein TAT, R8 conjugates TAT, penetratin, transportan conjugates TAT-NLS-dendrimer complex TAT, Rnconjugates K8 conjugates TAT, R7, KLA conjugates R8 Liposome incorporation IRQ Liposome incorporation R8 Liposome incorporation TAT Liposome incorporation

Plasmid

TAT-10 histidine-2 cysteine-fusion peptide/DNA complexes

Plasmid P53 protein

DNA-cationic peptide(46)–4(3)E complex HA2-P53-R11 fusion protein

Assay

Reference

In vitro In vitro In vitro

87 142 143

In vitro In vitro In vitro In vitro In vitro In vivo In vitro

144 142 145 57 146, 147

In In In In

vitro vivo vitro vivo

149

In vitro In vivo In vitro In vitro

150

148

78

151 152

415

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416 TABLE 22.4

Continued

Endosomal Disruptor HA2-penetratin

Cargo PNA

EB1 siRNA GALA Plasmid KALA Plasmid Membrane-disruptive polymers 2-kDa PEI Oligonucleotide 25-kDa PEI 25-kDa PEI Dendrimer a

Plasmid Plasmid Oligonucleotide

Endosomal Disruptor’s Linkage to Cargoa

CPPs and Their Linkage to Cargo

Assay

Reference

HA2-penetratin fusion peptide linked to PNA via disulfide bridge Penetratin analog EB1 complexed with siRNA Transferrin-PEG-GALA with condensed DNA nanoparticle PEI-DNA-KALA particle

In vitro

143

In vitro In vitro In vitro

54 153 154

PEG-PEI copolymers surface modified with TAT and then complexed to oligonucleotides R11-PEI conjugate and mixed with plasmid DNA DNA/Antp/PEI coacervation nanoparticles CPP-oligonucleotide conjugates and complexed with dendrimer

In vitro In vivo In vitro In vitro In vitro

155 156 157 158

CHEMS, cholesteryl hemisuccinate; DOPE, dioleylphysphatidylethanolamine; DOTMA, N-[1-(2; 3-dioleyloxy)propyl]-N;N;N-triethylammonium; DSPE, distearoylphosphatidyl-ethanolamine; EPC, egg phosphatidylcholine; GFP, green fluorescent protein; HZ, hydrazone linkage; PA, phosphatidic acid; PE, phosphatidylethanolamine; PEG, poly(ethylene glycol); PEI, polyethylene imine; PNA, peptide nucleic acid; Rn, n-arginine; SM, sphingomyelin.

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INTERNALIZATION MECHANISMS AND INTRACELLULAR TRAFFICKING

417

synthetic fusogenic peptides, such as GALA and KALA [94]. Poly-l-histidine can also trigger pH-responsive fusion, which correlates with the protonation of imidazole groups of histidine. The pKa of the imidazole group of histidine is around 6.0, and thus the imidazole group is protonated in acidic endosomes. Conventional CPPs can be modified with certain amino acids at different positions to obtain endosomolytic properties. For example, the endosomolytic CPP EB1 has been designed with histidine replacement of specific amino acids in the penetratin sequence to adopt an alpha helix conformational change upon protonation in the endosomes. EB1 is more effective in delivering biologically active siRNA (i.e., it obtains better gene silencing effect than its parent peptide) [54]. 22.2.2 Transduction Although the reevaluation of the internalization mechanisms of CPPs emphasized endocytosis as the main uptake mechanism for CPPs carrying large cargoes, in several other cases, arginine-rich CPPs alone or with small cargoes may still undergo direct membrane translocation, also called transduction. This is observed when endocytosis is inhibited (i.e., at 4 °C and in the presence of endocytosis inhibitors) [95], in the presence of counteranions such as pyrene butyrate, which may lead to charge neutralization [96], or at higher peptide concentration [97]. The exact mechanisms are not clear at present, but important determinants for transduction are suggested to be (1) accumulation of peptides on the cell surface to a certain extent, (2) transmembrane potentials, and (3) hydrogen bonding [98, 99]. Two hypothetical models exist for the transduction mechanism: direct membrane penetration [100] and the formation of inverted micelles [101]. 22.2.3 Intracellular Trafficking Up until today, endocytic uptake is the most accepted pathway for many CPPs and the cargoes they carry, despite some exceptions, as discussed above. However, very little is known regarding the intracellular fate of CPPs after endocytic uptake. The concerns that the CPP–cargo complexes might be trapped in the endosomes have led researchers to speculate on several strategies for endosomal escape (Table 22.4). Recently, an alternative possibility for cytosolic CPP release has been proposed based on the mechanism of toxin uptake [102]. After endocytosis, toxins such as Shiga toxin enter the cytosol by retrograde transport to the Golgi apparatus and then the ER. This is particularly attractive for cytosolic drug delivery since it is desirable to bypass the acidic pH and the hydrolytic environment of the lysosomes. Whether this retrograde trafficking pathway hypothesis is applicable for conventional CPPs is still doubtful given the contradictory results for Tat and R8 by Jones et al. [103]. However, the targeting properties of A/B type toxins such as Shiga toxin reside in their B subunit [104], which

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has the potential to be coupled into the current drug delivery system to obtain a nontoxic retrograde drug delivery. To make the picture more complex, certain CPPs are suggested to stimulate intracellular signaling cascades involving selected Rho-family GTPases that induce their uptake and regulate their endocytic trafficking [105].

22.3 DESIGN OF CELL-PENETRATING PEPTIDES FOR INTRACELLULAR CARGO DELIVERY In order to direct delivery of therapeutic cargoes to the cytosol, many of the same approaches have been taken as for optimization of CPP to facilitate cell entry. In general, the positive charge is essential for initial interactions with membranes [14], so increasing the number of R-amino acids in particular [106–108] and controlling the spacing between the guanidinum groups in the CPP lead to increased membrane interaction. Once taken up in the cell, limited endosomal escape hampers the cytosolic delivery of the cargo carried by the CPP. For increasing delivery specifically to the cytosol, various synthetic approaches have been investigated. The general recognition that increased proteolytic stability is important for uptake and processing in cells [109] has fueled studies of single residue exchanges in the CPP in order to increase the proteolytic stability. d-Amino acid exchange for the more labile l-amino acid is often investigated [110]. Other approaches to stabilize the CPP structure are to alter the chirality and structure of these CPPs, resulting in differences in cellular localization as well as systemic distribution. Thus novel CPPs, some rich in guanidino moieties, have been prepared. These include oligomers of β-amino acids (β-peptide), carbamates, peptoids [111], branched-chain CPPs [112], dendrimers, sugarbased and biphenyl-based CPPs, and amphiphilic proline/silaproline-based peptides [98]. However, single residues in a naturally derived CPP may be critical for transfer to biological membranes. As an example, exchanging tryptophan with phenylalanine decreased the cellular uptake of penetratin, even though the interaction with a lipid vesicle membrane was not changed [113]. Letoha et al. [114] showed the same dependency and further revealed that penetratin distributed to the cytosol upon cellular uptake. It has, however, been shown that substitutions of hydrophobic residues in penetratin did not alter the cytosolic and nucleic distribution pattern of the peptide [115]. Incorporating unnatural substituents such as the aminohexanoic acid (Ahx) into an arginine sequence (R-Ahx-R)n and analogs hereof has proved efficient in increasing the endosomal escape of the construct [116]. Also, terminal conjugation of six arginine residues to penetratin greatly enhanced the efficiency of the conjugated PNA without use of endosomolytic agents [60]. Although the presence and localization of positive charges seems to be the main important criteria for efficient CPP uptake and translocation, the

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hydrophobic content of the peptide appears to influence internalization [110]. An example is the complexation of R side chains in a CPP to aromatic moities of pyrene butyrate, thereby inducing cytosolic delivery of the CPP-assisted cargo [96]. Derivatization of CPPs with acyl chains increases the overall hydrophobicity, and the SAP sequence branched with fatty acids (C10 and C14) in itself increased cellular uptake and cytosolic localization several-fold [110]. Also, C18-penetratin significantly increased the splice correction readout as compared to penetratin upon administration in a complex with an oligonucleotide [117]. The splice correction was, however, further enhanced in the presence of chloroquine, indicating that cytosolic delivery was not complete. Similarly, the cholesteryl-conjugated R9 [118] and also silaproline derivatization of the SAP peptide increased the hydrophobic content as well as the cellular uptake significantly [110]. Several conditions may be encountered in the endosomal escape of CPP with cargo; first, the decrease in pH in endosomes as opposed to extracellular fluid, changes in membrane potential, and the up-concentration of the CPP construct in the endosomes may all play roles in the mechanism for endosomal escape [119]. Design of pH-triggerable CPP (e.g., histidine-rich) fusogenic peptide structures is a direct approach to increase endosomal release due to alterations in the CPP charge and structure (examples were given previously). Thus polymorphism of the CPP in different media and thereby modulation of the exposed hydrophobic part to the membrane has recently been discussed to be an important activity parameter for many CPPs [120]. Although for CPPs, such as penetratin, which is believed to enter cells partly via nonendocytic and non-energy-dependent mechanisms [115], the alpha-helical structure is not mandatory for internalization and intracellular processing [121]. Some studies suggest that the coupling of a CPP to a high molecular weight cargo forces it into a macropinocytotic route of entry, regardless of whether this is the case for the CPP alone [15, 87]. As an example, R8-modified liposomes [122] are thus believed to be internalized by macropinocytosis and to a large extent avoid lysosomal degradation due to leakiness of the macropinosomes or specific escape herefrom. This correlated with the observations by Torchilin [123], who observed that Tat-grafted liposomes migrated in the same manner. Other studies reported that CPPs linked to avidin were taken up by both clathrin-dependent and clathrin-independent endocytosis [124] and that different CPPs conjugated to PNA utilized different internalization routes corresponding to their amphipathic nature, but not to the biological readout [125]. Regardless of the route of delivery, the efficiency of the CPP on drug delivery may depend on whether direct conjugation, complexation, or incorporation into a delivery vehicle is applied. Sufficient stability of the delivery system and controllable cleavability are critical, thus size and type of linker such as disulfide or amide bonds for covalent linkage to amino acid-based drugs, and disulfide and thioester bonds for nucleic acid-based drugs, can be optimized.

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CELL-PENETRATING PEPTIDES FOR CYTOSOLIC DELIVERY OF BIOMACROMOLECULES

METHODOLOGIES

Since the cellular uptake of CPPs is in general considered to be independent of cell type, and since there is no consensus regarding uptake mechanisms and pathways of different types of CPPs, this should continuously be studied in great detail in order to optimize subsequent design of novel CPPs. It is also important to recognize that attachment of or complexation with a cargo, or incorporation of the CPP into a delivery vehicle, presumably will alter the cellular uptake pathway. Studies should preferably be carried out with a given cargo, and as for other delivery systems, in vivo studies with intravenous injections and subsequent tissue preparation are optimal, and have also been performed to some extent with CPP delivery systems [126]. Alternatively, in vitro studies using well-established cell culture models characterized with regard to a desired biological readout such as splice correction or silencing (Table 22.2) can be very useful. Studies on functional activity of the cargo molecule may be supplemented with live-cell imaging and mechanistic studies using lipid bilayers, thereby providing additional insight into the uptake and processing mechanisms [127]. In vitro cell culture models are widely used to study the route of cellular uptake and processing, mostly by use of microscopy, often applying fluorophores attached to the CPP terminal end and tracking by observation of a cellular localization and colocalization with macropinosome fluid-phase markers or other fluorescent early or late endosomal markers [102]. Interpretation of patterns of distribution visualized by confocal microscopy and quantified by flow cytometry [128] is thus done under the assumptions that (1) the fluorophore does not alter the properties of the CPP, (2) it is still linked to the construct of interest, (3) the signal is not altered due to, for example, pH sensitivity or quenching, and (4) surface-bound CPP is not calculated as internalized CPP [127]. Fischer et al. [129] observed pronounced differences in uptake for fluorophore-labeled CPPs compared to nonlabeled CPPs. Alternatively, biotinylated CPPs can be used with success [115, 130], but, in that case, permeabilization and fixation of the samples are necessary. It is important to recognize that live cells should in general be used since fixation increases the risk of artifacts (such as localization of label in the nucleus) [81]. Electron micrographs can provide much more detailed resolution of the cell and can also be used to visualize endosomes and intracellular organelles. But, as with the fluorescent techniques, use of transmission electron microscopy would require that the CPP or CPP delivery system carry a contrast label or could be labeled after the experimental incubation, as described by PalmApergi et al. [131]. Inhibitors of endocytosis can elucidate the mechanism of cellular entry and to some extent also intracellular processing. It must, however, not be neglected that these inhibitors are not completely specific and might affect, for example, intracellular pH levels [81]. As stated previously, chemical endosomolytic agents (such as chloroquine, high concentrations of Ca2+, or sucrose) or pho-

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toactivation can be applied in vitro in order to gain insight into the degree of inefficiency depending on endosome entrapment. Alternatively, lysing and subsequent fractionation of the cells could be used to quantitatively determine the intracellular localization of the CPP at different time points. However, for sensitive detection this technique would also require the use of labeled CPP (e.g., with fluorophore or biotin). In addition to in vitro cell culture studies and in vivo studies, the use of model vesicle membranes applying biophysical techniques is important for elucidating mechanisms of membrane association, interaction, and perturbation, and additionally for comparative studies [132, 133]. For these types of studies, detection can to some extent be done spectroscopically on a native tryptophan residue or a residue introduced terminally in the CPP, and thus avoiding the use of larger and more hydrophobic fluorophores. These studies are clearly justified, assuming that the binding and interaction with plasma membranes is the crucial first step in delivery using CPPs irrespective of the subsequent uptake pathway.

22.5

SIDE EFFECTS

Even though CPPs are associated with little or negligible toxicity at effective concentrations, some tendencies are briefly addressed here. Cellular toxicity is most often evaluated by assays to determine pertubation of the plasma membrane with consequent leakage from the cell interior [17, 19, 134], by assays determining intracellular enzymatic [21, 135] or ATP activity [17], or by nucleus staining [136]. In vitro, the cytotoxicity of most CPPs is in or above the micromolar range with some CPPs even being nontoxic at 100 μM [21]. In some cases, the effect seems to be cell type dependent [21, 137] and also dependent on the type of CPP [21], although some of the differences are minor. In another study, a similar degree of cytotoxicity was observed for Pep-1 in four different cell lines [12] and with Tat and penetratin in two cell lines [135]. When differentiating between the toxicity of different CPPs, a correlation between the degree of amphipathicity and membrane leakage was observed for a range of CPPs [47], and often Tat shows to be less nontoxic than, for example, transportan, penetratin, and polyarginines [21, 134]. Recent attention has been paid to the fact that the toxicity of CPP–cargo constructs might be different from the CPP, and the toxicity of Tat has been shown to increase when a peptide cargo (or even rhodamine) is conjugated to CPPs with a dependency on the specific cargo [21]. Similar findings were recently reported by Cardozo et al. [22], who concluded that the degree of cytotoxicity depended on the length of the cargo peptide as well as the dose applied to the cells. It has also been reported that the increased efficiency of YTA2 complexed with a protein as compared to the corresponding Tat– protein complex correlated with a higher level of toxicity [17]. Upon in vivo

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administration resulting in successful delivery of insulin, no release of intracellular lactate dehydrogenase was observed from intestinal tissue after application of Tat [138] corresponding to the observations of lack of toxicity in the initial in vivo studies with Tat–cargo constructs.

22.6

CONCLUSION

As evident from the present chapter, a significant number of experiments have to date been published concerning the screening of different types of CPPs with regard to cellular uptake and target validation with the overall conclusion that CPPs are promising carriers for a vast number of cargoes such as biomacromolecules and nanoparticles with cytosolic sites of action. Although some attention has recently been drawn to understanding the mechanisms of internalization and intracellular targeting, mainly pursued by the use of fluorophore-conjugated CPPs, many issues still have to be resolved. A major bottleneck for CPP-mediated delivery of biomacromolecules is endosomal escape, which is poorly understood. Also, in the rapidly evolving area of nanomedicine it is presently not clear which internalization mechanisms are responsible for the uptake of the CPPs associated with, for example, a biomacromolecular or colloidal cargo. We anticipate that current research will lead to delivery systems consisting of a CPP conjugated or associated to small molecule therapeutics, nucleic acids, peptide, or protein drugs or formulated in a more complex delivery system. Such CPP-based systems will also constitute valuable tools for research, diagnostics, and imaging.

22.7

LIST OF ABBREVIATIONS

Ahx APC CHEMS CPP CTL DC DOPE DOTMA DSPE EPC EPR ER GAGs GFP HIV HZ Kn

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Aminohexanoic acid Antigen-presenting cell Cholesteryl hemisuccinate Cell-penetrating peptide Cytotoxic T lymphocyte Dendritic cell Dioleylphysphatidylethanolamine N-[1-(2, 3-dioleyloxy)propyl]-N,N,N-triethylammonium Distearoylphosphatidyl-ethanolamine Egg phosphatidylcholine Enhanced permeability and retention Endoplasmic reticulum Glycosaminoglycans Green fluorescent protein Human immunodeficiency virus Hydrazone n-Lysine

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REFERENCES

MEND MHC NLS PA PE PEG PEI PMO PNA PSD PTD Rn RVG siRNA SM TAP TRP

423

Multifunctional envelope-type nanodevice Major histocompatibility complex Nuclear localization signal Phosphatidic acid Phosphatidylethanolamine Poly(ethylene glycol) Polyethylene imine Phosphoroamidate morpholino oligomer Peptide nucleic acid Poly(methacryloyl sulfadimethoxine) Protein transduction domain n-Arginine Rabies virus glycoprotein Short interfering RNA Sphingomyelin Transporters associated with antigen processing Tyrosine-related protein

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CHAPTER 23

Therapeutic Nano-object Delivery to Subdomains of Cardiac Myocytes VALERIY LUKYANENKO Department of Medicine, Johns Hopkins School of Medicine, Baltimore, Maryland

23.1

INTRODUCTION

Heart failure develops when the amount of blood pumped from the heart is inadequate to meet the metabolic demands of the body [1]. Heart failure is a syndrome with many different well-described causes, including myocardial infarction, pressure overload, volume overload, viral myocarditis, toxic cardiomyopathy, and mutations in genes encoding for sarcomeric or cytoskeletal proteins [1]. At least half of the causes lead to relatively slow deterioration in heart functioning, during which some cells are more damaged than others. The creation of nanocarriers to deliver specific markers, drugs, and therapeutic genes should make it possible to recognize sick cardiac cells and cure them individually. In other words, targeted delivery of analytic probes and therapeutic agents to cardiac myocytes and cellular compartments could significantly increase the efficiency of both diagnostic and treatment protocols for heart failure and have a significant impact on heart failure research. On the way to their cardiac targets, nano-objects must penetrate multiple barriers including capillary walls, the sarcolemma, and intracellular barriers, which are unique to the structure of cardiac myocytes [2]. The size and coating of the carriers must allow them to reach the target and go through intracellular barriers. The particles have to be coated with a shell that includes compounds preventing aggregation and nonspecific binding of nano-objects, and specific molecules that (1) improve recognition of the target cell, (2) promote binding to the cell membrane, (3) facilitate transportation through the cellular

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membrane, (4) defend the cargo compound from low lysosomal pH, and (5) have a “key” for entering the intracellular structure. For nanotechnology involved in the therapy of vascular problems resulting in heart failure please see the reviews by Kong and Goldschmidt-Clermont [3] and Brewster et al. [4].

23.2

STRUCTURE OF VENTRICULAR CELL

Cardiac myocytes are rod-shaped cells ∼100 μm in length and 20–30 μm thick (Figure 23.1). The characteristic cardiac stripes seen with light microscopy are a combination of extracellular Z-grooves and cytoskeletal Z-lines (Figure 23.1A). The sarcolemma of ventricular cells (biggest cardiac myocytes) has multiple invaginations, called T tubules, longitudinally connected to the transverse-axial tubular system (TATS) [5, 6]. Figure 23.1B shows a complicated network of the TATS tubules penetrating the entire thickness of the ventricular myocyte. Z-lines separate the myocytes into structurally similar sarcomeres. A longitudinal section presented in Figure 23.1C shows the main structures of the sarcomere: myofilaments, the myofibrillar and intermyofibrillar mitochondria, network and junctional SR, and transverse tubules (T tubules) of the TATS. Thus the diffusion pathways for nano-objects inside a ventricular myocyte can be divided into two groups: (1) the tubules of the TATS and (2) sarcoplasmic aqueous diffusion pathways [7]. The former is not a true intracellular system because of its multiple contacts with the extracellular space through T tubules. The diameters of the TATS tubules usually fall within the range of 200–400 nm [6]. However, they are filled with the glycocalyx [6], so the real clearance of the TATS pathway for nano-objects is ∼10 nm [7]. Therefore under normal conditions about 50% of the sarcolemma in cardiac ventricular myocytes is not available for transport of nano-objects >10 nm. The ultrastructure of cardiac myocytes suggests the existence of intracellular barriers significantly restricting the diffusion of nano-objects toward functionally important cellular compartments and structures such as the nucleus, mitochondria, and the space between the junctional sarcoplasmic reticulum (jSR) and the T tubules (so-called junctional cleft) (Figure 23.1C). Figure 23.2A shows the primary targets and possible barriers and pathways for nano-objects within a cardiac myocyte. Targeting the nucleus and mitochondria is important because of their role in protein expression and cell energetics. The TATS sarcolemma contains numerous transporting structures and receptors crucial for cardiac function. The structural organization of the space around the jSR is a matter of special interest because of its direct involvement in excitation–contraction coupling [8]. Electron micrographs show how tightly the intracellular microstructures are packed

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(a)

(b)

(c)

Figure 23.1. Cardiac myocyte. (A) Image of a ventricular cell in transmitted light. (B) Three-dimensional confocal imaging of transverse-axial tubular system (TATS). Optical cut through the center of the cell. The TATS tubules are marked with di-8ANEPPS. (Parts (A) and (B) reproduced with permission from Ref. 2.) (C) Electron micrograph; longitudinal ultrathin section. (Copyright © 2006 Biophysical Journal [7].) Oblique section shows the localization of junctional SR (jSR) in relation to T-tubules and mitochondria (M). MF, myofilaments; T, T tubule of TATS; Z, Z line.

in this peri-T-tubule region (Figure 23.1C) [7, 8]. Under pathological conditions cardiac cells undergo so-called ultrastructural remodeling, which increases the distance between the structures and makes them available for nano-objects.

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A

mDNA

Anti-apoptotic

Sarcoplasmic reticulum JC

Antigens T-tubule

Mitochondrion Nucleus Antisense oligos, siRNA, shRNA, drugs, CeO2 DNA B Transendothelial channel ~75 nm

Nano-object Plasmalemmal vesicles (caveolae) ~75 nm

Intercellular space ~10 nm

Capillary lumen

~200 nm

Adherens junction

Endothelial cells (internal tunic)

Tight junction

Basement membrane (middle tunic) Cells and fibers (adventitial tunic) Interstitial (perivascular) space C

E

D M M

M Z

N Z TT

Z

Figure 23.2. Nano-objects and barriers for their delivery to subdomains of cardiac myocytes. (A) Nano-objects and their cardiac intracellular targets. Simplified schematic representation of intracellular targets for nano-objects in ventricular cell. (Reproduced with permission from Ref. 2.). JC, Junctional cleft; mDNA, mitochondrial DNA; siRNA, synthetic small interfering RNA. (B) The capillary wall in the heart. (Copyright © 2006 Biophysical Journal [7].) Diagram schematically represents three mechanisms of cardiac vascular permeability: (1) transport by plasmalemmal vesicles, (2) transendothelial channel made by the fused vesicles, and (3) intercellular space. (C–E) Distribution of gold nanoparticles in permeabilized ventricular myocytes. Representative electron micrographs show the distribution of 3- (C) and 6-nm (D,E) particles within ventricular cells. M, mitochondrion; N, nucleus; Z, Z line, TT, T tubule. Ovals mark particles located deeper in the section and found with digitally enhanced contrast.

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CARDIAC VASCULAR PERMEABILITY

23.3

437

NANO-OBJECTS AND THEIR ANALYTICAL APPLICATIONS

All nanostructures with important analytical applications could be divided into two groups: (1) nano-objects (nanotubes, spherical nanoparticles, macromolecules, etc.) and (2) nanodevices (nanocapacitors, nanopores, nanocantilevers, etc.) [2]. Nano-objects can be used in a variety of bioanalytical formats [2]: (1) as quantitative tags, such as the optical detection of quantum dots and the electrochemical detection of metallic nanoparticles; (2) as substrates for multiplexed bioassays (encoded nanoparticles such as striped metallic nanoparticles); (3) as controllers of signal transduction (e.g., in colloidal goldbased aggregation assays); or (4) as catalyzers or inducers of biological processes (i.e., nonviral vectors). Chemotherapeutic and imaging nano-objects are usually conjugated to a chemotherapeutic drug (i.e., folic acid, paclitaxel, doxorubicin, etc.) and/or imaging agent (FITC, GFP, etc.). Nano-objects that have been suggested for biomedical research have diameters from 0.8 to 400 nm [2, 9]. Note that the two most popular groups of viral vectors for gene transfer are 20 nm (adeno-associated virus) and 60–90 nm (adenovirus) in diameter [10]. Although all nano-objects within the 2–100-nm size range were found to alter signaling processes essential for basic cell functions, 40- and 50-nm nanoparticles demonstrated the greatest effect [11]. Nano-objects could be injected directly into the tissue, but a pumping heart is not a convenient target for an injection. Therefore intravenous injection is the most appropriate way for delivery of nano-objects to the myocardium. Note that only nano-objects <100 nm were shown to be optimal for intravenous injection [12, 13]. The pathway for nano-objects from the bloodstream to the target intracellular subdomain in vivo has three barriers: (1) vascular wall, (2) cell membrane (sarcolemma), and (3) intracellular barriers. Experiments with gold nanoparticles showed that they have about 1 hour to pass the capillary endothelial barrier before being ingested by macrophages [14].

23.4

CARDIAC VASCULAR PERMEABILITY

One of the most critical issues for successful nano-object delivery is the ability of the nano-objects to penetrate the vascular wall. The most vulnerable place for the penetration of the nano-objects from the blood into myocardium is a thin blood capillary wall. Penetration of nanoparticles also occurs in arterioles and venules; however, in this case, they are rapidly ingested by phagocytic elements located along the vessels [14]. The penetration of particles through the capillary wall depends on transcapillary filtration pressure (Starling forces) and permeability pathways [15, 16]. The capillary wall (Figure 23.2B) has three layers: internal tunic (endothelial cells), middle tunic (basement membrane), and adventitial tunic (cells and fibers). However, structural details of the capillaries are different for different tissues and could lack the internal tunic and/or adventitial tunic [14].

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In the myocardium the blood capillaries are characterized by an extremely flat internal tunic (100–200 nm) and an irregular adventitial tunic. The middle tunic (20–50 nm) of the cardiac capillary wall consists mainly of collagenous and noncollagenous glycoproteins [14, 17]. Blood capillaries do not have a fenestrated endothelium encountered in many viscera, where the middle tunic is a major barrier between the vascular lumen and the interstitium [14]. In the cardiac capillary under normal conditions the primary barrier for nano-objects is the internal tunic, while the basement membrane is not a barrier at least for particles that can penetrate the internal tunic [14, 18]. There are three pathways for nano-objects through the capillary endothelial barrier: passage between endothelial cells, transporting vesicles (caveolae), and transendothelial channels [14, 19]. The endothelial cells are tightly connected by tight, gap, and adherens junctions [18, 20–22]. The density of the junctions varies between organs, and they are maximally enriched in the brain (blood–brain barrier), where strict control of endothelial permeability is exerted between the blood and the central nervous system. In the heart the distance between endothelial cells is ∼10 nm, and the gap is occupied by material of moderate density [14], which is probably junction structures and an overlay of two glycocalyxes (200–500 nm each [16, 18]). It was shown that the passage between endothelial cells is available only for objects <2 nm [19]. Nano-objects up to 50–70 nm in size could pass the endothelial barrier in the heart through membrane transporting vesicles, caveolae (the average diameter is ∼70 nm), and transendothelial channels formed by the fused vesicles [14, 19, 20]. It is important to note for drug delivery purposes that caveolae vesicles do not contain any enzymatic cocktail and the cargo to be delivered by nano-objects would not be degradated by lysosomal enzymes [23]. Taking into account that the thickness of endothelial cells in a heart capillary could be only 100 nm [19], the transendothelial channel could be a primary pathway for 2–50-nm particles. In other words, theoretically under normal conditions, the capillary barrier should be “transparent” for particles having dimensions of 50 nm. However, practically, even 15–30-nm particles were not found in the heart after intravenous injections [24, 25]. So far, only the delivery of cerium oxide nanoparticles (average particle diameter 7 nm) to cardiac myocytes was confirmed after intravenous injection [26]. Transportation of 30–50-nm particles by the transporting vesicles could be significantly improved with coating by albumin and/or opening of additional pathways with specific agents [15, 19, 27, 28]. The passage between endothelial cells could be “unlocked” and also used for delivery of nano-objects. For instance, the opening of interendothelial junctions (up to 2 μm) in capillary and venular endothelium could be induced with nitric oxide synthase inhibitor L-NAME [28]. Under some pathological conditions the cardiac capillary barrier for nanoobjects practically vanishes. For instance, hyperpermeability in the heart was shown to be triggered by inflammation or ischemia [20]. Also, during tissue inflammation the endothelial junctions were shown to widen (to 0.1–3 μm) in localized areas [18]. Acute inflammatory mediators, such as histamine and

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serotonin, significantly increase transportation of nano-objects by the vesicles and formation of transendothelial channels in cardiac endothelial cells [14]. Baldwin and Thurston [18] showed that histamine destroyed the endothelial barrier within 10 minutes. Thus the inflammatory mediators could be used to improve the transcapillary transport of nano-objects. The consequences of vascular permeability in vivo vary by location and depend on the situation. In the heart under normal physiological conditions the permeability response is tightly regulated and reversible. However, if platelets and leukocytes are recruited to the site of the leak, the tissue damage could be irreversible [20]. 23.5 TRANSMEMBRANE PATHWAYS FOR INTERNALIZATION OF NANO-OBJECTS Translocation of nano-objects across cell membranes depends on particle size, surface chemistry (coating), surface charge, and shape [2, 23]. There are four mechanisms for delivery of nano-objects through the cell membrane [2, 23, 29]: (1) clathrin-mediated endocytosis, (2) caveolae-mediated endocytosis, (3) macropinocytosis, and (4) nonspecific transport. Clathrin-mediated endocytosis is a primary mechanism of internalization of nano-objects <200 nm [30]. It occurs in a membrane region enriched in clathrin. Formation of an endocytotic vacuole is driven by polymerization of clathrin units [23, 29]. Note that this transporting mechanism leads nano-objects to degradative lysosomes. Therefore clathrin-mediated endocytosis should be avoided for drug and gene delivery unless the cargo is protected or this is necessary (e.g., for target delivery of lipophilic agents, or disruption of endosomes through pH-sensitive mechanisms) [2, 23, 31, 32]. This transportation could be facilitated by the inclusion into the nano-object coating of virus coat proteins (i.e., TAT from HIV-1) or their analogs, or cell adhesion molecules (CAMs): integrins, cadherins, selectins, and immunoglobulins [31, 33, 34]. Caveolae-mediated endocytosis is predominant for nano-objects >200 nm [30]. Caveolae are membrane invaginations lined by caveolin [29]. They are abundant in cardiac myocytes [35]. Unlike clathrin-mediated endocytosis, caveolae-mediated endocytosis is a highly regulated process, involving complex signaling [23]. Caveolae do not contain any enzymatic cocktail and could be employed as a primary transporting mechanism for drug and gene delivery with nano-objects. It was suggested that the switch between caveolae- and clathrin-mediated endocytosis could be made by pluronic P85 in a certain aggregation state: P85 unimers were shown to internalize through caveolaemediated endocytosis, while P85 micelles internalize through clathrinmediated endocytosis [36]. Macropinocytosis occurs via the formation of actin-driven membrane protrusions generating endocytic vesicles >1 μm in diameter [23, 29]. Macropinosomes do not have a specific coating. They can fuse with lysosomes and acidify. Macropinocytosis occurs in cardiac myocytes [37] and theoretically could be used for delivery of nano-objects up to 1000 nm in size.

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Nonspecific transport of nano-objects includes (1) clathrin- and caveolaeindependent endocytosis and (2) translocation that does not involve total enveloping of the nanoparticles by the membrane. The former involves cholesterol-rich microdomains, “rafts,” which are ∼50 nm in diameter and diffuse freely on the cell surface [29]. The latter depends on their electrostatic interactions with charged membrane domains governing the adsorption of the nanoparticles onto the cell membrane [23, 38]. This mechanism of uptake was shown for nano-objects of 20–60 nm in size [39, 40]. However, the use of cationic coating for nano-objects is problematic in the case of intravenous injection due to interaction with negatively charged serum proteins and red blood cells. Thus theoretically, for ventricular myocytes, there are four possible mechanisms for transport of nano-objects through the cell membrane. Although the mechanisms of nano-object transport through the sarcolemma of cardiac myocytes remain to be clarified, some morphological data suggest that it is similar to that described above. Two types of vesicles, coated and uncoated (∼100 and ∼50 nm correspondingly), that could be involved in transporting particles into cells have been demonstrated in cardiac myocytes [5, 41]. Due to the specific organization of ventricular myocytes, more than 50% of their sarcolemma belongs to T tubules (Figure 23.1B). The tubules are filled with glycocalics and under normal conditions are available only for particles <10 nm [7]. However, under some pathological conditions T-tubule dilation makes this sarcolemma opened for transportation of nano-objects. Independent of the mechanism of internalization, disruption of the endosomal membrane must occur to release the nano-objects. In the case of clathrin-mediated endocytosis such disruption could be induced by pH-sensitive mechanisms (such as hydrazone linkers or DOPE [23]). Also, mechanisms of osmotic swelling to disrupt lysosomes, or pH buffering to protect the cargo, could be employed [23, 38]. In the case of targeted gene delivery, it was found that, without the disruption of endosomes, at most only one plasmid out of 100 was effective [38]. However, even after a nano-object escapes the lysosome, its future could be questionable due to mechanical restrictions for diffusion. For example, it was shown that plasmid DNA (>20 nm in diameter) injected into the cytoplasm of rat myotubes mostly remained at its site of injection, probably being constrained by the actin cytoskeleton [38].

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INTRACELLULAR BARRIERS FOR NANO-OBJECTS

Nano-objects can reach an intracellular target by passive diffusion or actively by utilizing the microtubule network (retrograde transport to the nucleus), as has been shown for DNA [38]. Such an active microtubule-mediated cytosolic transport could probably also be employed for delivery to the nuclear pore complex (NPC) of similarly sized nano-objects [23]. The incorporation of Fab fragments of antibodies against the molecular motor protein dynein into the

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nano-object coating should significantly facilitate this transportation [38]. Recently, retrograde transport to the nucleus was shown to be enhanced with pluronics [42]. Free diffusion of nano-objects within cardiac myocytes has some restrictions [2, 7]. Moreover, some cellular subdomains such as mitochondrial intermembrane space, intermitochondrial contacts, mitochondria-junctional SR contacts, and junctional clefts are practically unavailable even to particles as small as 3 nm [2, 7, 43]. The 3-nm nano-objects diffuse freely to the nucleus and under some conditions can enter the mitochondrial intermembrane space (Figure 23.2C). Nanoparticles >6 nm in size cannot diffuse into the nucleus or mitochondrial intermembrane space (Figure 23.2D,E) due to the clearance within the NPC (∼5.5 nm) and porin (or VDAC; ∼3 nm) [2]. Certain parts of “free” intracellular space within cardiac myocytes are relatively unavailable for naniobjects >3 nm [7]. For instance, Figure 23.2E shows that after silver enhancement 6-nm particles can be found with much higher probability along Z-lines than along myofibrils. The 3-nm clearance excludes passive diffusion of nanoparticles into mitochondrial intermembrane space. Moreover, the 3-nm objects probably cannot enter the mitochondrial matrix, the zone of highest interest for nanomedicine [43, 44]. Therefore the use of the phenomenon of mitochondrial fusion was suggested for nanoparticle delivery [2, 23]. The most important target for contemporary bioresearch and therapy is the delivery of nano-objects to the nucleus. Cardiomyocytes do not undergo mitotic reorganization of the nuclear envelope, and for nano-objects the NPC is the only gate to the nucleus. There are two mechanisms of translocation across the NPC: passive diffusion and active transport. Although the NPC central channel has a limiting diameter of ∼25–30 nm [38], it is known that molecules <60 kDa (<3.5 nm [45]) and nano-objects smaller than 5.5 nm can freely diffuse through the NPC due to a polymeric mesh in the pore [46–48]. Contemporary polymeric gene delivery systems are between 40 and 400 nm in size [9]. This mainly keeps them out of the nucleus. So far only nano-objects <40 nm were shown to enter the nucleus [33]. Such transportation required the presence of a translocation signal (also known as a protein transduction domain or nuclear localization) in the coating of the nano-object to be recognized by the NPC transport receptors called karyopherins (importins α and β) and to form a nuclear pore targeting complex [38, 48]. There are many classes of signal localization sequences. Some of the signals (such as the peptide nuclear localization signal derived from the large T antigen of the SV40 virus, a peptide derived from the HIV Tat protein, adenovirus fiber protein, and modified integrin binding domain peptide) were already shown to be helpful in the delivery of nano-objects to the nucleus [33, 49–52]. Active nuclear transport is regulated by many mechanisms (such as phosphorylation/ dephosphorylation and protein cofactors [38, 48]) that have to be employed to facilitate the delivery of nano-objects to the nucleus.

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23.7 CHANGES IN VENTRICULAR CELL ULTRASTRUCTURE ASSOCIATED WITH HEART FAILURE The symptoms of heart failure include hypertrophy [53], defined as an enlarged heart size and muscle mass during dilated (congestive) cardiomyopathy. On the ultrastructural level, heart hypertrophy is characterized by lobulated nuclei, multiple intercalated disks, dilated T tubules, abnormal I bands, myofibrillar lysis, abnormally small mitochondria, and increased numbers of ribosomes [54–57]. Electron microscopy shows a significant reduction in the density of intermyofibrillar mitochondria and a reduction of mitochondrial number and size [2]. Dilated T tubules should make available for anchoring of nano-objects some proteins that are preferentially located within the TATS (such as L-type calcium and brain-type Na+ channels [58, 59]). These heart failure-related changes have to result in significant changes in the pathways for targeted nano-objects delivery [2]: (1) dilated T tubules approximately double the surface of the sarcolemma available for transport of nanoparticles and, consequently, the probability of delivery to failed cells, (2) diffusion pathways empty of glycogen and small mitochondria give more freedom for nano-objects to diffuse within failing cells, and (3) transformed mitochondria become much more available for nanoparticles. On the whole, even taking into account the slowing of physiological processes, the probability to deliver nano-objects to a certain subdomain of a failing cell could be significantly higher than for a healthy myocyte.

23.8 METHODS FOR STUDYING THE DELIVERY OF NANO-OBJECTS TO INTRACELLULAR SUBDOMAINS OF CARDIAC MYOCYTES Delivery of genes and drugs requires the creation of reliable delivery systems. The methods available for monitoring the delivery of nano-objects to intracellular subdomains could be divided into visualization of nano-objects and retrieval of the products of the nano-object delivery. The latter is especially important in the case of gene delivery when the synthesis of certain proteins (or their inhibition) clearly confirms the delivery. This method, however, has some restrictions. Overexpression of any protein in cardiac myocytes can result in heart failure, probably due to obstruction of diffusion pathways [60]. Therefore protein synthesis must be inhibited with doxycycline [61, 62]. This, however, reduces the synthesis of other cell proteins as well. The former, visualization, clearly shows the location of nano-objects. Unfortunately, the popular method for intracellular and intraorganellar localization of nano-objects with confocal microscopy is significantly complicated in cardiac myocytes due to multiple membrane invaginations, low resolution of confocal systems, and out-of-focus light [63]. To avoid these problems we

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improved the electron microscopy method developed by Feldherr [2, 7, 64]. The method employs electron microscopy, water-soluble resin for cell polymerization, and silver enhancement within ultrathin sections. This approach allows precise localization of 2-nm gold particles that could be tagged to any nano-object.

23.9 CONFIRMED DELIVERIES OF NANO-OBJECT TO CARDIOMYOCYTES IN VIVO Torchilin’s laboratory reported the successful delivery of DNA- or ATPloaded immunoliposomes specific for cardiac myosin to cardiomyocytes of ischemic myocardium after intravenous injection [65, 66]. This significantly improved heart contractility after global ischemia. Kolattukudy’s laboratory demonstrated cardioprotective effects of cerium oxide (CeO2) nanoparticles in cardiomyopathy [26]. The nanoparticle treatment attenuated progressive cardiac dysfunction and remodeling in a murine model of ischemic cardiomyopathy. Authors attributed this beneficial effect of CeO2 nanoparticles to their autoregenerative antioxidant properties inhibiting myocardial oxidative stress, ER stress, and inflammatory processes. Two groups reported successful cardiac transgene expression in vivo using an adeno-associated virus and cardioselective promoters for intravenous injection. Although under these conditions all organs were shown to be infected, the promoters allow activation of delivered genes only in cardiomyocytes [67]. Koch’s group targeted myocardial beta-adrenergic receptor signaling and calcium cycling with an adeno-associated virus [68]. They showed a stable myocardial-specific expression of a therapeutic transgene, the calcium Ca2+sensing S100A1, which resulted in functional heart failure rescue in a rat model of heart failure [69]. Recently, Metzger’s laboratory performed in vivo transgene expression of a cytosolic Ca2+ buffer parvalbumin and adult cardiac Troponin I isoform cTnI A164H [50, 70]. The former significantly accelerated relaxation of the heart without affecting cardiac morphology or systolic function [70]. The latter significantly improved systolic and diastolic cardiac function and mitigated reperfusion-associated ventricular arrhythmias [10, 50, 70].

23.10 CONCLUSION To exert their therapeutic action, many pharmacological agents and large regulatory molecules (i.e., anti-apoptotic drugs, enzymes, siRNA, shRNA, etc.; Figure 23.2A) have to be delivered intracellularly. Precise delivery of different biologically active molecules to cellular subdomains of failing cardiac myocytes can benefit the entire heart function. Although the delivery of nanoobjects to subdomains of a cardiac cell is complicated and many aspects of the

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delivery remain to be clarified, the contemporary state of the field is very promising.

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CHAPTER 24

Design Parameters Modulating Intracellular Drug Delivery: Anchoring to Specific Cellular Epitopes, Carrier Geometry, and Use of Auxiliary Pharmacological Agents SILVIA MURO Center for Biosystems Research, University of Maryland Biotechnology Institute, and Fischell Department of Bioengineering, University of Maryland, College Park, Maryland

VLADIMIR R. MUZYKANTOV Department of Pharmacology and Targeted Therapeutics Program of Institute of Translational Medicine and Therapeutics, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania

24.1 INTRODUCTION: INTRACELLULAR DELIVERY OF DRUG CARRIERS Subcellular destination of drugs determines their effects, metabolism, and duration. Every drug action requires specific subcellular localization. For example, RNA interference should take place in the cytosol, transgene induction in the nucleus, shift of pro-/anti-apoptotic balance in the mitochondria and cytosol, modulation of protein synthesis in the endoplasmic reticulum and the Golgi complex, and certain types of enzyme replacement therapy in endosomal and lysosomal vacuoles. However, subcellular targeting of drugs and their carriers is an extremely challenging and still largely elusive goal. Coordinated efforts of experts in diverse fields are necessary to achieve this goal. The nature of a drug dictates both desirable cellular compartments for therapeutic action and, in some cases, the entry path. For example, viruses Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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developed very effective means of delivering nucleic acids into the cells, providing a basis for gene therapies. Small soluble drugs diffuse through the cellular membranes and utilize transmembrane transporters including ion channels, thereby getting access to these compartments. Conjugating drugs with small ligands having specific affinity to certain organelles (e.g., mitochondrial or nuclear localization sequences) facilitates their accumulation in these compartments. These approaches are described in other chapters. However, the nonviral biotherapeutics (therapeutic proteins, enzymes, and genetic materials) and drug carriers have no free intracellular access, nor affinity to compartments of interest. One strategy to deliver these cargoes and carriers into a cell is to couple them with peptides containing sequences of protein transduction domains (PTDs), for example, those derived from the HIV transcription factor, Tat [1, 2]. Tat basic charge mediates binding to the negatively charged components of plasmalemma, facilitating its uptake into the cells [1–3], which may occur via passive endocytosis of surface-bound Tat carriers or other still elusive mechanisms involving PTD insertion into the plasmalemma and direct transmembrane transfer of cargoes [1]. This strategy, described in other chapters, may find interesting applications in experimental biomedicine. Arguably, an ideal drug delivery system should provide specific binding to a target cell (unavailable for PTDs, most of which bind promiscuously to diverse cell types). From this standpoint, binding drugs or drug carriers to specific surface determinants or epitopes expressed preferentially on cells of a given type or phenotype (e.g., pathologically altered cells) represents an attractive strategy. Furthermore, binding of drugs or drug carriers to these epitopes may facilitate intracellular delivery. It has been known for several decades that binding to certain cellular receptors, such as Fc receptor in phagocytes or transferrin receptor in nonphagocytic cells, facilitates internalization of the ligands via endocytic uptake into cellular vesicles [4]. More recent studies, including those discussed in this chapter, found that depending on the cellular mechanism of endocytosis, the nature of engaged epitopes, and such specific features of a drug delivery system as size, shape, and valency of binding sites, one can get a high degree of control of the mechanism and rate of endocytic internalization, as well as the subsequent intracellular trafficking, sorting, and final destination of the delivered drug carriers and cargoes. This chapter will focus on the most recent findings in this area, that is, exploitation of a novel endocytic mechanism, cell adhesion molecule- or CAM-mediated endocytosis, for intracellular delivery of biotherapeutics into vascular endothelial cells. The monolayer of endothelial cells lining the luminal surface of blood vessels controls vascular tone, blood fluidity, and extravasation [5]. In particular, endothelium plays a central role in inflammation and vascular oxidative stress, syndromes involved in the pathogenesis of stroke, ischemic heart disease, acute lung injury, atherosclerosis, hypertension, diabetes, and genetic

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diseases [6]. Targeted delivery of drugs to desired compartments within endothelial cells holds significant promise as a means to improve pharmacological management of these conditions [7, 8]. Examples of controlled subcellular transport of therapeutic enzymes (e.g., endosomal addressing of antioxidant enzymes vs. lysosomal addressing of enzyme replacement therapy for lysosomal storage diseases) illustrates this paradigm.

24.2 VESICULAR ENDOCYTIC PATHWAYS: NATURAL GATEWAYS FOR INTRACELLULAR TRANSPORT AND DELIVERY OF DRUGS Endocytic pathways, naturally useful for intracellular drug delivery, include (1) macropinocytosis and phagocytosis, the main pathways in macrophages allowing uptake of extracellular fluid or large particulate ligands, respectively [9–12]; (2) classical clathrin- and caveolae-mediated endocytosis, used by most cell types for internalization of diverse ligands and turnover of plasmalemma components [13, 14]; and (3) various nonclassical pathways bypassing clathrin and caveolar endocytosis [15–17] including the recently defined endothelial CAM-mediated endocytosis, described in more detail below. To some extent, every pathway serves certain specific ligands (e.g., transferrin for clathrin pathways, albumin for caveolae, IL-2 for nonclassical pathways) and exerts preferential sensitivity to pharmacological inhibitors (Table 24.1). The internalization pathway determines the destination of the internalized materials, for example, macropinocytosis phagocytosis and clathrin-mediated pathways generally deliver materials via endosomes to lysosomes [4, 18], while caveolae-related endocytosis delivers materials to other compartments including the Golgi complex or the endoplasmic reticulum (ER), and to a lesser extent to lysosomes [19, 20]. Endocytic vesicles internalized via clathrin- and more typically caveolae-mediated endocytosis can also traverse the cell body, thus transporting materials across cells, a process known to as transcytosis [20, 21]. Targeting drugs to the cellular receptors naturally associated with specific endocytic pathways often leads to internalization and intracellular trafficking typical of these pathways [22]. For example, liposomes targeted to the selectins whose turnover in endothelium naturally occurs via clathrin-mediated endocytosis [23, 24] enter cells via clathrin-coated pits and traffic to lysosomes [25, 26]. Imaging probes and drugs conjugated to transferrin receptor antibodies bind to this receptor expressed on the luminal surface of brain endothelium, enter cells via clathrin-coated pits, and traverse the endothelial monolayer, similar to the endogenous ligand transferrin, thus supporting drug delivery through the blood–brain barrier [27]. Drug delivery systems exploiting caveolae-mediated endocytosis—for example, compounds conjugated to antibodies to gp90, a 90-kDa glycoprotein located in the caveolae—provide

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TABLE 24.1

Endothelial Pathways for Endocytic Internalizationa

Pathway Phagocytosis

Macropinocytosis

Clathrinmediated

Caveolaemediated CCI

CAM-mediated

Inhibitors Dynamin and actin disruption (cytochalasin), nocodazole, H-7, wortmannin Amiloride, nocodaloze, BIM-1, H-7, staurosporine, wortmannin Dynamin and actin disruption (cytochalasin), K+ depletion, MDC, amantidine Dynamin disruption, cyclodextrin, filipin, genistein Dynamin and actin disruption

Dynamin and actin disruption (latrunculin), amiloride, EIPA, Y27632, radicicol

Ligands

Initial Vesicle Size (nm)

Bacteria, opsonized particles, immune complexes

Predominantly 1–10 μm

Fluid-phase uptake; activated by EGF

Predominantly 1–10 μm

TF, VCAM-1, ACE

Up to 250 nm

Cholera toxin, caveolar ligands

Up to 70–100 nm

High dose of EGF?, anthrax toxin?, IL-2β, fibroblast growth factor 2 via syndecan 4, some GPI-anchored proteins Multivalent ligands binding to PECAM-1 or ICAM-1

Varies from 50 to 250 nm

From ∼50 nm to 5 μm

a

Phagocytosis and pinocytosis represent the oldest evolutionary pathways. Pinocytosis includes micro- and macropinocytosis, which yield vesicles of different sizes. In some instances, clathrinand caveolae-mediated encodytosis lead to uptake of fluid phase along with ligands, hence contributing to micropinocytosis. Profound disruption of dynamin (e.g., by dominant-negative constructs) and actin (by latrunculin that more severely than cytochalasin inhibits actin reorganization) impairs all vesicular pathways, as these proteins control formation and pinching off of all types of vesicles. Nocodazole affects microtubuli. Wortmannin inhibits PI3 kinase. Staurosporine is a broad spectrum kinase inhibitor. BIM-1 and H-7 inhibit PKC. Genestein inhibits tyrosine kinases, mostly those involved in caveolar endocytosis. Y27632 inhibits Rho-dependent kinase ROCK. Radicicol inhibits Src kinase. MDC or monodansyl cadaverin inhibits clathrin lattices arrangement; amantidine inhibits budding of clathrin-coated pits. Cyclodextrin and filipin deplete and sequester, respectively, cholesterol in lipid rafts and caveolae. Amyloride inhibits sodium– proton exchangers, NHE, and other ion pumps; EIPA more specifically inhibits NHE1 in the plasmalemma TF, transferrin; ACE, angiotensin converting enzyme; CCI, clathrin- and caveolaeindependent endocytosis is not well defined, but in some instances shares some features of the classical pathways, and does not have specific inhibitors.

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transcellular delivery across endothelium, especially effectively in the pulmonary microvasculature [28]. The lysosome is the most typical final destination for most ligands taken up by cells via vesicular pathways [29]. When ligands are taken into the nascent subplasmalemmal vesicles and early endosomes, transmembrane H+ATPases and Cl− channels acidify the endosomal lumen, while the Na+–H+ exchanger-6 (NHE6) favors H+ efflux to help regulate the endosomal pH [30]. This mechanism maintains a mildly acidic pH in the early endosome lumen (e.g., 6.3–6.5), favoring separation of ligands from their receptors and binding of coat proteins to the endosomal membrane, necessary for lysosomal biogenesis [31]. The specific itinerary of intracellular sorting vesicles and the intracellular destinations of internalized ligands may vary depending on the type of receptor, ligand size, endocytic pathway, cell type, and status [32–36]. As discussed above, caveolar endocytosis delivers some ligands to the recycling endosomes, ER, and subendothelial space, in addition to lysosomes [14]. Yet components of the plasmalemma and their ligands taken via vesicular pathways can meet in common sorting vesicles including early endosomes. However, our present understanding of the internalization mechanisms and subsequent trafficking pathways of nanoconjugates targeted to endothelium is grossly incomplete. For instance, vascular cell adhesion molecule-1 (VCAM1) is naturally internalized from endothelial surface and transported to lysosomes via clathrin-mediated endocytosis [37]. VCAM-1 antibodies have been tested for targeting of imaging agents or biodegradable carrier particles to endothelial cells involved in inflammation, both in vitro and in vivo [38–40]. Yet the phage-display library approach helped to identify diverse epitopes on VCAM-1 molecule providing a differential rate of internalization [41]. Furthermore, the itinerary of intracellular trafficking, final destination, and the fate of drugs targeted to VCAM-1 and most other identified epitopes remain to be determined. In addition, potential side effects of engaging of a cellular epitope involved in a given endocytic pathway represent an important safety issue in the context of drug delivery. Examples of such side effects include masking of a receptor and/or inhibition of its biological function, as a result of stimulation of its shedding from the plasmalemma or its endocytosis. Obviously, side effects that defeat the purpose of therapeutic intervention must be avoided. Targeting endothelial surface determinants other than endocytic receptors can also provide intracellular delivery of nanoconjugates via induction of endocytic processes that normally are not utilized by target cells. Targeting of nanoconjugates to transmembrane glycoproteins ICAM-1 and PECAM-1 (intercellular adhesion molecule-1 and platelet-endothelial cell adhesion molecule-1, i.e., cell adhesion molecules or CAM) primarily expressed by endothelial cells, illustrates this scenario and provides an interesting paradigm for safe and effective intracellular delivery of diverse drugs and carriers [42].

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24.3 CAM-MEDIATED ENDOCYTOSIS: AN INDUCIBLE PATHWAY FOR INTRACELLULAR DELIVERY OF A WIDE RANGE OF DRUG CARRIERS PECAM-1 and ICAM-1 are Ig-superfamily endothelial transmembrane glycoproteins involved in leukocyte adhesion and transmigration, cellular recognition, and signaling [43–45]. They represent good epitopes for endothelial targeting of drugs [46]. Leukocyte adhesion to endothelium is an important step in vascular inflammation involved in the pathogenesis of other diseases [47–49]. ICAM-1 and PECAM-1 blockade suppresses this phase of inflammation [50, 51]; hence drug targeting to these molecules may suppress inflammation. Studies in diverse animal species revealed no harmful effects of drug targeting directed to PECAM-1 [8, 52–58]. Endothelium constitutively stably expresses approximately 2 × 105 and 106 copies of ICAM-1 and PECAM-1, respectively, on the surface of one cell; ICAM-1 expression is further enhanced upon pathological conditions [52, 59]. Enzymes, genetic materials, nanocarriers, liposomes, and other cargoes and carriers conjugated or fused with ICAM-1 and PECAM-1 antibodies (antiICAM and anti-PECAM, generally termed anti-CAM thereof) bind to endothelial cells in culture and, more importantly, in the vasculature in intact animals [8]. In particular, the pulmonary endothelium is a preferential site for CAM-targeted drug delivery, because (1) it is the first major capillary network encountered by drugs after IV injection; (2) it contains 30% of endothelial in the body; and (3) it receives the whole cardiac output of venous blood [8]. Drugs, enzymes, DNA, and diverse nanocarriers conjugated with antiPECAM and anti-ICAM accumulate in the lungs after intravascular injection, providing delivery to the pulmonary endothelium [8, 52–58]. Docking of natural ligands (e.g., leukocytes) to ICAM-1 or PECAM-1 does not prompt uptake by endocytosis, but results in proteolytic shedding of these molecules from the plasmalemma and ligand release [60, 61]. Similarly, monomeric ICAM-1 or PECAM-1 antibodies are not internalized by endothelial cells [52, 62, 63]. This feature, that is, anchoring on the cell surface without speedy internalization, ideally suits the needs for targeted delivery of antithrombotic agents and other drugs that are supposed to act in the vascular lumen. For example, monovalent recombinant proteins fusing plasminogen activators with a single chain fragment of anti-PECAM bind to endothelium and stimulate dissolution of pathological intravascular blood clots in animal models [54, 64, 65]. However, when antibodies to PECAM-1 or ICAM-1 bind to cells as multivalent complexes (e.g., as multimeric protein conjugates or nanocarriers coated by anti-CAM, which, for the sake of simplicity, will all be referred to as “nanocarriers” throughout the text below, unless specified otherwise), they cluster their antigens in the endothelial surface and enter cells very rapidly (t1/2 = 15–20 min) and efficiently (90% internalization of the cell-bound nanocarriers) [63, 69].

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Of note, cell culture studies are marred with artifacts associated with loss of cellular phenotypes adequately reflecting cellular status in vivo and prolonged exposures to nonphysiologically high doses of drugs and nanocarriers. For example, endothelial cells in culture display a rather degenerative phenotype due to loss of flow adaptation, including loss of actin stress fibers maintaining flow alignment, restricted plasmalemma deformability to minimize compromising the endothelial permeability barrier, and increased activity of certain endocytic pathways and related signaling platforms. Importantly, both flow-adapted endothelial cells in vitro and vascular endothelium in lab animals displayed effective and nondamaging uptake of diverse carriers targeted to ICAM-1 and PECAM-1 in both the large vessels and the microvasculature [67, 68]. Internalization of anti-CAM conjugates by endothelial cells requires dynamin, a large GTPase typically involved in phagocytosis and clathrin- and caveolar-mediated endocytosis [69]. However, the cellular mechanism of uptake and subsequent intracellular trafficking of anti-ICAM or anti-PECAM conjugates have unique features that distinguish this pathway from previously known endocytic pathways. Summarizing a decade of research in this area by our laboratories, anti-PECAM and anti-ICAM conjugates bind multivalently to CAM and trigger CAM-mediated endocytosis, defined by the following unique features [22, 55, 66, 69, 70]. 1. It is mediated by tyrosine phosphorylation of the CAM cytosolic domain (particularly described for PECAM-1), which provides the endocytosis signal [55]. 2. It is not sensitive to inhibitors of clathrin- and caveolae-mediated pathways and does not colocalize with ligands or endocytic markers of these pathways [69, 71]. 3. It is suppressed by inhibitors of the plasmalemmal sodium–proton exchanger NHE1 (e.g., amiloride and EIPA), but not by wortmannin and nocodazole, which suppress macropinocytosis and phagocytosis via inhibition of PI-3 kinases and microtubules, respectively [69, 72]. 4. It is not affected by the ionophore monensin, indicating that ion exchange, important for regulating osmolarity during macropinocytosis, is not involved in CAM-mediated endocytosis, and NHE1 rather anchors actin filaments needed for the uptake [69, 72]. 5. It is blocked by the actin fiber-disrupting agent latrunculin A (but not cytochalasin D that caps short actin filaments and inhibits macropinocytosis and clathrin-mediated endocytosis) and is driven by the formation of actin fibers (not actin cups involved in macropinocytosis and phagocytosis) and endocytic vesicles mediated by Rho-dependent kinase (ROCK), Src kinase, PKC, and dynamin mediated signaling [69]. Figure 24.1 schematically illustrates CAM-mediated endocytosis, involving activation of PKC, Src kinases, and Rho-mediated pathways, leading to

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Figure 24.1. Intracellular drug delivery into vascular endothelial cells using anti-CAM nanocarriers. Multivalent anti-CAM/nanocarriers and conjugates bind to and cluster cell surface CAMs. This activates PKC, Src kinases, and Rho-dependent kinase (ROCK), enzymes that regulate the activity of the GTPase, dynamin, and the amiloridesensitive sodium-proton exchanger, NHE1, which induces formation of actin stress fibers. CAMs and NHE1 form a complex, which may help crosslink actin fibers to the CAM cytosolic tail through α-actinin (α-act) and ezrin/radixin/moesin (ERM) family proteins. After internalization, anti-CAM/nanocarriers traffic via a microtubuledependent process to endosomes (1–2 h), which can be identified by early endosome antigen-1 (EEA1). Endosomes are enriched in NHE6, an intracellular ion exchanger that helps regulate endosome acidification by vacuolar H+-ATPase. Anti-CAM/nanocarriers dissociate from CAMs and NHE1 in endosomes and CAM recycles to the plasma membrane. Nanocarriers arrive at lysosomes about 3 hours after their internalization, which can be identified by colocalization with lysosome-associated membrane protein 1 (LAMP1). In lysosomes, NHE6 becomes inactive, favoring further acidification and degradation of delivered proteins by acidic proteases. Nocodazole (which disrupts the cell microtubule network), chloroquine (a mild base that inhibits lysosomal acidification), and monensin (which enhances Na+–H+ exchange in endosomes and induces recycling of anti-CAM nanocarriers to the plasma membrane) alter the intracellular itinerary of anti-CAM/nanocarriers and prolong their effects. From reference 114.

formation of actin stress fibers, apparently associated with nascent endocytic vesicles [69, 72]. CAM-1 clustered by anti-CAM/nanoconjugates interacts with NHE1 (particularly described for ICAM-1) [69, 72], which may serve as an adaptor for the cytoskeletal proteins α-actinin and ERM family proteins (ezrin, radixin, and moesin), favoring formation of bundles of actin filaments at the plasmalemma sites where conjugates are bound. Cytoskeletal

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reorganization and dynamin activity induce invaginations that pinch off the plasma membrane, driving anti-ICAM/nanoconjugates and, concomitantly, ICAM-1 and NHE1, into vesicular compartments in the cytosol [42, 69, 72]. Within an hour after uptake, anti-CAM/nanoconjugates traffic to early endosomes (Figure 24.1), containing early endosome antigen 1 (EEA1) and an intracellular NHE isoform, NHE6 [72]. NHE6 regulates ion balance and maintains a moderately low pH in the endosome, required for ligand dissociation from receptor and lysosomal biogenesis [73, 74]. Indeed, anti-ICAM/ nanoconjugates dissociate from ICAM-1 within this compartment and ICAM1 recycles back to the plasma membrane, while nanoparticles arrive to lysosomes about 3 h after internalization [42, 66]. ICAM-1 recycling supports recurrent targeting of subsequent doses of anti-ICAM/nanoconjugates, since a single target molecule can undergo many rounds of uptake of circulating conjugates, enhancing the efficacy of delivery [42]. Typically, ligands internalized via other pathways including clathrinmediated mechanisms, macropinocytosis, and phagocytosis are delivered to lysosomes within minutes [10, 75]. Underscoring mechanistic differences from these pathways, cargoes entering cells via CAM-mediate endocytosis arrive in the lysosomes at a relatively slow pace, approximately 2 hours after the uptake in most cases [66]. Thereafter, NHE6 becomes inactive within lysosomes, and the pH drops due to the continuous activity of the H+-ATPase [30], which activates lysosomal acidic hydrolases and, in most cases, results in proteolytic degradation of the delivered proteins (Figure 24.1).

24.4 EXAMPLES OF INTRACELLULAR DRUG DELIVERY VIA CAM ENDOCYTOSIS: ANTIOXIDANT AND LYSOSOME REPLACEMENT ENZYMES Presumably, biomedical use of a plethora of therapeutic, diagnostic, and imaging agents can benefit from their targeted delivery into endothelial cells— nucleic acids, molecular beacons, inhibitors of cellular regulatory proteins, and reporter and therapeutic enzymes. Here we consider intracellular delivery strategies for two types of therapeutic enzymes—antioxidants (developed by Muzykantov’s lab) and lysosomal hydrolases (developed by Muro’s lab). 24.4.1 Antioxidant Enzymes Abnormally high levels of influx of reactive oxygen species, such as hydrogen peroxide (H2O2) and superoxide anion, cause oxidative stress and cellular damage. A prolonged administration of megadoses of nonenzymatic antioxidants (e.g., N-acetyl cysteine) may provide some degree of alleviation of chronic and relatively subtle oxidative stress, such as in early stages of atherosclerosis, but provide no significant protection in acute severe conditions, such as ischemia reperfusion, stroke, and acute lung injury [76]. Importantly, endothelium represents the key therapeutic target in these conditions. In theory,

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the extremely potent antioxidant enzymes catalase and superoxide dismutase SOD (that decompose H2O2 and superoxide, respectively) could provide more effective protection. Furthermore, intracellular delivery of antioxidant enzymes would help to detoxify oxidants otherwise poorly accessible for circulating drugs. Alas, catalase and SOD have no medical utility, in large part due to inadequate pharmacokinetics (both enzymes get eliminated within 5–10 min) and the lack of effective means for their delivery into the target cells, including endothelial cells [77]. CAM-mediated endocytosis, taken with other features of drug targeting to ICAM-1 and PECAM-1 (e.g., anti-inflammatory effect of blocking leukocyte adhesion to endothelial cells) provides the basis for intracellular delivery of antioxidant enzymes to endothelium. Indeed, catalase and SOD conjugated with anti-CAM (but not naked, PEG-derivative or IgG-conjugated enzymes) bind to endothelial cells in culture, ex vivo in perfused isolated lungs and in vivo after intravascular injection [52, 57, 78]. Anti-CAM/catalase conjugates enter endothelial cells and protect against damage by H2O2 in vitro, in perfused organs and in intact animals [52, 57, 58, 79, 80]. Similarly, superoxide dismutase conjugated with anti-PECAM (i.e., anti-PECAM/SOD, average size ∼300 nm) also bound to human endothelial cells and attenuated necrotic and apoptotic types of oxidative stress caused by either extracellular or intracellular influx of superoxide anion [78]. Furthermore, anti-PECAM/catalase injection in donor rats prior to procurement of the lung graft markedly improves the outcome of transplantation, including normalization of blood oxygenation of the recipient animals [57]. The relatively slow lysosomal traffic of anti-CAM/nanocarriers permits reasonably prolonged activity of targeted therapeutics, such as several hours of antioxidant protection conferred by delivered catalase [66]. Since H2O2 easily diffuses through cellular membranes, including the membranes of intracellular vesicular compartments such as endosomes, targeted catalase trafficked into endosomes degrades H2O2 and protects cells against oxidative stress without the need for endosomal escape [66]. However, the effect of catalase delivered intracellularly by anti-CAM/nanoconjugates decays within 3 hours after internalization due to lysosomal trafficking and degradation (Figure 24.2) [66]. Encapsulation of catalase into biocompatible polymer nanocarriers selectively permeable to H2O2, but not to proteases, permits a prolonged antioxidant effect (Figure 24.2). These carriers thus act as a protective cage, capable of therapy without traditional “drug release.” A freeze–thaw modified double emulsion allows encapsulation of active catalase into such protective nanocarriers with controlled size and shape, appropriate for vascular delivery into endothelial cells [81, 82]. Typical carrier morphologies are nanospheres with diameters <500 nm [81] or flexible filaments that are a few microns in length and only ∼50 nm in cross section [82]. Morphology and loading of nanocarriers are controlled by chemistry of the polymers utilized, for example, molecular weight and ratio of hydrophilic to hydrophobic domains in the used

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(A)

(B)

Figure 24.2. The concept of vascular delivery of catalase by semipermeable polymer nanocarriers targeted to CAM. (A) Catalase conjugated with antibodies against endothelial cell adhesion molecules (CAM) bind to endothelial cells, get internalized via CAM-mediated endocytosis, and remain active intracellularly for a few hours, thus protecting cells against injurious effects of H2O2, either produced by endothelial cells (e.g., in mitochondria, M) or diffusing into endothelium after being released by activated white blood cells (WBCs). In addition to detoxification of H2O2, blocking CAM by conjugates inhibits WBC adhesion. (B) Once immunoconjugates are trafficked into the lysosomes, proteases degrade catalase and H2O2 escapes enzymatic interception (left). In contrast, catalase loaded into permeable, biodegradable polymer nanoparticles, PNC, will remain protected against lysosomal proteolysis and decompose H2O2 even after degradation of proteins on the PNC surface.

amphiphilic diblock copolymers (e.g., PEG-PLGA). Not only does this approach enable significant loading of active catalase, but the cargo enzyme is protected against external proteolysis. Catalase-loaded polymer nanocarriers targeted to PECAM-1 deliver active cargo to the pulmonary vasculature after IV injection in animals and protect endothelial cells against H2O2-induced injury for durations on the order of many hours [83]. 24.4.2 Lysosomal Enzyme Replacement Therapy Lysosomal destination of anti-CAM conjugates offers an advantage in the context of other therapeutic interventions, such as enzyme replacement

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therapies for lysosomal storage disorders, a group of syndromes primarily caused by inherited deficiency of lysosomal hydrolases, several of which present a marked endothelial component [84, 85]. Enzyme replacement therapy is one the most viable treatments for these diseases [85, 86]. This strategy relies on the presence of certain sugar residues in lysosomal enzymes, which provides binding to cell surface receptors such as mannose or mannose-6-phosphate receptors, leading to enzyme uptake and delivery to lysosomes by clathrin-mediated endocytosis [84, 87]. However, inadequate glycosylation of recombinant enzymes and/or lack of appropriate receptors in LSD cells are obstacles affecting delivery and efficacy of enzyme replacement therapies [88]. Such glycosylation requirements can be bypassed by delivering these recombinant enzymes using CAM-targeted carriers, which efficiently bind to CAMs both in cell cultures and when injected IV in laboratory animals [68, 89, 90]. As an example of this type of application, coupling of recombinant acid sphingomyelinase (ASM, an enzyme whose genetic deficiency in types A and B Niemann–Pick disease results in lysosomal accumulation of sphingomyelin) to anti-ICAM/nanocarriers resulted in a marked elevation of the levels of enzyme delivered to body organs (e.g., one to two orders of magnitude in the case of the lungs [89]). After binding to endothelial cells, anti-ICAM/ASM/carriers efficiently entered cells via nonclassical CAM-mediated endocytosis, providing a means to markedly enhance effective enzyme delivery to endothelial cell lysosomes [84, 89, 90]. This in turn resulted in a significantly enhanced attenuation of the sphingomyelin storage typical of Niemann–Pick disease [68, 84, 89, 90]. Interestingly, even in the presence of glycosylated enzymes on the carrier surface, which could potentially bind to their natural receptors and be internalized via mechanisms of clathrin-mediated endocytosis, anti-ICAM/carriers bearing recombinant lysosomal enzymes are transported into endothelial cells only via CAM-mediated pathway, indicating the dominant efficiency of this route and the potential utility if this strategy [90].

24.5 OPTIMIZATION OF INTRACELLULAR DRUG DELIVERY VIA CAM-MEDIATED ENDOCYTOSIS USING PHARMACOLOGICAL AGENTS, SELECTION OF THE EPITOPE, AND CARRIER GEOMETRY Understanding the mechanisms of interactions between anti-CAM conjugates and endothelial cells permits one to optimize subcellular delivery and, likely, therapeutic effects of these conjugates. For instance, the NHE1 inhibitor amiloride delays internalization of anti-CAM/nanoconjugates by endothelial cells [69], which may suit therapeutic interventions intended for the vascular lumen [62]. Furthermore, disruption of the cell microtubular network by nocodazole inhibits lysosomal trafficking of anti-CAM/ nanoconjugates, while inhibition of lysosomal acidification by the mild base, chloroquine, protects

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delivered proteins against proteolysis (Figure 24.1) [66]. Monensin which enhances Na+–H+ exchange in early endosomes, favors sorting of nanoconjugates to a recycling pathway [72]. All these pharmacological agents markedly prolonged the antioxidant effect of anti-ICAM/nanoconjugates of catalase [11, 66, 72]. Therefore auxiliary pharmacological interventions may help modulate the subcellular delivery, longevity, and effects of therapeutics targeted to endothelial cells by targeted nanoconjugates. This goal can also be achieved by selection of optimal binding sites on the target cellular determinant. It has became evident in recent years that anchoring to different epitopes on the same cellular molecule may offer different rates of intracellular uptake. For example, as mentioned above, phage display screening identified peptides that bind to VCAM-1 and differentially facilitate constitutive endocytosis of this target determinant [37]. This feature seems to enhance vascular VCAM-1 imaging in animal models of inflammation [41]. Studies of CAM-mediated endocytosis described below even further expanded this important paradigm. 24.5.1 Control of Internalization and Intracellular Trafficking by Selecting CAM Epitopes for Carrier Binding Given that signaling via PECAM-1 cytosolic domain is involved in endothelial uptake of anti-PECAM/nanocarriers via CAM-mediated endocytosis [55, 69], it seemed plausible that selection of specific PECAM-1 epitopes for antiPECAM/nanocarrier binding might play a role in the subsequent internalization process. The extracellular portion of human PECAM-1 consists of 574 amino acids containing six Ig-like domains, numbered from 1 to 6, from the most membrane-distal to the most membrane-proximal domain (Figure 24.3) [91]. PECAM-1 molecules in neighboring endothelial cells interact in a homophilic manner, and specific PECAM-1 epitopes differentially involved in this interaction have been identified [92, 93, 95]. Monoclonal antibody mAb62 directed to PECAM-1 Ig domain 1 efficiently binds to endothelial cells and delivers to endothelium submicron and micron sized protein conjugates and nanocarriers [52, 57, 63, 69, 78, 83, 92, 95]. We have compared endothelial binding, uptake, and trafficking of model spherical carriers (100-nm diameter) coated with mAb62 with four other mAbs directed to different PECAM-1 extracellular epitopes: mAb4G6 binds to membrane-proximal domain 6, mAbGi34 binds between domains 2 and 3, and mAb35 and mAb37 bind to membrane-distal domain 1 (Figure 24.3). Nonconjugated mAb62 and mAb37 displayed the highest binding level, mAb35 and mAb4G6 (recognizing the most distal and proximal PECAM-1 domains, respectively) bound slightly less effectively, whereas mAbGi34 recognizing a middle area of PECAM-1 showed the lowest level of binding to endothelial cells. However, nanocarriers coated with these antibodies (mAb/NC, ∼200-nm diameter) showed a binding pattern different from that

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PECAM-1 Homophilic N-tss ss interaction s

s s s s s s s Extracellular domain ss

Cytosolic domain C-t

Figure 24.3. Epitope controlled subcellular destination of anti-CAM nanocarriers. PECAM-1 structure is comprised of six Ig-like extracellular domains, a transmembrane region, and a cytosolic tail. The extracellular domain mediates homophilic interaction with PECAM-1 molecules in neighboring cells, which particularly occurs through certain epitopes of the more membrane-distal Ig domain. Monoclonal antibodies (Abs), which bind to these domains, and their effect on PECAM-1 homophilic interaction, are shown. Nanocarriers targeted to different PECAM-1 epitopes using these Abs showed differential binding to endothelial cells, endocytic uptake by the cells, and intracellular transport to lysosomes. N/D, not determined; N/A, not applicable.

of their corresponding free counterparts: mAb62/NC showed the highest binding, followed by mAb37/NC, mAb35/NCs, and mAbGi34/NC, whereas mAb4G6/NC did not bind to endothelial cells, despite the effective binding of the corresponding antibody [55]. This could reflect low accessibility of the most membrane-proximal PECAM-1 domain to the carriers. However, among anti-PECAM/nanocarriers that effectively bound to endothelial cells, mAb35/ NC, mAb62/NC, and mAbGi34/NC were quickly internalized, whereas mAb37/NC was internalized rather slowly and incompletely [55]. Following internalization, the majority of mAb62/NC was transported to lysosomes within 3 h after transient and rapid passage through endosomes. In contrast, a small fraction of the internalized mAb35/NC arrived to the lysosomes at this time, whereas a major fraction resided in prelysosomal compartments for prolonged periods of time [55]. Therefore the PECAM-1 epitope recognized by antibody-coated nanocarriers regulates their intracellular uptake and trafficking. There appears to be a high degree of specificity in this regulation, since the epitopes for mAb37, mAb62, and mAb35, providing three distinct patterns of nanocarrier uptake and trafficking, are located in the same domain of PECAM-1. The exact mechanisms of differential subcellular addressing attained by anchoring to distinct PECAM epitopes are not known. Importantly, mAb62, mAb35, and mAb37, respectively, inhibit, activate, and have no effect on PECAM-1 homophilic interaction, which indicates that engaging these epitopes within the same domain causes distinct functional responses [92]. Of note, five monoclonal antibodies to different epitopes on rat angiotensin-converting enzyme (ACE, explored as a target for drug delivery to the pulmonary endothelium)

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have been studied [96]. In that study, efficacy of endothelial binding and potency of ACE inhibition were distinctly epitope dependent [96, 97], yet the internalization was similar for all tested ACE antibodies [96]. However, endothelial cells internalize naked anti-ACE [98], in contrast with antiPECAM that does not enter cells unless it is coupled to a multivalent carrier [52]. It is logical to postulate that the role of epitopes in regulation of intracellular delivery is dependent on the target molecule, cell type and status, and features of a drug carrier. At the present time, we lack sufficient knowledge of mechanisms regulating cellular uptake and trafficking to make predictions on the fate of carriers targeted to specific epitopes. Therefore determination of differential binding, internalization, intracellular transport, and subcellular destination offered by selected epitopes represents a powerful but empirical means for optimization of intracellular drug delivery.

24.5.2 Control of Internalization and Intracellular Trafficking by Carrier Geometry There are, however, elements of design of a drug delivery system that can be rationally modified in order to optimize subcellular targeting. Some of these elements, including the valency of binding moieties, decoration with membrane-permeating and organelle-specific leading peptides, have been discussed above. In this section we consider the role of the carrier geometry (i.e., size and shape) in intracellular delivery. Intuitively, one can expect that the size and shape of a drug carrier can modulate not only its circulation in the blood and distribution in organs, but also cellular uptake, intracellular transport, and fate. Previous studies in cell cultures lend indirect support to this notion. For example, macrophages internalized IgG-coated spheres with diameters of 0.2 versus 2 μm with similar kinetics, but via different pathways (clathrin endocytosis vs. phagocytosis, respectively) and, as a result, micron particles traffic to lysosomes faster than smaller counterparts [34]. The shape of micron sized polystyrene particles also modulates the rate of phagocytosis by macrophages [99]. Of note, uptake of IgG-coated particles by macrophages occurs via immune receptors (quite different in functions from conventional cell surface proteins used as targets) and represents rather an obstacle for most drug and gene delivery systems. In contrast to macrophages, parenchymal cells internalize <100-nm particles faster than >500-nm carriers [100], unless the carriers are not rigid but rather represent an elastic carrier shell or capsule [101]. However, the role of carrier geometry in uptake and trafficking by nonphagocytic cells has not been systematically addressed until recent studies, prompted by development of polymer carriers with diverse geometries (spheres, disks, filaments, and tubes of diverse sizes), which naturally ignited an acute interest in the role of carrier geometry in drug delivery [102–104]. (See Figure 24.4.) For example, internalization of model nonspherical carriers has been investigated in diverse cell types. For instance, short single wall nanotubes (SWNT,

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Elliptical disks 0.1x1x3 μm

Spherical carriers 1-10 μm 0.1 μm Rapid uptake Rapid uptake

Delayed uptake

Partial transport to lysosomes

Effective transport to lysosomes

Partial transport to lysosomes CAM-ENDOCYTOSIS IN ENDOTHELIAL CELLS

Figure 24.4. Controlled subcellular destination of anti-CAM carriers by particle geometry. All model carriers targeted to ICAM-1, including polystyrene submicrometer spheres, micrometer size-range spheres, and elliptical disks coated by surface absorption with anti-ICAM, entered endothelial cells via CAM-mediated endocytosis. The kinetics of uptake by the cells was similar for anti-ICAM spherical carriers, regardless of their size, but this was reduced for anti-ICAM disks, indicating that shape rather than size governs CAM-mediated endocytosis. Contrary, only submicrometer sizes anti-ICAM carriers were efficiently transported to intracellular lysosomes, whereas micrometer size-range anti-ICAM carriers were retained within prelysosomal compartments, independent of their shape, indicating that size rather than shape governs intracellular transport mediated by the CAM pathway.

that have dimensions of ∼5–20 nm diameter and ∼200 nm length) coated with BSA or DNA are internalized by cancer cells (immortalized HeLa cell line), via a pathway that apparently involves predominantly clathrin-coated pits [105]. Furthermore, cells appear to excrete internalized DNA-coated SWNT via exocytosis with similar kinetics as endocytosis [106]. HeLA cells also appear to internalize PEG-decorated cylindrical polymer particles with dimensions of 1 μm in diameter and ∼0.7 μm in height via clathrin-mediated endocytosis and macropinocytosis in HeLa cells [102]. Filomicelles produced from PEG-containing diblock copolymers (diameter ∼50 nm, length 10–20 μm) appear to undergo macropinocytosis in pulmonary epithelial cells, followed by rapid supercoiling and fragmentation, before trafficking of the resultant fragments to the perinuclear region [107]. These results support the notion that there are potentially many mechanisms for internalization and intracellular trafficking of drug carriers controlled by their geometry, both common and specific for diverse cell types and target molecules. However, most of the

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studies on the role of carrier geometry in the uptake and subsequent intracellular trafficking employed nontargeted devices that relied predominantly on nonspecific mechanisms of binding (e.g., electrostatic or hydrophobic adhesion) and uptake (i.e., fluid-phase uptake or passive entrapment into endocytic vesicles of materials adhered to the plasmalemma). In this context, CAM-mediated endocytosis in endothelial cells represents an especially interesting paradigm for analysis of the role of geometry in subcellular addressing of targeted carriers. As we have noted above, endothelial cells do not internalize monoclonal anti-PECAM and anti-ICAM, but very effectively internalize multivalent protein anti-PECAM conjugates within the size range (100–500 nm), although the rate of the endocytic uptake dramatically decreases if size of polymorphous protein conjugates of the same content exceeds this range [63]. However, this restriction does not apply to polymer carriers of spherical and elliptical shape: endothelial cells in vitro and in vivo effectively and safely internalize micron sized carriers targeted to ICAM-1 and PECAM-1 [68, 70]. Interestingly, the selection of epitopes engaged and “stimulated” by anti-PECAM/nanocarriers seems to play a more important role in determining the efficacy of internalization than carrier size: spherical particles within the size range from 0.1 to 5 μm carrying mAb62, mAb35, and mAbGi34 were similarly internalized by endothelial cells [70]. We have recently found that prototype anti-ICAM carriers of diverse shape (spheres vs. elliptical disks) and size (mean diameters from 0.1 to 10 μm) are internalized by endothelial cells in culture and pulmonary endothelial cells after vascular injection in vivo [68]. Therefore delivery via CAM-mediated endocytosis differs from that via classical endocytic pathways restricted to much smaller objects. For example, ∼1-μm phage particles expressing a peptide with high affinity to a lung endothelial, caveolae-specific protein neither bind to endothelium nor enter caveoli in vivo due to the size restriction (<100 nm) of the caveolar neck [108]. Studies with both anti-ICAM and antiPECAM carriers confirmed extraordinary internalization capacity of CAM-mediated endocytosis for spherical carriers with diameter up to several microns [68, 70]. However, the shape of anti-ICAM/carriers influenced their internalization. At the cellular level, spheres seem to be the most permissive shape for endocytosis of anti-ICAM carriers ranging from 0.1 to 5 μm, whereas uptake of anti-ICAM elliptical disks is delayed [68]. Slow uptake of anti-ICAM disks may be due to higher surface-to-volume ratio of disks versus spheres, given that this parameter dictates the need for higher plasmalemma deformability to adapt to disks, which have more pronounced and varying curvature. A similar phenomenon was recently observed for phagocytosis of IgG-opsonized particles by macrophages, which was governed by formation and progression of actin cups beneath the plasma membrane [99]. However, in contrast with phagocyte uptake of IgG-opsonized particles, anti-ICAM carriers did not elicit actin polymerization into cups around spherical or disk particles, regardless of anti-ICAM carrier geometry and the progression of internalization.

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Instead, anti-ICAM carriers induced formation of actin stress fibers [68, 69, 72] and their endocytosis was inhibited by the actin filament inhibitor latrunculin A, but not by cytochalasin D that typically affects classical actindependent endocytosis. A similar reorganization of the actin cytoskeleton has previously been observed for 0.1–0.2 μm diameter anti-ICAM spheres, which is mediated by RhoA activation and ROCK signaling [55, 69]. This and results of probing the endocytic pathways with preferential pharmacological inhibitors indicate that endocytosis of anti-ICAM carriers with different geometries operates via a similar, nonclassical uptake pathway, namely, CAMmediated endocytosis [68, 69]. In contrast to endocytosis, mainly ruled by carrier shape, intracellular transport of internalized anti-ICAM carriers to lysosomes is controlled by carrier size. Submicron sized anti-ICAM carriers show more efficient transport to lysosomes, whereas micron sized anti-ICAM spheres or disks reside within prelysosomal compartments for longer periods of time [68]. This is unlike lysosomal trafficking of IgG-opsonized particles by macrophages, which increases with increasing particle size [34]. Perhaps less effective lysosomal trafficking of micron sized anti-ICAM carriers by ECs may reflect more restrictive dimensions of intracellular vesicular machinery within this cell type. Endosomal versus lysosomal addressing of anti-ICAM carriers, regulated by modulating carrier size, highlights the variability and potential utility of this paradigm for therapeutic applications that require corresponding intracellular localization. As an example with relevant implications in the design of antioxidant therapies, utilization of micron sized anti-ICAM carriers delivering catalase provided prolonged antioxidant protection by catalase residing in endosomes [68]. As a distinct example relevant to the treatment of lysosomal storage disorders, utilization of submicron sized anti-ICAM carriers delivering recombinant ASM into endothelial lysosomes optimized targeted attenuation of sphingomyelin storage condition [68]. Therefore, rational design of drug delivery nanodevices, including control of such engineered parameters as their size and shape, helps to guide the uptake and trafficking in a way that optimizes the therapeutic effects of the targeted drugs.

24.6

CONCLUSION

The means providing precise control of subcellular destination of drug delivery in the selected target cells remain an elusive goal, the Holy Grail of modern biomedicine. It is tempting to speculate that this goal can be achieved via rational design of drug carriers, for example, by modulating such controllable parameters as the carrier size, shape, charge, elastic properties, rate, and pH dependence of degradation and deployment of auxiliary tools guiding the intracellular uptake, vesicular transport, release, and subsequent navigation through the cytosol and organelles. Some of these design parameters have

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been discussed in this and other chapters of the book, whereas some remain to be uncovered and employed. On the other hand, our understanding of the cellular and molecular mechanisms that govern uptake, turnover, and intracellular trafficking of diverse target molecules, expressed on the cellular surface and their natural and artificial ligands, continues to excel. Although still grossly incomplete, this knowledge helps us to select particular cellular determinants providing uptake and cellular transport of the anchored ligands via specific endocytic and trafficking pathways. Rational selection of such molecular target determinants for drug delivery will both provide specific recognition of cell types and phenotypes of interest and help to address the drug carriers and their cargoes to selected subcellular localization. Of course, the fate of a carrier or a drug, even anchored to the same target molecule, may vary in different cells (e.g., macrophages vs. endothelial cells) and even in different phenotypes of the same cell type (e.g., quiescent vs. pathologically activated endothelial cells). Remarkably, even anchoring to distinct epitopes localized in the same extracellular domain of a target molecule can provide variable subcellular delivery of the same drug carrier. This intriguing paradigm has been illustrated in this chapter by the case of PECAM-1. Conceivably, combining precision of subcellular targeting attainable by modulation of the carrier design parameters with that provided by optimal selection of the anchoring epitope on target molecules holds a promise for unprecedented versatility and accuracy of subcellular addressing of drugs. In this context, it seems plausible to postulate that certain target molecules may serve a variety of delivery purposes. For example, selection of target epitopes combined with modulating the carrier geometry and valency of anchoring on these epitopes provides a versatile framework for optimization of subcellular addressing of diverse therapeutic enzymes. This principle is illustrated in this chapter by surface anchoring of monovalent antithrombotic drugs on the endothelial luminal surface [54] and variably optimized intracellular delivery of antioxidant and lysosomal enzymes [68], all targeted to the same molecules, CAMs. It is tempting to believe that such a versatile portfolio of addresses is not unique for this particular endothelial target molecule. No doubt, ongoing and future studies will uncover similarly versatile outcomes of anchoring given drug delivery systems to a variety of cell surface molecules that can be exploited for precise subcellular drug delivery in diverse cell types.

ACKNOWLEDGMENTS This work was supported by Pennsylvania NTI core project, PENN TAPITMAT grant, grants from NHLBI RO1 HL087036, PO1 HL079063, and HL73940 to V.M., and by Nano-Biotechnology DEBD, UMCP Minta Martin, and AHA 09BGI2450014 grants to S.M.

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CHAPTER 25

Uptake Pathways Dependent Intracellular Trafficking of DNA Carrying Nanodelivery Systems IKRAMY A. KHALIL, YUMA YAMADA, HIDETAKA AKITA, and HIDEYOSHI HARASHIMA Faculty of Pharmaceutical Sciences, Hokkaido University, Hokkaido, Japan and CREST, Japan Science and Technology Agency (JST), Japan

25.1

INTRODUCTION

Gene therapy is a promising approach for the treatment of a number of inherited and acquired diseases that cannot be treated effectively by conventional medicine [1–4]. The approach relies mainly on the production of therapeutic proteins within cells and requires the efficient delivery of nucleic acids to intracellular target sites. The simplest form of gene therapy involves the delivery of DNA to the nucleus and its subsequent transcription and translation, leading to the production of therapeutic proteins. Although simple in theory, gene therapy is more complicated in practice [3]. DNA and other nucleic acids are generally large, charged molecules and hence they do not have the ability to reach their intracellular target sites on their own. An appropriate gene delivery system is thus required to deliver nucleic acids to their intracellular targets. Current gene delivery systems can be divided into viral and nonviral vectors. Viral vectors are generally more efficient since they utilize the natural machinery associated with a viral infection to deliver therapeutic genes [5]. However, they suffer from major drawbacks, particularly related to their high toxicity and immunogenicity in addition to the difficulties associated in their bulk production. Nonviral vectors are attractive alternatives to viral vectors owing to their relatively low toxicity and immunogenicity and their ease of production. However, they are relatively less efficient than viral vectors [6–9]. Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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Although gene therapy is highly promising as a possible future medical treatment for several diseases, clinical success continues to be far below theoretical expectations [3]. As explained above, this is mainly due to the poor toxicological profile of the efficient viral vectors and the inefficient delivery of nucleic acids to their intracellular target sites of action in the case of nonviral vectors [10–13]. Efforts are ongoing to solve both of these obstacles; however, increasing attention is directed toward improving the performance of nonviral vectors. One strategy for improving nonviral gene delivery is the synthesis of novel materials and compounds that are capable of efficiently delivering genes to intracellular sites of action. Another strategy involves studies of the interactions of vectors with cellular components and developing strategies for overcoming different biological barriers [12, 13]. The problem is more complex than simply how therapeutic genes can pass through plasma membranes. Recent studies have shown that nonviral mediated cellular internalization is actually an efficient process; however, the level of therapeutic genes produced is still low compared to what would be expected from the amount of material that is internalized [14]. This indicates that intracellular trafficking is still inadequate and efforts should be directed to methods for improving it. Therefore it is important to identify the uptake pathways of DNA carrying systems and, more importantly, to understand the intracellular barriers that limit the efficient delivery of DNA to intracellular target sites. In this chapter, we briefly introduce the different uptake pathways involved in the internalization of macromolecules in general and discuss the uptake pathways dependent intracellular trafficking of common classes of nonviral DNA carrying nanosystems. The endosomal escape of internalized genes is discussed in more detail, since this is currently a major obstacle to gene delivery. We also discuss intracellular organelle targeting, particularly mitochondrial targeting and DNA nuclear transfer. We focus on our own efforts to rationally design improved DNA carrying systems based on our studies of their intracellular trafficking. A novel multifunctional envelope-type nanodevice (MEND) developed by us, in an attempt to overcome several of the biological barriers to efficient gene delivery to intracellular targets, is reviewed and future directions for further development are also discussed.

25.2

CELLULAR UPTAKE PATHWAYS OF MACROMOLECULES

Before we discuss the different uptake pathways and their related intracellular trafficking of DNA carrying nanodelivery systems, we briefly introduce the cellular uptake pathways of macromolecules in general. The plasma membrane of a living cell is a dynamic structure that is relatively lipophilic in nature. As a result, it restricts the entry of large, hydrophilic or charged molecules. Endocytosis (the vesicular uptake of extracellular macromolecules) has been established as the main mechanism for the internalization of

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macromolecules into cells [15–17]. However, endocytosis is not the sole entry port for macromolecules. A class of cationic peptides that contain protein transduction domains (PTDs), such as the TAT, penetratin, and VP22 peptides, may have the ability to be taken up by cells without endocytosis [18–20]. It was initially suggested that these peptides are able to penetrate cell membranes directly by an energy-independent route [21–24]. A mechanism involving the direct penetration of the lipid bilayer caused by the localized positive charge of the peptide was proposed to explain this uptake [21]. However, we have previously shown that the mechanism of uptake of an octaarginine peptide, a prototype of the PTDs, was dramatically changed by N-terminal stearylation as well as by complexation with DNA [25]. This raised a serious issue concerning the ability of a peptide to retain its activity after modification, conjugation, or complexation with other molecules. Nevertheless, based on a recent reevaluation of the uptake of these peptides and their cargoes, the evidence suggests that endocytosis is the major uptake pathway [26–30]. The possibility of the endocytosis-independent uptake of these peptides and their cargoes, however, cannot be fully excluded [31]. In addition to this mysterious PTD-mediated nonendocytic delivery, several nonendocytic internalization pathways have been described, including microinjection, cell permeabilization, and electroporation [32–34]. However, these techniques are highly invasive and cannot be used for in vivo gene delivery. Therefore the focus of this discussion is mainly on well-characterized endocytic uptake pathways. Endocytosis refers to the cellular uptake of macromolecules and solutes into membrane-bound vesicles derived by the invagination and pinching off of pieces of the plasma membrane [16]. Kinetically, three modes of endocytosis can be defined: fluid-phase, adsorptive, and receptor-mediated endocytosis [16]. Endocytosis can also be classified into two broad categories—phagocytosis or cell eating (the uptake of large particles) and pinocytosis or cell drinking (the uptake of fluids and solutes) [16]. At least three morphologically distinct pinocytic pathways have been characterized: clathrinmediated endocytosis, caveolae, and macropinocytosis (Figure 25.1). They differ in the composition of the coat (if any), in the size of the detached vesicles, and in the fate of the internalized particles. 25.2.1 Clathrin-Mediated Endocytosis Clathrin-mediated endocytosis (CME) is the major and most well-characterized endocytic pathway [17, 35]. CME occurs constitutively in all mammalian cells and carries out the continuous uptake of essential nutrients, antigens, growth factors, and pathogens. The most common examples of molecules that are internalized by CME are the cholesterol-laden low-density lipoprotein (LDL) that binds to LDL receptors, and iron-laden transferrin (Tf), which binds to Tf receptors [36, 37]. Generally, the first step of internalization via CME is the strong binding of a ligand to a specific cell surface receptor. This

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Clathrin-mediated Endocytosis Dynamin

Ligand

Receptor Clathrin-coated pit

Caveolae

Macropinocytosis

Caveolin

Clathrin coat

Recycling Clathrin-coated vesicle (CCV)

H+ H+

Caveosome ?

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Leakiness H+

Late endosome (Low pH) Endosomal release Nuclear delivery

Cytosolic release H+

Lysosome

Degradation

Ruffling

Actin Recycling

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Nucleus

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Figure 25.1. Different endocytic pathways. Clathrin-mediated endocytosis (CME) is the major and best-characterized endocytic pathway. Caveolae and macropinocytosis are two clathrin-independent endocytic pathways. (See color insert.)

results in the clustering of the ligand–receptor complexes in coated pits on the plasma membrane, which are formed by the assembly of cytosolic coat proteins, the main assembly units of which are clathrin, which forms a polygonal lattice in the surface of the membrane (Figure 25.1). The coated pits then invaginate and pinch off from the plasma membrane to form intracellular clathrin-coated vesicles (CCVs). CCVs carry concentrated receptor–ligand complexes into cells. They range in size from ∼100 to 150 nm in diameter and are characterized by the presence of a polygonal clathrin coat. The clathrin coat then undergoes depolymerization, resulting in early endosomes, which fuse with each other or with other preexisting endosomes to form late endosomes that further fuse with lysosomes. Molecules entering via this pathway rapidly experience a drop in pH from neutral to pH 5.9–6.0 in the lumen of early endosomes, with a further reduction to pH 5 during the progression from late endosomes to lysosomes [38]. Within the endosomes, ligands and receptors are sorted and are sent to their appropriate cellular destinations, such as lysosomes, the Golgi apparatus, the nucleus, or the cell surface membrane. Macromolecules that are internalized through CME are usually trapped in endosomes followed by enzymatic degradation in lysosomes with little or almost no access to intracellular target sites. Actually, entrapment and degradation can be regarded as two separate barriers, since preventing lysosomal degradation results in the accumulation of internalized molecules in intracellular vesicles without enhancing cytosolic release.

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25.2.2 Caveolae-Mediated Endocytosis Caveolae are small, hydrophobic membrane microdomains that are rich in cholesterol and glycosphingolipids [39, 40]. Classically, caveolae were defined as flask-shaped invaginations of the plasma membrane, but they can also exist as flat, tubular, or detached vesicles. Caveolae are present in many cell types and are especially abundant in endothelial cells. They are involved in a number of cellular processes, including cholesterol homeostasis and glycosphingolipid transport. Caveolae are also involved in transcytosis and endocytosis of certain viruses such as the simian virus 40 (SV40), as well as some bacteria and bacterial toxins (e.g., cholera toxin). Caveolae are characterized by their association with a family of cholesterol-binding proteins called caveolins, which function to create and/or mediate these structures. The mechanisms of caveolar internalization were elucidated by visualizing the trafficking of the SV40 that utilizes caveolae to gain entry into the cells [41] (Figure 25.1). SV40 initially associates with the cell membrane and then becomes trapped in relatively stationary caveolae. The subsequent uptake of the virus leads to its delivery to intracellular organelles that are distinct from classical Tf-labeled endosomes. The presence of caveolin in these organelles gave rise to the name “caveosome.” SV40 then becomes segregated from caveolin and is sorted out of caveosomes for delivery to the endoplasmic reticulum (ER). In general, caveolae (∼50–60 nm) are highly stable and are only slowly internalized, in contrast to the rapid and dynamic nature of Tf-labeled endosomes [15]. Another major difference is that caveolar uptake is a nonacidic and nondigestive route of internalization [42]. Caveolae do not suffer a drop in pH and most pathogens that are internalized by caveolae can be transported directly to the Golgi and/or ER, thus avoiding normal lysosomal degradation [39, 42]. It is generally believed that caveolar uptake does not lead to lysosomal degradation. However, caveolae are slowly internalized, are small in size, and their fluid-phase volume is low. Thus it is unlikely that they contribute significantly to constitutive endocytosis, although the situation is different in endothelial cells in which caveolae constitute 10–20% of the cell surface [15].

25.2.3 Macropinocytosis Macropinocytosis refers to the formation of large endocytic vesicles that are irregular in size and shape and are generated by actin-driven envagination of the plasma membrane [16, 43, 44]. Macropinocytosis is usually accompanied by cell surface ruffling that is induced in many cell types upon stimulation by growth factors or other signals. A ruffle is formed by a linear band of outwarddirected actin polymerization near the plasma membrane, which lengthens into a planar extension of the cell surface. After stimulation by a mitogenic factor, the ruffles become longer and broader, and frequently close into large macropinosomes (Figure 25.1). Macropinosomes have no coat and do not concentrate receptors. They vary in size, sometimes reaching diameters as

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large as 5 μm. Because the macropinosomes are relatively large, macropinocytosis is an efficient route for the nonselective endocytosis of solute macromolecules [15]. Macropinocytosis fulfills diverse functions, especially when massive fluid-phase endocytosis is necessary. This route facilitates the bulk uptake of soluble antigens by immature dendritic cells and some pathogens trigger macropinocytosis to facilitate their own uptake [15]. After the formation of macropinosomes, these vesicles lose their F-actin and their intracellular fate differs, depending on the cell type [44, 45]. In macrophages, they move toward the center of the cell, shrink by loss of water, become acidified, and then completely merge into the lysosomal compartment [45]. In human A431 cells, aside from macropinosomes, they do not interact with other endocytic compartments. They constitute a distinct vesicle population, which eventually recycles most of their contents back to the cell surface [44]. Although the pH of macropinosomes decreases, they are less directed to lysosomes than CCVs. Macropinosomes are thought to be inherently leaky vesicles compared to other types of endosomes [45, 46]. This pathway provides some advantageous aspects such as the increased uptake of macromolecules, the avoidance of lysosomal degradation, and the ease of escape from macropinosomes due to their relatively leaky nature.

25.3 INVOLVEMENT OF ENDOCYTIC PATHWAYS IN CELLULAR ENTRY OF DNA CARRYING NANODELIVERY SYSTEMS The current status of our knowledge regarding the contribution of each endocytic pathway to the cellular uptake of common classes of DNA carrying nanodelivery systems and how these systems are processed by cells to achieve transgene expression is discussed in this section. We focus mainly on four typical DNA carrying nanosystems: cationic lipids, cationic polymers, ligandbased, and peptide-based systems. Both cationic lipids and polymers form complexes upon mixing with DNA. Cationic lipid/DNA complexes are denoted as “lipoplexes” while cationic polymer/DNA complexes are denoted as “polyplexes.” Both systems can be further modified with targeting ligands and specific peptides to improve their performance and increase their specificity to certain cell populations. 25.3.1 Lipoplexes The first step in the internalization of DNA carrying systems is cell binding. Unless a specific targeting ligand is incorporated into the system, the binding of lipoplexes (and also polyplexes) to the cell surface is the result of a nonspecific ionic interaction between positively charged complexes and a negatively charged cell surface. Negatively charged cell surface constituents, which include heparan sulfate proteoglycans (HSPGs) and integrins, play a role in the cellular binding of positively charged lipoplexes [47]. For example, the

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cellular binding of lipoplexes in proteoglycan-deficient mutant cells is reduced [47]. The presence of soluble heparin and heparan sulfate in the medium competitively inhibits binding. At this point, HSPGs may act as nonspecific receptors for cationic macromolecules, but their exact role in mediating cellular uptake is currently unclear. Some evidence exists to show that the transmembrane proteins, syndecans, may cluster to form focal points at the plasma membrane during binding to cationic particles and this clustering induces their interaction with the actin cytoskeleton, probably resulting in the formation of tension fibers. This tension provides the energy required for the particles to be engulfed [48]. Regarding the mechanism by which lipoplexes are internalized, early reports suggested that fusion between the lipids and the plasma membrane is responsible for delivering DNA directly to the cytosol [49, 50]. It was proposed that the interaction between the liposomes and DNA or the cell membrane destabilizes the liposomes, thus facilitating their fusion with each other and with other membranes. However, most of the following experimental evidence supports the involvement of endocytosis as the main entrance route. For example, the use of endocytosis inhibitors significantly reduces gene expression. Furthermore, interference of the endocytic pathway with lysosomotropic reagents such as chloroquine was found to enhance gene expression. The strongest evidence arises from electron microscopy imaging of gold-labeled DNA, which clearly shows the presence of DNA in intracellular vesicles, typical of entry via endocytosis [51]. In general, it is currently believed that membrane fusion is important for transfection but that most of the uptake occurs through endocytosis. Membrane fusion occurs as a result of endosome acidification and is responsible for releasing the contents of an endosome to the cytosol. Almofti et al. [52] proposed that the uptake of lipoplexes occurs by endocytosis, but that the majority of membrane fusion (72%) occurs at the plasma membrane level and is essential for endocytosis to proceed. Assuming that endocytosis is the main entry route for lipoplexes, the question then is which pathway of endocytosis is responsible for uptake. The available data show diverse results and the mechanism appears to be dependent on the cationic components used as well as on the cell type. Rejman et al. [53] reported that the uptake of lipoplexes formed between a cationic lipid (DOTAP) and DNA is inhibited by chloropromazine and potassium depletion but is unaffected by filipin and genestein, suggesting that uptake occurs solely by CME. Furthermore, they showed that particles that are internalized by CME are eventually degraded in lysosomes. In an earlier study, Zhou et al. [54] proposed that the uptake of lipopoly-l-lysin (LPLL) lipoplexes occurs through CME, since the presence of the actin-depolymerizing reagent cytochalasin B increased transfection activities. In contrast to Rejman et al. [53] they suggested that CME is the most productive pathway for internalization. Zuhorn et al. [55] showed that lipoplex-mediated transfection occurs through cholesterol-dependent CME. Cholesterol depletion by treatment with methylβ-cyclodextrin decreased the activities of SAINT-2/DOPE lipoplexes. This is

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largely indicative of nonclathrin endocytosis, however, inhibitors of caveolae such as filipin and cytochalasin D had only a slight effect on internalization. Colocalization with Tf (CME marker) and inhibition by potassium depletion further confirmed the involvement of CME. In contrast to these reports, Matsui et al. [56], using poorly differentiated airway epithelial cells, proposed that the uptake of LipofectACE lipoplexes occurs through phagocytosis. Similar to the uptake of large (2-μm) microspheres, which were used as markers for phagocytosis, they found that the uptake of lipoplexes was inhibited by cytochalasin B as well as by potassium depletion. 25.3.2 Polyplexes The uptake of polyplexes also occurs through endocytosis, but without fusion with the cell membrane. It is generally believed that the uptake of poly-llysine (PLL) and polyethylene imine (PEI) complexes occurs through CME. Goncalves et al. [57] showed that the uptake of histidylated poly-l-lysin (His-pLK) occurs through both clathrin-dependent and clathrin–independent pathways, the latter being mostly macropinocytosis, since it was inhibited by amiloride and stimulated by phorbol esters. Furthermore, they found that macropinocytosis of the polyplexes and the recycling of DNA impaired the transfection and concluded that CME is the most productive pathway. Rejman et al. [53] also proposed that the uptake of PEI-polyplexes occurs through both clathrin-dependent and clathrin–independent pathways, but they favored the latter mechanism, which involves the caveolae, since it was inhibited by filipin and genestein. Another difference arises regarding their finding that CME is less productive since caveolar internalization does not proceed via lysosomes, thus leading to efficient transfection. The diversity of results suggests that a variety of factors may affect the actual mechanism. 25.3.3 Ligand-Mediated Internalization Receptor-mediated DNA delivery is an attractive feature since it allows the cell-specific delivery of DNA. A ligand is attached to the DNA delivery system so as to be recognized by particular receptors on the cell surface to induce receptor-mediated endocytosis (RME). For example, hepatocytes exclusively express large numbers of high-affinity cell surface receptors that bind to and subsequently internalize asialoglycoproteins [58]. Introducing a galactose moiety into a gene delivery system can achieve liver-parenchymal-cell specific gene transfection. Mannose receptor-mediated gene transfection is another approach for targeting macrophages that overexpress mannose receptors on their surface [59]. Transferrin (Tf), an iron-binding glycoprotein, has been used as a tumor-targeting ligand in gene delivery systems [37, 60]. T freceptors are overexpressed in rapidly dividing cells due to the increased cellular need for iron. The folate receptor is another example of receptors that are overexpressed in tumor cells and it can be used for tumor targeting [61].

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Most DNA systems carrying a targeting ligand to specific receptors are internalized by CME. For example, a DNA vector carrying the targeting ligand Arg-Gly-Asp (RGD) was shown to be internalized by CME. This is evident from electron microscopy (EM) images showing colocalization with clathrin-coated pits in addition to the inhibitory effects of CME inhibitors such as potassium depletion and hypertonic treatment [62]. The uptake was not inhibited by cytochalasin B, which blocks noncoated pit formation. Similar results showing CME appeared using Tf-modified DNA delivery systems [63]. However, other pinocytic pathways are capable of selective RME events [64]. For example, the internalization of interleukin-2 (IL-2) into lymphocytes was partially inhibited by treatments that disrupt CME, suggesting that a clathrinindependent mechanism contributes significantly to the efficient internalization of IL-2 receptors [65]. In addition, Tf-targeted lipoplexes were shown to be internalized through nonclathrin-dependent endocytosis, probably due to their large size, which cannot be internalized via CME [66]. Although RME is a promising approach for drug targeting, most of the currently used ligands are internalized by CME, which is highly linked to lysosomal degradation, leading to limited transfection activities for these systems. Therefore functional devices that increase the cytosolic delivery of genes are needed. For example, in a previous study, we reported that the intracellular fate of T fliposomes was improved by adding GALA, a pHsensitive fusogenic peptide, which enhances endosomal escape in response to the low pH in endosomes [60]. In addition, we also showed that Tf-modified DNA carrying nanoparticles did not yield high transfection activities unless the fusogenic peptide GALA is used [67]. Exploring and targeting new receptors that can be internalized by clathrin-independent endocytosis are likely to provide more efficient systems because these uptake mechanisms are relatively less affected by lysosomal degradation. 25.3.4 Peptide-Mediated Internalization Cationic peptides such as arginine-rich peptides (RRPs) were previously used to improve gene delivery through different strategies [18]. The direct complexation of TAT or R8 peptides to DNA was shown to produce complexes that are capable of transfection [68]. The addition of hydrophobic moieties to short RRPs was shown to further enhance the activities [68]. The TAT peptide facilitated gene transfer in combination with cationic liposomes and the activities of Lipofectin lipoplexes could be further improved by simply mixing the liposomes and DNA with the TAT peptide [69]. Torchilin et al. [70] reported an efficient gene delivery in vitro and in vivo with reduced cytotoxicity mediated by complexes of TAT liposomes and DNA. In a previous study, we described the development of R8-modified liposomes (R8-Lip) for use as an efficient drug delivery system [71]. The N-terminal amino acid residue of the R8 was attached to a stearyl group to form stearylated-R8 (STR-R8). The STR-R8 peptide was mixed with other lipids

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during the formation of the liposomes. The hydrophobic stearyl group would be expected to become anchored on the lipid bilayer of liposomes, thus leaving the R8 exposed on the surface. The modification of liposomes with low or high densities of the R8 peptide dramatically enhanced the cellular internalization of a model drug encapsulated in the liposomal aqueous phase compared to the use of the free drug or when the drug was encapsulated in liposomes lacking the R8 peptide [71, 72]. When double-labeled liposomes were used, high intracellular fluorescence was observed and both liposomal labels (aqueous and lipid markers) were colocalized in the cytosol of cells. The intracellular colocalization of the markers excludes the possibility of fusion between the liposomal membrane and the cell membrane. In addition, the intracellular fluorescence appeared as punctuated signals in the cytosol, indicating the possible contribution of endocytosis, a finding that is further supported by the strong inhibition in uptake after energy depletion [71]. To investigate the potential of the R8-Lip as a DNA carrying system, we adopted a technology based on forming a core–shell structure, which resembles an envelope-type virus [73]. The DNA was condensed with a polycation and the complexes were coated with a lipid envelope that had been modified with the R8 peptide. The encapsulation of DNA inside the liposomes is more advantageous than the method involving the direct complexation of the cationic liposomes with DNA as in lipoplexes, since it provides more protection for the DNA and the lipid envelope can control the topology of the functional devices to exert their activities. The DNA is separated from the outer R8 peptide by the lipid envelope. As a result, the peptide remains free and available for interaction with the plasma membrane. R8-Lip with a lipid envelope consisting of egg phosphatidylcholine (EPC) and cholesterol (Chol) efficiently internalized the DNA encapsulated in their cores. However, the internalized R8-Lip modified with a low density of R8 (R8-Lip-LD) did not show any significant gene expression [71]. The reason for this low transfection is inadequate intracellular trafficking rather than a lack of cellular uptake. This was confirmed by the high transfection activities of the same particles in the presence of the endosome disrupting drug chloroquine (unpublished data). This is consistent with our analysis of intracellular trafficking, which showed that the particles are extensively degraded in lysosomes [71]. In contrast, R8-Lip modified with a high density of R8 (R8-LipHD) produced remarkable gene expression, even in the absence of any device for enhancing endosomal destabilization, indicating that the particles are able to escape from endosomes on their own. The difference in gene expression levels between DNA-containing R8-Lip-LD and R8-Lip-HD was about three orders of magnitude, while the difference in internalization was only about sevenfold [71]. This confirms our conclusion that the intracellular fate of the particles is improved by increasing the peptide density. In the presence of chloroquine, the difference in gene expression was around eightfold (unpublished data), which can be explained by the difference in the amount internalized in each case.

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The difference in intracellular trafficking of R8-Lip-LD and R8-Lip-HD can be explained by differences in the internalization mechanism, which confirmed that intracellular trafficking is closely related to the entry pathway [71, 72]. Incubation of cells with R8-Lip-LD or R8-Lip-HD in the presence of a mixture of metabolic inhibitors, which inhibit all types of endocytosis through energy depletion, strongly inhibited the cellular uptake of both types of liposomes. This indicates that the uptake process is highly energy dependent. Moreover, incubation in the presence of a hypertonic medium strongly inhibited the uptake of R8-Lip-LD, whereas it inhibited the uptake of R8-Lipo-HD by about 35%. This indicates that CME is the major uptake pathway for R8Lip-LD, whereas the uptake of R8-Lip-HD involves different pathways as a major entrance route. On the other hand, in the presence of the macropinocytosis inhibitor amiloride, the cellular internalization of R8-Lip-HD was strongly inhibited (approximately 80%), while more than 50% of the R8-LipLD particles were taken up by the cells. These results indicate that only liposomes modified with a high density of R8 peptide use macropinocytosis as the major entrance route, whereas those modified with a low density of R8 mainly use the classical endocytosis. The caveolar inhibitor filipin inhibited the uptake of both types of liposomes only slightly, indicating a minor contribution of caveolae in the uptake. Furthermore, cytochalasin D and nystatin, both of which inhibit macropinocytosis and caveolae, inhibited the uptake of R8-LipHD. Because the presence of the specific caveolae inhibitor filipin caused only a minor inhibition in uptake, these results further confirm the involvement of macropinocytosis in the cellular uptake of R8-Lip-HD. Only R8-Lip-HD was able to stimulate the uptake of the macropinocytosis marker dextran (70 kDa). In addition, nonlabeled R8-Lip-HD increased the internalization of rhodamine-labeled R8-Lip-LD in a concentrationdependent manner and this increase was nearly completely inhibited in the presence of amiloride [71]. This confirms that R8-Lip-HD has the ability to stimulate macropinocytosis and suggests that a certain peptide concentration is required to accomplish this. We next examined the intracellular trafficking of R8-Lip for further optimization, in an attempt to achieve the maximum activity. The intracellular fate of particles internalized via CME or macropinocytosis is an important issue. Particles internalized via CME eventually undergo degradation in lysosomes while macropinosomes do not extensively fuse with lysosomes in most cell types [16]. This would be expected to enhance the survival of the particles inside the cells. In accordance, the intracellular fluorescence in the case of R8-Lip-LD containing a rhodamine aqueous phase was dramatically reduced after a chase time of 6 h. In the case of R8-Lip-HD, a highly punctuated intracellular fluorescence was observed after the same chase time, indicating that the liposomes remained intact during that period [71]. However, using this approach, it is not possible to precisely determine the extent of escape from macropinosomes. The detected fluorescence may represent liposomes in the cytosol or trapped in macropinosomes. In either case, the liposomes would appear to be resistant to degradation. In accordance

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with the hypothesis that macropinocytosis is responsible for the enhanced survival of R8-Lip-HD, the intracellular behavior of R8-Lip-LD was changed when internalization was shifted to macropinocytosis using nonlabeled R8Lip-HD. To confirm the avoidance of lysosomal degradation in the case of macropinocytosis, we examined the colocalization between different R8-Lip preparations and LysoSensor, which stains acidic compartments such as lysosomes. R8-Lip-LD were highly colocalized with lysosomes, supporting the hypothesis that they are degraded in lysosomes. This also indicates that these liposomes do not have the ability to escape efficiently from the endocytic compartment. Meanwhile, R8-Lip-HD was only partially colocalized with lysosomes. The fraction that was not colocalized with lysosomes, the dominant fraction, may represent liposomes either in macropinosomes or those that had escaped to the cytosol. Wadia et al. [46] showed that the TAT-Cre fusion protein could escape from macropinosomes, probably due to the inherently leaky properties of the macropinosomes compared to other intracellular vesicles. The enhanced activity of the fusion protein in the presence of a pHsensitive fusogenic peptide suggests that the particles were present in an acidic compartment and also indicates that the escape from macropinosomes was an inefficient process. In the case of R8-Lip-HD, at least a fraction was found to escape from macropinosomes and migrate toward and bind to the nucleus (unpublished data). Liposomes that are located in close proximity to the nucleus are useful, since they will be internalized more readily when the cell divides. The results presented before indicated that uptake via macropinocytosis is more productive in terms of gene expression than the classical CME since it tends to avoid lysosomal degradation. Consistent with this assumption, blocking the internalization of R8-modified nanoparticles through macropinocytosis caused a ∼95% inhibition in gene expression while blocking CME caused a ∼35% inhibition [71]. These data confirm that uptake via macropinocytosis is the main contributor to efficient gene expression, indicating the significance of macropinocytosis as a potential entrance pathway for improving transfection. Based on our studies of the uptake-dependent intracellular trafficking of R8-Lip, we prepared an artificial virus system that we refer to as a multifunctional envelope-type nanodevice (MEND) [73, 74]. The transfection activity of R8-MEND was increased in the presence of dioleoylphosphatidyl ethanolamine (DOPE), indicating that devices that can enhance endosomal escape appear to be essential for achieving high transfection activity [74]. The use of the optimized R8-MEND system with an efficient combination of R8 and DOPE resulted in high transfection activities in vitro. The activities were comparable to adenovirus, and were superior to the frequently used lipofection reagent, Lipofectin [72, 74]. In addition, the system produced minimum cytotoxicity and it was found to be safer than both adenovirus and the Lipofectamine Plus reagent as judged by measuring the protein content and by an MTT assay of cell viability [74].

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We recently introduced liposomes modified with a novel peptide sequence (IRQRRRR) (IRQ-Lip) [75]. This novel peptide sequence was identified with an in vivo phage display to target the skeletal muscles. The peptide induced liposome internalization through a novel pathway, caveolar endocytosis, in parallel with CME. The uptake of IRQ-Lip was inhibited by hypertonic treatment (56%) and filipin (60%) while it was not affected by the macropinocytosis inhibitor amiloride. Compared to R8, which directs the liposome entry mainly via macropinocytosis, the IRQ peptide is similarly rich in arginine, albeit it contains fewer residues than R8. It is interesting to note that substituting only two arginine residues in R8 with isoleucine and glutamine dramatically changed the uptake mechanism. Five arginine residues in the IRQ peptide sequence may remain available to interact with HSPGs; however, the interaction may not be sufficiently strong to induce the cytoskeleton rearrangement required for macropinocytosis stimulation. These results suggest that there is a specific sequence in IRQ that is recognized by receptor molecules in caveolae. The other part of the sequence may be responsible for uptake via CME. As a gene delivery system, optimized IRQ-modified MEND encapsulating siRNA produced 52% gene silencing in cell cultures [75]. The mechanism of endosomal escape of IRQ-modified systems will be discussed below.

25.4 INTRACELLULAR TRAFFICKING OF DNA CARRYING NANODELIVERY SYSTEMS Considering that the major part of DNA carrying systems are internalized through endocytosis, the internalized DNA exists in the endosome with no access to the cytosol or the nucleus. These endosomes either fuse with lysosomes for degradation or their contents are recycled back to the cell surface. Therefore escape from endosomes is essential for efficient transfection. Most DNA carrying systems contain a specific machinery to allow escape from endosomes. Once released from endosomes, the DNA must reach the nucleus for transcription. The general belief is that DNA escapes from endosomes and travels within the cytosol to reach the nucleus periphery [76]. However, cytoplasmic injection of naked DNA does not result in transfection [77]. DNA in the cytosol may suffer from cytosolic degradation and it must be protected in order to reach the nucleus in a viable form. This role is believed to be performed by cationic lipids or polymers that remain associated with the DNA after endosomal escape. Endocytic vesicle movement to the periphery of the nucleus followed by DNA release near the nucleus is also a possibility [77]. The cytoskeleton elements share in the transport of vesicles from one organelle to another and hence they may direct the vesicles to desirable or undesirable locations. Microtubule disruption generally enhances lipoplex-mediated gene transfer, presumably by preventing transit to lysosomes. In the following section, we focus mainly on endosomal escape and nuclear transfer as the two

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most important barriers that limit transfection efficiency. We also discuss the possibility of mitochondrial targeting as a means to treat a variety of mitochondrial diseases. 25.4.1 Endosomal Escape The most important functional devices used to enhance endosomal escape and cytosolic delivery are briefly discussed. 25.4.1.1 Fusogenic Lipids Transfection mediated by cationic lipids is usually enhanced by the presence of DOPE, a fusogenic lipid [49, 78]. Lipoplexes containing DOPE can release the associated DNA into the cytosol through fusion with endosomal membranes. DOPE forms a stable lipid bilayer at physiological pH ∼7; however, at an acidic pH 5–6, it undergoes a transition from a bilayer to an inverted hexagonal structure, which fuses and destabilizes the endosomal membrane, releasing its contents into the cytosol [79]. We have previously shown that replacing EPC with DOPE in the R8-MEND system enhanced the transfection activities by severalfold [74]. Evidence exists to show that fusion with the endosomal membrane is essential for DOPEcontaining lipoplexes [79]. It is possible that only DNA or the lipoplex as a whole is released to the cytosol after fusion. If lipoplexes are released, the dissociation of DNA must occur in the cytosol or even at the nuclear membrane to achieve transfection. We recently developed multilayered nanoparticles for penetrating the endosome and nuclear membrane via a stepwise membrane fusion process [80]. The developed tetralamellar MEND (T-MEND) consists of a DNA/ polycation condensed core covered with two nuclear membrane fusogenic inner envelopes and two endosome-fusogenic outer envelopes. A screening of different fusogenic lipids revealed that the combination DOPE/cardiolipin (CL) (1/1) is optimum for nuclear fusion while the combination DOPE/phosphatidic acid (PA) (7/2) was optimum for endosome fusion. It is worth mentioning that the endosome fusion lipids required a high percentage of DOPE and the presence of the R8 peptide while the nuclear fusion lipid functioned optimally with lower amounts of DOPE and without R8. Intracellular FRET analyses showed that DNA is delivered to the nucleus synergistically and involves the function of both endosome-fusogenic and nucleus-fusogenic envelopes. This novel multilayered structure resulted in dramatic levels of gene expression, even in nondividing cells, making it highly promising for in vivo gene delivery applications. 25.4.1.2 Fusogenic Peptides DNA carriers lacking fusogenic lipids are not released efficiently into the cytosol unless additional functional devices for endosomal release are used. Viruses such as the influenza and adenovirus utilize the acidic pH of endosomes to induce endosomal disruption or fusion. Similarly, functional devices, which take advantage of the acidic pH of the

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endosome to induce their rupture, are incorporated in these systems. An example of such functional devices is the pH-sensitive fusogenic peptides. These peptides are derived from viruses, such as the peptide derived from the N-terminal sequence of the influenza virus hemagglutinin subunit HA-2, or prepared synthetically, such as GALA [81, 82]. The incorporation of the influenza HA-2 subunit augmented Tf-PLL-mediated gene transfer. The synthetic peptide GALA is comprised of a Glu-Ala-Leu-Ala repeat, designed to mimic the membrane penetrating activity of viruses. The peptide undergoes conformational changes at the low pH in the endosomes to interact with and perturb the endosomal membrane [82]. We have previously shown that the presence of the cholesterol-GALA peptide on the liposomal membrane effectively enhances the endosomal release of the liposome contents [60]. The cholesterol moiety was essential for attaching GALA to the liposome surface since the use of GALA in the liposome containing medium or its encapsulation in the liposomes failed to induce cytosolic release. Our studies with in vitro energy transfer and dynamic light scattering suggest that the endosomal escape of the liposomal aqueous phase of Tf-modified liposomes can be attributed to a pH-dependent membrane fusion. We further extended this study by developing the Tf-polyethylene glycol (PEG)-MEND modified with GALA [67]. Surprisingly, cholesterol-GALA and a PEGylated GALA were found to interact synergistically to induce membrane fusion between the liposomes and endosomes and transfection activities were enhanced 100 times. The presence of flexible PEG spacers is favorable for ligand recognition by Tf receptors; however, PEG may interrupt the interaction of GALA and the endosomal membrane due to steric hindrance. Therefore the introduction of GALA on the tip of PEG is important for enhancing endosomal escape in the presence of PEG since the interaction of GALA and the endosomal membrane is facilitated. 25.4.1.3 Membrane Disruptive Polymers The polycation PEI has an intrinsic ability to cause endosomal release without membrane fusion [83]. This is evident by the observation that transfection with PEI-polyplexes is not improved by fusogenic peptides or chloroquine [84]. A proton sponge hypothesis was proposed by Behr and co-workers to explain this phenomenon. This hypothesis proposes that PEI becomes more protonated at a low pH as in endosomes. This protonation triggers an influx of Cl− ions with protons leading to an influx of water and finally the swelling and rupturing of the endosomes [85]. Blocking vascular proton pumps using bafilomycin A1 inhibited PEImediated transfection. Earlier polycations without a proton sponge mechanism such as PLL were much less efficient than PEI. 25.4.1.4 Role of R8 in Endosomal Escape The R8-MEND system containing DOPE in its lipid composition showed high transfection activities in cultured cells. The activities were not improved in the presence of the endosome disrupting reagent chloroquine, suggesting that endosomal escape is a

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relatively efficient process. The transfection activities of R8-MEND were dramatically reduced when the R8 was replaced with a similar positive sequence octalysine (K8) [86]. This prompted us to consider the possibility that the effect of R8 is not solely dependent on a positive charge. We investigated the mechanism of the improved activity of the R8-MEND compared to the K8-MEND. The extent and rate of uptake of R8- and K8-modified nanoparticles (NPs) were similar and both were taken up by cells mainly through macropinocytosis. This indicates that the main difference between R8- and K8-NPs lies mainly in their intracellular trafficking. The colocalization of DNA encapsulated in R8-NPs with endosomes was lower than that encapsulated in K8-NPs. A silencing study using specific siRNA showed that R8-NPs are much more efficient that K8-NPs. Since the silencing effect is independent of nuclear delivery, this confirms that R8 plays an important role in enhancing the endosomal escape of DOPE-containing lipid vesicles. Since both R8- and K8-modified NPs contain DOPE, fusion was postulated as the main mechanism of endosomal escape. Using live cell spectral imaging, we showed that fusion in the case of R8-NPs, but not K8-NPs, was inhibited in the presence of bafilomycin A1, indicating that fusion is acidic dependent only in the case of R8-NPs [86]. Lipid mixing studies confirmed this finding, since R8-Lip is able to fuse efficiently to artificial membranes at both neutral and acidic conditions while fusion in the case of K8-Lip occurred only at a neutral pH. Consistent with this finding, R8-Lip caused the efficient release of calcein from liposomes at both acidic and neutral pH while K8-Lip was functional only at neutral pH. The superiority of R8-NPs over K8-NPs and the positive role of R8 in endosomal escape can be explained in the following model (Figure 25.2). At neutral pH, both R8- and K8-NPs fuse efficiently to endosomes, due to the possible approach of liposomal and endosomal membranes through ionic interactions. Upon maturation, the endosomes become more acidic and this decreases the net positive charge of K8 due to the deprotonation of some adjacent amino groups, resulting in a reduced interaction between liposomal and endosomal membranes with a subsequent decrease in fusion. In the case of R8, the total positive charge of R8 remains high due to the complete protonation of all arginine residues. Therefore electrostatic interactions with endosomal membranes remain high and fusion continues to proceed. The binding between R8 and the endosomal membrane is also aided by bidentate hydrogen bonding between R8 and amphoteric components in the endosomal membrane. Since the transit time of endosomes with a neutral pH (immediately after uptake) is short, it would be expected that overall endosomal escape is higher for R8-NPs, which continue to escape after endosome maturation. Similar to R8, the novel IRQ peptide with a lipid composition DOPE and cholesteryl hemisuccinate (CHEMS) caused efficient endosomal escape, as judged by confocal microscopy and by the silencing effect of entrapped siRNA [75]. Interestingly, the same endosomal escape of siRNA could be observed, even in the presence of the nonfusogenic lipid EPC/CHEMS. This indicates

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R8-MEND Fusiondependent Cytosolic release

Macropinosome

Acidification

Acidification

Degradation Recycling

Degradation Recycling Stop cytosolic release

Continue cytosolic release

Figure 25.2. A schematic diagram of endosomal escape of R8- and K8-MEND. After internalization, membrane fusion occurs at a neutral pH in both cases. Upon acidification, the membrane fusion stops in the case of K8-MEND while it continues in the case of R8-MEND. Extensive interactions between endosomal and liposomal membranes occur in both neutral and acidic pH in the case of R8-MEND. (See color insert.)

that the peptide promoted endosomal escape even in the absence of DOPE, or under nonfusogenic conditions. Although both lipid compositions showed the same endosomal escape activity, the detected siRNA in cytosol was associated with the lipids in the case of EPC/CHEMS while more free siRNA was detected in the cytosol in the case of DOPE/CHEMS. This suggests that the decoating of the nucleic acid core is more efficient when DOPE is used. Some of the lipid coat may be consumed during fusion with the endosomal membrane, which is not the case with the nonfusogenic EPC/CHEMS. Consistent with this finding, the silencing effect of IRQ-MEND with DOPE/CHEMS was higher than that with EPC/CHEMS. In the previous discussion, we emphasized the positive role of R8 and IRQ peptides in enhancing endosomal escape of gene delivery systems. We are currently investigating these mechanisms, in attempts to further improve gene delivery. 25.4.2 Intracellular Targeting 25.4.2.1 Mitochondrial Targeting Mutations and/or defects in mitochondrial DNA (mtDNA) have been implicated in a variety of human disorders, the so-called mitochondrial diseases [87–89]. Therefore mitochondrial gene delivery leading to the complementing of normal mtDNA, repair of mutated mtDNA, inhibition of replication for mutated mtDNA, and/or the degradation of mutated mtDNA would be a novel strategy for the cure of such diseases. The loci of mtDNA mutations are known for many mitochondrial diseases [90–92]. Accordingly, the precise site-specific correction of mtDNA mutations

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is a sophisticated strategy for the treatment of genetic disorders. Using a cellfree DNA repair assay with oligodeoxynucleotides (ODNs), Chen et al. [93] demonstrated that isolated rat liver mitochondria contain the machinery required for the repair of genomic single-point mutations and concluded that ODN might provide a novel approach to the treatment of certain mitochondrial-based diseases. Taylor et al. [94] proposed the selective inhibition of mutant mtDNA replication, which would allow propagation of only wild-type DNA, as a treatment for mitochondrial diseases. Using an in vitro replication runoff assay, they showed that antigenomic peptide nucleic acids (PNAs) specifically inhibited the replication of mutant, but not wild-type, mtDNA templates. As described above, the delivery of therapeutic ODN and PNA to the mitochondrial matrix holds promise for the treatment of mitochondrial diseases that result from mtDNA mutations. The conjugation of a mitochondrial targeting signal (MTS) peptide to exogenous small linear DNAs including either ODNs, double-stranded DNA, or PNA was found to aid their delivery to the mitochondria [95–97]. Seibel et al. [96] showed that these conjugates are imported into mitochondria through the outer and inner membranes via the protein import machinery. Reportedly, DNA 17 to 322 bp in length can be used in this strategy. A similar strategy has also been developed for the mitochondrial delivery of PNA. Using membrane permeability toxin as a device for cytoplasmic delivery, MTS-conjugated PNA can be imported into mitochondria. This method provides a viable strategy for the modification of mitochondrial DNA in cultured cells, animals, and humans [95]. Conjugation of an MTS peptide to small linear DNAs was found to aid their delivery to the mitochondria [95–97], but this strategy is severely limited by cargo size [98–100]. As a result, MTS cannot deliver macromolecules, such as mtDNA and plasmid (pDNA), to mitochondria. Weissig and co-workers attempted to deliver pDNA to mitochondria using DQAsomes, which are mitochondriotropic and cationic “bola-lipid”-based vesicles [101, 102]. In previous studies, DQAsomes–DNA complexes (DQAplexes) were shown to selectively release pDNA upon contact with isolated mitochondria derived from the rat liver [102]. In addition, DQAplexes apparently escape endosomes without losing their pDNA, and specifically release pDNA proximal to mitochondria [101]. More recently, Weissig and co-workers attempted the delivery of pDNA to the mitochondrial matrix by means of improved DQAsomes [103]. In the future, the effective delivery of circular DNA to the mitochondrial matrix using such novel approaches holds considerable promise for use in mitochondrial gene therapy. We recently proposed the MITO-Porter as a MEND for mitochondrial delivery [104–106]. The MITO-Porter is a liposome-based nanocarrier that delivers cargo to mitochondria via a membrane-fusion mechanism (Figure 25.3). Mitochondrial delivery using MITO-Porter requires the following three steps: (1) the carrier must be delivered to the cytosol; (2) intracellular trafficking of the carrier, including mitochondrial targeting, must be regulated;

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Figure 25.3. A schematic diagram of mitochondrial delivery via membrane fusion by MITO-Porter. R8, octaarginine; OM, outer membrane; IMS, intermembrane space; IM, inner membrane. (See color insert.)

and (3) mitochondrial delivery via membrane fusion must be regulated. We are hopeful that MITO-Porter can be used to deliver a wide variety of carrierencapsulated molecules, regardless of size or physicochemical properties, to the mitochondrion. The first barrier to intracellular targeting is the plasma membrane. Therefore the transduction activity of MITO-Porter should be required for mitochondrial delivery. We previously showed that high-density R8-modified liposomes are taken up mainly through macropinocytosis and delivered to the cytosol while retaining the aqueous phase marker [71]. Therefore we chose R8 as a cytosol delivery device for MITO-Porter. We also expected that R8, which mimics TAT, might have mitochondrial targeting activity [107, 108]. We first screened different lipids to find a liposome composition with high fusion ability using fluorescence resonance energy transfer (FRET) analysis. Two highly fusogenic lipid compositions were used to construct the MITOPorter. The green fluorescence protein (GFP) was used as a model macromolecule, which permitted us to verify its delivery to mitochondria by CLSM. GFP was efficiently delivered to mitochondria. Thus the MITO-Porter holds promise as an efficacious system for the delivery of therapeutic molecules to mitochondria.

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In this section, recent reports concerning mitochondrial gene delivery and therapy were described. We also described MITO-Porter, a liposome-based nanocarrier that introduces cargoes into mitochondria via a membrane fusion mechanism. We believe that mitochondrial gene delivery via MITO-Porter has potential for use as a novel therapeutic strategy. Studies concerning mitochondrial gene delivery using MITO-Porter are currently in progress. 25.4.2.2 Nuclear Transfer of DNA In mitotic cells, pDNA primarily enters the nucleus during the period, M-phase, when the nuclear membrane structure breaks down [109–112]. In contrast, in nondividing cells, the nuclear membrane structure remains intact. Therefore the nuclear membrane is the ultimate barrier to be overcome for a successful nuclear delivery in nondividing cells. The mutual transport of various types of molecules between the cytosol and nucleus was carried out via nuclear pore complex (NPC), which allows the passive transport of gold nanoparticles with sizes of <9 nm [113]. Wolff and co-workers investigated the effect of pDNA size on nuclear transport activity using digitonin-permeabilized cells [114–115]. Short DNA (<200bp) fragments were effectively imported into the nucleus, whereas nuclear import decreased when the size of the pDNA increased and became negligible at >1500 bp. Generally used plasmid DNA is too large to pass through the nuclear pore complexes. In fact, microinjection studies demonstrated that more than a 100-fold larger amount of pDNA is essential to achieve a transgene expression by cytoplasmic microinjection comparable to that by nuclear microinjection, suggesting that less than 1% of the cytoplasm-microinjected plasmid DNA reaches the nucleus [116]. Therefore efficient systems for the nuclear delivery of pDNA would be highly desirable for developing an efficient gene delivery system. The mechanism for the nuclear localization signals (NLSs)-dependent nuclear import of proteins is well understood [117]. One of the most defined NLSs is a peptide (PKKKRKV) derived from the SV40 T-antigen (NLSSV40). The endogenous or chemically modified NLSs bind to importin-α and are subsequently recognized by importin-β, which facilitates the nuclear import of cargo via its efficient interaction between hydrophobic phenylalanine-glycine (FG)-rich domains inside the nuclear pore [118]. One of the most important characteristics of NLS-dependent transport is that the threshold size of the NPC would be expanded to >39 nm in size [119–120]. Moreover, the latter research indicates that the larger size of NLS-displaying phages (∼55 nm) would be acceptable for NPC-dependent transport [121]. In the light of these findings, numerous attempts have been made to improve nuclear delivery (Figure 25.4). First, NLSs were chemically linked to the pDNA itself [122]. However, in this strategy, >100 NLS peptides/1 kbp are required to induce the nuclear delivery of pDNA, severely inhibiting transcription. Similarly, several groups attempted to conjugate NLSs on the edge of linearized pDNA. However, microinjection studies [123, 124] have not validated the utility of this strategy. Since it uses highly cationic NLSSV40, this

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(A) Direct modification

(B) Insertion of transcription factorbinding sequence

(D) NLS-modified envelope-type nanoparticle

(C) Complex formation with NLS-modified polycation or NLS-peptides

(E) Envelope type nano particle encapsulated with endosome- and nuclear membrane-fusogenic membrane

Figure 25.4. Strategies to overcome a nuclear member barrier. (A) Direct modification of NLS peptides and/or nucleus-targeting proteins to pDNA by chemical linkage or avidin–biotin interaction. (B) Insertion of nuclear-transcription factor-binding sequences (i.e., NF-κB) in the pDNA. (C) Condensation of pDNA with NLS itself or NLS-modified polymers. (D) Surface modification of NLS on the surface of lipid-based particle. (E) Encapsulation of pDNA in NLS-fusogenic lipid envelope. (See color insert.)

may be explained by assuming that its recognition by importin is prevented by electrostatic interactions with the negatively charged pDNA. One excellent strategy for overcoming this issue is to modify the pDNA with a NLS-possessing protein or importin-α via avidin–biotin interactions [115, 125]. Alternatively, specific DNA sequences can be introduced into the plasmid DNA (i.e., NF-κB binding sequence [126–129], SV40 enhancer [130, 131], and smooth muscle gamma actin promotors [132]), which allow the binding of nuclear transcription factors in the cytosol, thus supporting nuclear import. This strategy would be advantageous in controlling nuclear import in a signal-dependent and cell type-dependent manner. A second approach is to condense pDNA with an NLS-modified polycation. One example of this is a NLSSV40-modified PLL. The condensation of pDNA with this polycation slightly enhanced transgene expression compared with nonmodified PLL [133, 134]. To avoid electrostatic interaction between

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NLS and DNA, a novel NLS, derived from heterogeneous nuclear ribonucleoprotein-A1 (M9) was conjugated to the cationic peptide (scattered SV40 NLS). Complex formation with M9-modified polycations exhibited enhanced transfection activity when transfected by lipofection [135]. An alternate approach is to condense pDNA with the NLS peptide itself, or NLS-like peptides. Various NLSs, such as tetramers of SV40 NLS [136], TAT oligomers [137], Mu peptide derived from the adenoviral core [138, 139], and protamine [140, 141], are currently being investigated. DNA condensation with these peptides enhances transgene expression. Of note, a cytoplasmic microinjection study showed that transgene expression is enhanced by condensation with protamine, a peptide possessing four possible NLS-like regions consisting of basic amino acid and proline or serine residues, depending on the charge ratio (+/−) [141]. At a low charge ratio, the basic amino acids of protamine may be dominantly consumed in the condensation of pDNA, and, as a result, NLS-like functions would be limited. By increasing the charge ratio, the protamine displayed on the surface of particles functions as an NLS, allowing the particles to be recognized by nuclear transport-associating proteins. These data are important examples that demonstrate the significance and simultaneous difficulty of controlling the topology of an NLS. For an NLS to function efficiently, it must be displayed on the surface of the particle. However, the density of its surface display is an accidental event determined by the condensation status. We recently proposed a third strategy for nuclear delivery, which mimics the nuclear gene delivery system of adenoviruses. In adenoviruses, the NLS is spontaneously displayed on the adenovirus particle via a well-ordered assembly of capsid proteins around the DNA core. To control the topology of the NLS on the particle surface, a lipid derivative of a nuclear-targeting device was synthesized with the lipid moiety incorporated into the envelope and the NLS spontaneously oriented outward from the particle surface. This strategy resulted in an effective transfection of nondividing cells (i.e., primary dendritic cell cultures) [142]. Moreover, our group has also focused on sugars as nuclear-targeting devices. In recent decades, it has been reported that bovine serum albumin (BSA) modified with certain sugars accumulate in the nucleus [143]. Modifying the surface of R8-MEND with sugars increased transfection activity 10- to 100-fold [144]. Finally, we introduced a novel concept for overcoming intracellular membrane barriers using a stepwise membrane fusion, as described in an earlier section. To realize this concept, an innovative nanotechnology was developed to create a multilayered nanoparticle, which we refer to as a tetralamellar MEND (T-MEND) [80]. The critical structural elements of T-MEND are a DNA/polycation condensed core coated with two nuclear membranefusogenic inner envelopes and endosome-fusogenic outer envelopes, which are shed in a stepwise fashion as the core is trafficked through the cell. The R8-modified T-MEND can induce macropinocytosis, and thus the outer membranes were optimized to fuse with macropinosomes. The internal double-lamellar structure is required for nuclear delivery via stepwise membrane

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fusion, independent of the nuclear pore complex. Furthermore, we found that coating the core with the minimum number of nucleus-fusogenic lipid envelopes (i.e., two) is essential for facilitating transcription. As a result, the T-MEND achieves dramatic levels of transgene expression in nondividing cells, and with facile modifications it can be engineered to deliver non-DNA cargoes (e.g., siRNAs, proteins) to other organelles.

25.5

CONCLUSION

A nonviral gene delivery system with high transfection activities and minimum cytotoxicity is highly desired. The system should have the ability to target specific cell populations and to function after systemic administration. The most important barrier against efficient nonviral gene delivery is inadequate intracellular trafficking rather than low cellular internalization. Overcoming lysosomal degradation and endosome entrapment is a challenging task. Enhancing nuclear delivery is an additional task in nondividing cells. Recently developed nonviral vectors produce efficient gene delivery in vitro; however, their in vitro performance is not well correlated with in vivo behavior. Therefore parallel efforts should be directed toward improving the targeting ability of the systems and overcoming various extracellular barriers in in vivo situations. The rationalized design of novel components and systems is the key for the development of smart nanosystems for future medicine.

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132. Miller, A. M. and Dean, D. A. Cell-specific nuclear import of plasmid DNA in smooth muscle requires tissue-specific transcription factors and DNA sequences. Gene Ther. 15(15): 1107–1115 (2008). 133. Chan, C. K. and Jans, D. A. Enhancement of polylysine-mediated transferrinfection by nuclear localization sequences: polylysine does not function as a nuclear localization sequence. Hum. Gene Ther. 10(10): 1695–1702 (1999). 134. Chan, C. K., Senden, T., and Jans, D. A. Supramolecular structure and nuclear targeting efficiency determine the enhancement of transfection by modified polylysines. Gene Ther. 7(19): 1690–1697 (2000). 135. Subramanian, A., Ranganathan, P., and Diamond, S. L. Nuclear targeting peptide scaffolds for lipofection of nondividing mammalian cells. Nat. Biotechnol. 17(9): 873–877 (1999). 136. Ritter, W., Plank, C., Lausier, J., Rudolph, C., Zink, D., Reinhardt, D., and Rosenecker, J. A novel transfecting peptide comprising a tetrameric nuclear localization sequence. J. Mol. Med. (Berlin) 81(11): 708–717 (2003). 137. Rudolph, C., Plank, C., Lausier, J., Schillinger, U., Muller, R. H., and Rosenecker, J. Oligomers of the arginine-rich motif of the HIV-1 TAT protein are capable of transferring plasmid DNA into cells. J. Biol. Chem. 278(13): 11411–11418 (2003). 138. Keller, M., Harbottle, R. P., Perouzel, E., Colin, M., Shah, I., Rahim, A., Vaysse, L., Bergau, A., Moritz, S., Brahimi-Horn, C., Coutelle, C., and Miller, A. D. Nuclear localisation sequence templated nonviral gene delivery vectors: investigation of intracellular trafficking events of LMD and LD vector systems. Chembiochem 4(4): 286–298 (2003). 139. Akita, H., Tanimoto, M., Masuda, T., Kogure, K., Hama, S., Ninomiya, K., Futaki, S., and Harashima, H. Evaluation of the nuclear delivery and intra-nuclear transcription of plasmid DNA condensed with micro (mu) and NLS-micro by cytoplasmic and nuclear microinjection: a comparative study with poly-l-lysine. J. Gene Med. 8(2): 198–206 (2006). 140. Sorgi, F. L., Bhattacharya, S., and Huang, L. Protamine sulfate enhances lipidmediated gene transfer. Gene Ther. 4(9): 961–968 (1997). 141. Masuda, T., Akita, H., and Harashima, H. Evaluation of nuclear transfer and transcription of plasmid DNA condensed with protamine by microinjection: the use of a nuclear transfer score. FEBS Lett. 579(10): 2143–2148 (2005). 142. Nakamura, T., Moriguchi, R., Kogure, K., Minoura, A., Masuda, T., Akita, H., Kato, K., Hamada, H., Ueno, M., Futaki, S., and Harashima, H. Delivery of condensed DNA by liposomal non-viral gene delivery system into nucleus of dendritic cells. Biol. Pharm. Bull. 29(6): 1290–1293 (2006). 143. Duverger, E., Carpentier, V., Roche, A. C., and Monsigny, M. Sugar-dependent nuclear import of glycoconjugates from the cytosol. Exp. Cell Res. 207(1): 197–201 (1993). 144. Masuda, T., Akita, H., Nishio, T., Niikura, K., Kogure, K., Ijiro, K., and Harashima, H. Development of lipid particles targeted via sugar-lipid conjugates as novel nuclear gene delivery system. Biomaterials 29(6): 709–723 (2008).

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CHAPTER 26

Cellular Interactions of PlasmonResonant Gold Nanorods QINGSHAN WEI and ALEXANDER WEI Department of Chemistry, Purdue University, West Lafayette, Indiana

26.1

INTRODUCTION

Colloidal gold has a long history of clinical use, both as immunocytochemical probes for ex vivo applications [1–4] and also as colloidal adjuvants for in vivo radiotherapies [5–8]. In recent years, developments in nanoscale materials chemistry have led to the efficient syntheses of anisotropic gold nanoparticles with tunable surface plasmon modes, with optical resonances ranging from visible to near-infrared (NIR) wavelengths [9–12]. Several types of gold nanostructures are currently being investigated as contrast agents for various optical imaging modalities, but those with plasmon resonances in the NIR region between 750 and 1300 nm are particularly favorable for biomedical imaging, as shorter wavelengths are extinguished by hemoglobin or other endogenous pigments, and longer wavelengths are strongly attenuated by water [13, 14]. Gold nanorods (GNRs) are especially attractive for their straightforward and highly reproducible synthesis and for the tunability of their plasmon resonances, which can be centered at any position within the NIR window (Figure 26.1) [15]. GNRs also have narrower linewidths relative to spherical gold nanoparticles at comparable resonance frequencies, due to reduced radiative damping effects [16]. GNRs have been efficiently synthesized using seeded growth conditions, with various modifications in reaction conditions allowing for fine control over aspect ratio, scalability and uniformity, optical and colloidal stability, and absorption and scattering properties [10, 15, 17]. The latter define the types of optical imaging modalities that can be empowered by the

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Figure 26.1. (A) Gold nanorod (GNR) with longitudinal and transverse plasmon resonances (LR and TR). (B) Simulations of λLR in GNRs with different aspect ratios (R), based on modified Mie theory [20]. (C) Transmission electron microscopy (TEM) image of NIR-absorbing GNRs, prepared by a seeded growth method [21]. (Reproduced with permission of the American Chemical Society.)

use of GNRs as contrast agents [18]. The remarkable optical properties of GNRs are also matched by an equally remarkable capacity to mediate localized photothermal effects, giving rise to the popular concept of theranostics (a combination of diagnostics and therapeutic action) [19]. Finally, GNRs have very recently become commercially available, increasing the opportunities for application to a broad range of biomedical areas. Here we discuss some recent advances in the chemistry and photophysical properties of GNRs, with a focus on biological imaging and photothermal activity at the cellular level. While much attention has been focused on the synthesis and structure–function relationships of GNRs, the process chemistry supporting their biological applications (i.e., detoxification and surface functionalization) is also critical and cannot be overlooked. The multifunctional activity of functionalized GNRs will be illustrated through several examples that also demonstrate the impact of surface chemistry on targeting, cytotoxicity, and their subsequent implications for clinical use.

26.2

SURFACE CHEMISTRY OF GNRS

The synthesis of GNRs will not be discussed in detail, as this subject has already been reviewed elsewhere [10, 15, 17]. However, it is important to point out that NIR-absorbing GNRs are typically synthesized under micellar conditions using cationic surfactants such as cetyltrimethylammonium bromide (CTAB), which introduces some complications regarding cytotoxicity and nonspecific cell uptake (to be discussed below). Surface modification is thus critical in the development of GNRs for biomedical applications, for improving biocompatibility and minimizing cytotoxicity as well as for cellular and subcellular targeting (Figure 26.2). With respect to impending clinical applications, GNRs and other nanoparticle-based agents will need to meet multiple criteria: (1) dispersion stability

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Figure 26.2. Some surface functionalization and bioconjugation methods applied toward GNRs [18]: (A) electrostatic adsorption onto polyelectrolyte (PE)-coated GNRs [22–24]; (B) covalent attachment via carbodiimide coupling [25]; (C) “click” bioconjugation [26, 27]; (D) chemisorption using thiols [28, 29]; and (E) chemisorption using in situ dithiocarbamate (DTC) formation [19, 30]. (Reproduced with permission of Wiley Interscience.)

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in blood and other physiological fluids, (2) functionalization methods for siteselective targeting and/or cell uptake, (3) resistance against nonspecific cell uptake and protein adsorption, (4) sufficiently long circulation lifetimes to allow efficient delivery to the region of interest, and (5) low cytotoxicity and inflammatory response. Nanoengineered systems must also be robust against chemical degradation while under biological exposure, to avoid opsonization and loss of biocompatibility. Altogether, these requirements present a daunting set of obstacles for engineered nanomaterials. For this reason we will pay considerable attention to the surface chemistry of GNRs, and their subsequent impact on biological systems.

26.2.1 Methods for GNR Coating and Functionalization Decades of research have already been devoted toward the bioconjugation of colloidal gold nanoparticles, many of which have been targeted toward cellsurface biomarkers for ex vivo immunolabeling studies by optical or electron microscopy [1]. However, many of those protocols are not straightforwardly adapted toward the bioconjugation of GNRs, which are typically synthesized or stabilized in micellar CTAB solutions [10, 15, 17]. In this section we describe some recently developed methods for the functionalization of CTAB-stabilized GNRs (Figure 26.2), all of which are applicable for short-term in vitro studies in a laboratory setting. However, it is important to note that most of these have not been evaluated for long-term stability or cytotoxicity due to the leaching of CTAB or other agents. Indeed, CTAB presents significant challenges in the preparation of biocompatible GNRs and can thwart efforts to meet the aforementioned requisites for preclinical evaluation and subsequent in vivo studies, if appropriate steps are not taken to ensure its complete removal [31]. 26.2.1.1 Physisorptive Coatings The physical or electrostatic absorption of polystyrenesulfonate (PSS) and other anionic polyelectrolytes is the simplest and most direct method of coating CTAB-stabilized GNRs (Figure 26.2A). GNRs coated with multiple layers of polyelectrolyte can form stable dispersions at various pH or ionic strength, and are stable against dilution effects [23, 32]. The latter is important because it enables most of the soluble CTAB to be removed from aqueous suspensions of GNRs by multiple chloroform extractions. In the absence of such stabilizing agents, the GNRs have a strong tendency to flocculate when the CTAB drops below the critical micelle concentration (ca. 1 mM) [33]. The polyelectrolyte coatings prevent the GNRs from agglomerating and can also reverse their zeta potential from positive to negative, which substantially improves their compatibility with biological media. Generic “blocking” proteins such as bovine serum albumin (BSA) can also adsorb onto CTAB-coated GNRs by physisorption and provide good dispersion stability in physiological media [34–36].

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Polyelectrolyte coatings have also been used to support antibodies [24] or proteins such as myoglobin [37, 38], transferring [23], or streptavidin [39, 40], again by physisorption using a relatively low pH. Antibody-functionalized GNRs are ostensibly useful for targeting cell-surface receptors and antigens; examples include epidermal growth factor receptor (EGFR) [41, 42], prostate specific antigen (PSA) [43], the δ-opioid receptor [44], the CD11b antigen on activated macrophage [34], and an antigen of the parasitic tachyzoite Toxoplasma gondii [35]. Multilayer polyelectrolyte coatings can also be used to load molecular cargoes onto GNRs, as demonstrated with dye molecules [45] and hydrophobic proteins such BSA [46]. Covalent coupling on polyelectrolyte supports has been applied toward the bioconjugation of GNRs. For example, amine- and acetylene-terminated biomolecules have been incorporated onto GNRs via carbodiimide coupling [25] and “click” bioconjugation [26], by using polyelectrolyte coatings bearing activated N-hydroxysuccinimide (NHS) carboxylate esters or azides, respectively (Figure 26.2B,C). Covalent crosslinking may also further increase the stability of bioconjugated GNRs, although the long-term stability of the physisorbed polyelectrolyte continues to remain an open question for the time being.

26.2.1.2 Chemisorptive Coatings An alternative route for the surface bioconjugation of GNRs and other Au nanoparticles is by direct surface ligation, using functional groups with a strong affinity for Au. Thiols (–SH) are presently the most widely used functional group [47], and have been conjugated to proteins [29], oligonucleotides [48, 49] and DNA aptamers [50], and oligopeptides [27] for their immobilization on GNR surfaces (Figure 26.2D). Heterobifunctional ligands can be introduced for sequential bioconjugation, including mercaptoacetic acid [51–53], dihydrolipoic acid [54, 55], cysteine [52, 56–59], glutathione [56–58], and thiol-appended biotin [60, 61]. An interesting consequence of the chemisorptive approach is that many of these ligands adsorb preferentially onto the tips of GNRs, either on the {111} facets or along their edges. This anisotropic functionalization has enabled GNRs to be assembled in end-to-end fashion, with a strong effect on their plasmon resonances [49, 51–53, 56, 58–62]. When coupled with biomolecular recognition events, such assemblies can be used as optical switches for biosensing applications [49, 53, 56, 62, 63]. Despite their popularity for the functionalization of Au surfaces, chemisorbed thiols may have limited stability when exposed to physiological conditions [64, 65]. Several groups have shown that chemisorbed alkylthiols and dithiols are readily displaced by surface exchange by other molecules (including biogenic thiols such as glutathione) [66–70], and degrade under oxidative conditions or in the presence of electrolyte [71–73]. These pathways compromise the integrity of alkylthiol-based self-assembled monolayers [65, 67], with negative consequences for biological applications.

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Robust alternatives to thiols are currently being developed for the surface modification of GNRs and other nanoparticles. One promising development involves ligands based on the dithiocarbamate (DTC) group ( − NCS−2 ), which can be formed in situ by the condensation of alkylamines with CS2 under moderately basic conditions (Figure 26.2E) [74, 75]. This is a useful addition to existing bioconjugation methods, as it provides the option of attaching amine-terminated ligands directly onto Au surfaces, while retaining the simplicity of chemical self-assembly. In situ DTC formation can be performed in aqueous suspensions of GNRs; once adsorbed onto the metal surface, the DTC ligands are resistant to displacement by competing surfactants and to ambient oxidation, much more so than their thiolated counterparts. In situ DTC chemisorption has been demonstrated on GNRs using amine-terminated polyethylene glycol (PEG) [30] and with diamine-functionalized PEG, followed by conjugation with targeting ligands such as folic acid (Figure 26.2E) [19,76,99]. Other recent examples involving DTC chemisorption include the conjugation of heterobifunctional linkers [77–79], DNA oligonucleotides [80], and prolineterminated oligopeptides [75]. 26.2.1.3 Core–Shell Formation Another method of functionalizing GNRs is to encapsulate them within silica shells, by application of the Stöber process [81, 82]. The silica deposits preferentially onto the CTAB coating and in a nonuniform manner during the early stages of growth, often resulting in the formation of a mesoporous SiO2 shell [83]. Methods for functionalizing silica are well known and present more opportunities for endowing GNRs with biomolecular functionality. Chromophores can also be embedded within the silica matrix, as was done in a recent demonstration of silica-coated GNRs as SERS labels [84].

26.3

BIOCOMPATIBILITY AND CELLULAR UPTAKE OF GNRS

The cytotoxicity of nanomaterials has been a broadly discussed issue, attracting a great deal of attention worldwide [85]. With respect to GNRs, cytotoxicity studies have been few but are increasing in number. These toxicological studies are critical, as the potential of GNRs for biomedical applications can be derailed by the membrane-compromising effects of the cationic surfactant CTAB, which has a high acute cytotoxicity (IC50 < 10 μM) and has been shown to inhibit mitochondrial activity [86, 87]. As a result, considerable efforts have been made to reduce the toxic presence of CTAB [31, 32, 36, 88–90]. CTABstabilized GNRs coated with anionic polymers [22, 25], and serum proteins [32, 36], or treated with phosphatidylcholine [91] or synthetic cationic lipids [92] all have reduced cytotoxicity profiles relative to untreated GNRs, due in part to the shielding effect of those physisorbed materials. Furthermore, in vitro studies have shown that the CTAB-stabilized GNRs are not necessarily

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Figure 26.3. Nonspecific cell uptake of CTAB-stabilized GNRs [30]. (A) Overlay of transmission and TPL images of GNRs (green) internalized by KB cell, with nucleus outlined in red. (B) Trajectory of an internalized GNR exhibiting bidirectional motion, with overall displacement toward the cell nucleus. (C) TPL image of GNRs (yellow) segregated within inclusion bodies, 5 days after initial exposure; bar = 20 μm. (Reproduced with permission of the American Chemical Society.) (See color insert.)

cytotoxic by themselves, if the excess CTAB is removed from the solution. CTAB-coated GNRs that were internalized by K562 or KB cells (from human leukemia or nasopharyngeal carcinoma cell lines, respectively) did not produce an appreciable cytotoxic response even after several days of incubation [30, 88]. A second, more insidious, problem associated with CTAB is their role in mediating the nonspecific cell uptake of GNRs, even at very low surfactant levels. CTAB-stabilized GNRs have been observed to accumulate inside KB cells over the course of several hours [30]. The uptake of individual GNRs was monitored by two-photon excited luminescence (TPL), a nonlinear optical property intrinsic to the plasmon-resonant GNRs (Section 26.4.3). Singleparticle tracking analysis revealed a characteristic bidirectional motion with an overall trajectory in the direction of the nucleus, consistent with intracellular vesicular transport along microtubules (Figure 26.3). The CTAB-coated GNRs were ultimately segregated from the rest of the cell by compartmentation within inclusion bodies, and were passed down to daughter cells as the culture grew to confluence. Issues of cytotoxicity aside, the nonspecific uptake of GNRs remains a concern due to their powerful photothermal activities (Section 26.5), so the rigorous removal of CTAB is necessary to minimize collateral damage. The nagging problem of CTAB removal has been addressed by several different approaches. On the laboratory scale, it has been shown that ion-exchange resins can mediate an essentially complete exchange of CTAB with chemisorptive surfactants; the resin physically prevents GNRs from flocculation during the exchange process [93]. CTAB-stabilized GNRs have also been treated with thiol-terminated poly(ethylene glycol) (PEG-SH) [29, 89, 94, 95] and with amine-terminated PEGs in the presence of CS2 (in situ PEG-DTC

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70-kDa PSS CTAB-coated GNRs

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Figure 26.4. Detoxification of GNRs. (Left) Scheme for removing CTAB from GNRs using PSS exchange [31]. (Right) Cytotoxicity profile (using KB cells) of PSS-coated GNRs contaminated with CTAB (䊏), and after further exchange with unadulterated PSS (ⵧ). (Reproduced with permission of the American Chemical Society.)

formation) [30], and can be extensively dialyzed for the effective removal of CTAB. PEG-coated GNRs form stable dispersions in buffered solutions and have low cytotoxicity even at high concentrations, with 90% cell viability reported for a GNR concentration of 0.5 mM [89]. These GNRs are also amenable to in vivo biodistribution studies, with a circulation halflife of many hours [89, 96]. CTAB can also be removed from GNRs on a batch scale without resin, simply by applying sodium polystyrenesulfonate (PSS) as an adsorbent and detergent [31]. At first glance this may be somewhat surprising, given the popular use of polyelectrolytes to stabilize GNRs by electrostatic adsorption over the short term (Figure 26.2A). However, a longer-term study over a period of weeks has revealed that PSS-stabilized GNRs gradually leach a cytotoxic material, which has been postulated to be a persistent PSS–CTAB complex [31]. Fortunately, CTAB-laden PSS can be exchanged with fresh polyelectrolyte to produce biocompatible, CTAB-depleted GNRs with no significant cytotoxicity up to 85 μg/mL (Figure 26.4). These purification methods permit further development of surface-functionalized GNRs for cellular targeting, without concern for contamination by CTAB.

26.4

CELLULAR IMAGING WITH GNRS

GNRs can be used as optical markers in a variety of biophotonic applications, for microscopy as well as for biomedical imaging modalities with greater penetration depth [18]. GNRs are versatile contrast agents because their TR and LR modes support optical responses at visible and NIR frequencies, respectively. In particular, the LR plasmon mode can enhance the two-photon absorption cross section of GNRs to generate a bright TPL signal with pulsed NIR laser excitation [97, 98], which has been employed with great advantage toward cellular confocal microscopy [19, 28, 30, 76, 99].

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26.4.1 Resonant Light Scattering The sensitivity and resolving power of optical and optoelectronic systems continues to improve, driving the development of imaging modalities based on resonant light scattering. Plasmon-resonant light scattering from GNRs prepared by seeded growth can be visualized by either standard darkfield microscopy or by confocal reflectance microscopy [16, 100–105], despite the fact that their extinction cross sections are dominated by absorption [106]. GNRs have a strong appeal as optical contrast agents because their TR and LR modes are active in both the visible and NIR range, respectively. Darkfield microscopy is generally performed at visible wavelengths supported by the TR mode, but the LR mode is able to support polarized light scattering, allowing for the tracking of lateral and especially rotational motion of single GNRs on artificial biomembranes [107] or attached to biomolecular motors such as F1-ATPase [39, 108]. GNRs can be used in darkfield microscopy for targeted cancer cell imaging. For example, antibody-labeled GNRs were observed to label malignant carcinoma cell lines by recognition of their cell-surface EFGRs, with selectivity over normal human keratinocytes [41]. The same strategy has been used to monitor the targeted nuclear delivery of GNRs conjugated with transferrin [23, 109] or cell-penetrating peptides [27]. Darkfield microscopy with whitelight illumination can also support multiplex labeling strategies, as demonstrated by the simultaneous detection of GNRs with different aspect ratios, targeted toward separate cell-surface biomarkers on human breast epithelial cells [110], or pancreatic cancer cells such as Panc-1 and MiaPaCa [109]. Darkfield imaging with GNRs can even be used to measure biomechanical properties: longer GNRs (R∼15) were embedded in a cardiac fibroblast network to track local deformations induced by mechanical stress [111]. Strain distributions were measured by monitoring the positions of GNRs in real time [111, 112].

26.4.2 Linear Photoluminescence While the absorption and scattering properties of gold nanoparticles are firmly established, their photoemission properties are much less well known. Indeed, the photoluminescence (PL) of metal nanoparticles is often overlooked, as metals are better known for their ability to quench fluorescent molecules or particles by back-electron transfer. Nevertheless, weak yet detectable PL can be generated from Au nanoparticles and GNRs using laser excitation [113– 116]. It is worth mentioning that PL can also be generated by UV excitation of very small (<2-nm) gold nanoclusters [117, 118], whose density of states are discretized and more like that of a semiconductor than a metal. GNRs can be excited above plasmon resonance (λex = 480 nm) to produce linear photoemission spectra with λem ranging from 548 to 588 nm [113]. The emission wavelength increases with aspect ratio, and the quantum efficiency

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(φem) has a quadratic dependency on GNR length. Plasmons appear to enhance the PL effect, and the φem of GNRs has been estimated to be over 106 times compared with bulk Au, due to resonant coupling with local electromagnetic fields. Subsequent studies have attributed the increase in PL to ultrafast, plasmon-enhanced emission [114, 115], but the effect of higher aspect ratios on PL is somewhat less clear [119], and in partial conflict with the earlier study [113]. An independent report has shown the emission band intensity to increase when using even longer GNRs (R = 13) with lower energy excitation (λex up to 690 nm) [120]. The linear PL of GNRs has been used in biosensing applications, but with regard to biological imaging there have been no reports to date. Fortunately, TPL imaging has proved to be more than adequate for detecting GNRs in biological samples (see below). 26.4.3 TPL and Other Nonlinear Optical Properties GNRs exhibit several nonlinear optical (NLO) properties when excited by ultrashort (femtosecond) laser pulses. Two-photon excited luminescence (TPL), hyper-Rayleigh scattering (HRS), and second harmonic generation (SHG) have all attracted much attention for their capacity to produce optical contrast at visible wavelengths using NIR excitation, with minimal background fluorescence. TPL involves the simultaneous absorption of two photons (typically in the NIR range), followed by a three-step process: (1) excitation of electrons from the d to the sp band to generate electron–hole pairs, (2) scattering of electrons and holes on the picosecond time scale with partial energy transfer to the phonon lattice, and (3) electron–hole recombination resulting in photoemission [121]. Like linear PL, the intrinsic TPL efficiency of bulk gold is poor but can be greatly amplified by resonant coupling of the incident excitation with the LR mode (Figure 26.5A) [97, 98, 122]. The two-photon absorption cross section of GNRs (ca. 15 × 50 nm) is on the order of 2000 GM, intermediate between that of typical dye molecules (∼102 GM) [123, 124] and semiconductor quantum dots (∼104 GM). [125]. Other Au nanostructures such as nanospheres [126], nanoplates [127], nanoshells [128], and nanoparticle dimers [129, 130] have also been found to exhibit TPL activity, but GNRs likely have the highest TPL intensity per unit volume, as well as the added feature of polarization-dependent excitation [97, 131]. The TPL intensity of GNRs is quadratically dependent on excitation power (Figure 26.5B) but has little correlation with excitation frequency or the emission spectra, which are broad and contain several peaks associated with interband transitions within Au (Figure 26.5C) [97]. The TPL intensity of single GNRs also has a cos4 dependency on the polarization of incident light, another characteristic of a nonlinear absorption process [97]. The secondorder polarizability of GNRs has also been measured by hyper-Rayleigh scattering (HRS) [132, 133], sometimes called two-photon Rayleigh scattering due to the frequency-doubled nature of the output signal [134]. However, unlike

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1 λex=730nm 0

400 500 600 Wavelength (nm)

λex=780nm 400 500 600 Wavelength (nm)

λex=830nm 400 500 600 Wavelength (nm)

Figure 26.5. TPL generated from GNRs, using a femtosecond-pulsed Ti:sapphire laser: (A) excitation intensities superimposed onto an absorption spectrum; (B) quadratic dependence of TPL intensity with excitation power; and (C) TPL emission spectra from GNRs in aqueous solution, excited at 730, 780. and 830 nm respectively [97].

TPL, the HRS intensity is linearly dependent on excitation power and the incident polarization is retained [134]. Second harmonic generation (SHG) is another plasmon-enhanced NLO response and has been investigated in oriented GNR arrays as a function of excitation polarization and wavelength [135, 136]. Higher-order NLO responses such as four-wave mixing (FWM) and coherent anti-Stokes scattering (CAS) have been observed from short GNRs [137] and long Au nanowires [138], respectively. The FWM signal from GNRs is ∼39 times stronger than anti-Stokes Raman scattering (CARS) signals produced from molecular probes such as melamine [137]. The emergence and characterization of NLO properties in GNRs have provided fertile ground for biological imaging applications, and for good reasons. As mentioned in Section 26.1, NIR illumination has high transmittivity through biological tissues and can achieve greater penetration depth than

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Figure 26.6. Still-frame TPL image of several GNRs (indicated by arrows) passing through a mouse ear blood vessel, several minutes after a tail vein injection [97]. Blood vessel walls in transmission overlay enhanced for clarity.

visible light, and the very low autofluorescence under multiphoton excitation ensures a large signal-to-background ratio. Furthermore, NLO signal intensities are highly power dependent, which increases three dimensional (3D) spatial resolution and minimizes collateral photodamage. GNRs have been demonstrated as TPL contrast agents both in vitro and in vivo [97]. In a seminal in vivo TPL imaging study, a dilute solution of CTAB-stabilized GNRs were delivered into an anesthetized mouse by tail vein injection, then detected some minutes later passing through ear blood vessels after dilution in the blood pool (Figure 26.6) [97]. Continuous TPL monitoring revealed that the GNRs were cleared from the bloodstream within 30 minutes, presumably due to opsonization. Subsequent in vivo TPL studies with PEG-conjugated GNRs indicate a much longer blood residency time with a circulation half-life of several hours, in line with other recent reports involving PEG-coated GNRs [89, 96]. A three-dimensional TPL imaging modality has been developed for tissues using GNRs as contrast agents, with penetration depths up to 75 μm in a tissue phantom [42]. TPL imaging has been particularly useful to demonstrate the targeted delivery of GNRs to bacterial pathogens [28], tumor cells [19, 99], and activated macrophages [139]. GNRs coated with PEG chains by in situ DTC formation were not taken up by KB cells, as characterized by the near-absence of TPL signals [30], but GNRs conjugated with folic acid derivatives could be targeted to the high-affinity cognate receptor overexpressed on the surfaces of tumor cells. These were observed by TPL imaging to accumulate on the outer cell membrane for many hours, prior to their receptor-mediated endocytosis and delivery to the perinuclear region (Figure 26.7) [19, 99]. This image-guided

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(A)

(B)

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(C)

Figure 26.7. Targeted adsorption and uptake of folate-conjugated GNRs (red) by KB cells overexpressing folate receptors (imaged in transmission mode, grey) [99]. (A) Folate-conjugated GNRs were observed on the surface of KB cells with high density, after 6-h incubation at 37 °C. (B) GNRs were internalized into KB cells and delivered to the perinuclear region after 17-h incubation. (C) No GNR binding or uptake was observed by NIH-3T3 cells, which express folate receptors at a low level. Bar = 10 μm. (Reproduced with permission of Wiley-VCH.) (See color insert.)

delivery provides an opportunity not just to identify and target cancer cells for photothermally induced cell death (Section 26.5), but also to time the delivery of NIR dosage for maximum efficacy. 26.4.4 Surface-Enhanced Raman Scattering (SERS) The tunable plasmonic responses of GNRs have also been applied toward SERS, which is based on the Raman vibrational modes of chemical species adsorbed onto the surfaces of plasmon-resonant nanoparticles. In SERS, the normally weak Raman intensities can be amplified many orders of magnitude by local electromagnetic fields generated by the surface plasmons, which in turn are highly dependent on the supporting nanostructure [140,170]. In addition to local field enhancement effects, the electronic interaction between the Au substrate and absorbed molecules can also contribute to SERS, commonly referred to as a chemical enhancement effect [141, 142]. The SERS activities of GNRs have been investigated extensively and can support signal enhancement factors ranging from 107 to 109 when using an NIR excitation source [63, 143–151]. However, at present there are only a few examples of GNR-based SERS for bioanalytical applications. GNRs have been examined as SERS tags for in vitro cancer diagnostics, using antibodies (anti-EFGR) to select for oral carcinoma cells distributed in a population of healthy cells (Figure 26.8) [152]. Interestingly, it was claimed that CTAB provides a convenient Raman signature for SERS-based imaging. GNRs functionalized with oligopeptide ligands have also been used to detect nuclear translocation events by SERS [27]. With respect to in vivo SERS imaging, Au nanoparticles functionalized with antibodies and Raman-active dyes can serve as especially bright immunolabels based on surface-enhanced resonance

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HaCat normal cells

HSC cancer cells

(B)

Bright field

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Intensity

520

40 0 60 0 80 0 10 0 12 0 00 14 0 16 0 0 18 0 00

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Raman shift (cm-1) 30000 20000 10000 0

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Figure 26.8. Antibody-functionalized GNRs for SERS imaging [152]. (A) Bright- and darkfield images of anti-EGFR-conjugated GNRs on normal human keratinocytes (HaCat) and malignant squamous carcinoma (HSC) cells. (B,C) SERS spectra of GNRs incubated with HaCat and HSC cells, respectively. (Reproduced with permission of the American Chemical Society.)

Raman scattering (SERRS) [153]. Multiplex SERRS imaging with GNRs has been demonstrated very recently, using three different Raman-active dyes [154].

26.5

GNRS AS PHOTOTHERMAL AGENTS

GNRs are capable of producing intense photothermal effects, a property not typically associated with conventional imaging agents or fluorophores. The concept of using NIR-absorbing nanoparticles as highly localized heat sources has inspired a global research effort to develop photothermal agents for the treatment of cancer and other pressing concerns in human health. Here we survey recent studies in which the photothermal properties of GNRs have been applied for targeted cell death (photothermolysis). In particular, it is worth noting that more than one mechanism can be used to induce cell death, and the precise placement of these photothermal agents strongly influences the relative efficiency of the photothermolytic pathways. 26.5.1 Photothermal Action of Plasmon-Resonant Nanoparticles Metal nanoparticles are well known to be efficient converters of light energy into heat, and numerous in vitro examples have been reported using spherical

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Au nanoparticles conjugated to biomolecular recognition elements. The threshold laser fluence required for photoinduced damage can be remarkably low: one study demonstrated that antibody-labeled Au nanoparticles targeted toward CD8+ lymphocytes could induce cell death with a single 532-nm laser pulse of 0.35 J/cm2 [155]. However, NIR irradiation is preferred due to its greater penetration depth into biological tissue [13], as illustrated by the use of NIR-active gold nanoshells for in vivo photothermal imaging and therapy in a tumor mouse model [156], and also more recently by GNRs [95, 96]. With respect to photothermolysis at the cellular level, GNRs have been targeted against tumor cells [19, 41, 95, 99, 157–160], parasitic protozoans [35], macrophages [34], and pathogenic bacteria [28, 161]. Despite the intense interest in this subject, in vitro demonstrations of GNR-mediated photothermolysis are in fact quite recent, the earliest report having been published in 2006 [41]. In many of these studies, the tacit (or sometimes explicit) assumption has been that GNRs induce cell death (necrosis) by photoinduced hyperthermia, in which a few degrees is sufficient to cause cell and tissue malfunction. However, a closer inspection of the photophysical properties of GNRs suggest alternative mechanisms for photothermally induced cell injury and death. In particular, the thermalization of the phonon lattice occurs on the picosecond time scale [162], which can lead to superheating and localized cavitation effects [163, 164]. The scenario above suggests that GNRs can be targeted to tumor cell membranes and serve as “optoporation” agents, and induce cell necrosis by disrupting homeostasis. This has been investigated using folate-conjugated GNRs targeted to KB cells, in which cell necrosis was correlated with a compromise in membrane integrity and an intracellular influx of Ca ions (Figure 26.9) [99]. NIR irradiation in Ca-free media did not lead to any observable

(A)

before

(B)

after

(C)

Figure 26.9. “Optoporation” of tumor cell membranes by photothermally active GNRs [99]: (A,B) folate-conjugated GNRs (red) targeted to the membranes of KB cells, before and after a 1-min exposure to a scanning NIR laser (12 J/cm2); (C) ethidium bromide (red) and a Ca-sensitive dye (green) indicate a loss of membrane integrity and high levels of intracellular Ca2+, respectively. (Reproduced with permission of Wiley-VCH Publishing.) (See color insert.)

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changes in cell morphology, but the introduction of millimolar levels of Ca resulted in a dramatic membrane blebbing. 26.5.2 Subcellular Localization of GNRs If the inception of cell necrosis in Figure 26.9 is attributable to a loss of cell membrane integrity, then it stands to reason that the effect can be optimized by delivering GNRs with subcellular precision. This is indeed possible with folate-conjugated GNRs, which can accumulate on the cell membrane for many hours prior to intracellular uptake (Figure 26.7). It was subsequently demonstrated that the localization of the GNRs strongly affected their ability to mediate NIR-induced photothermolysis [99]. Using a continuous-wave laser source, the threshold power needed to induce membrane blebbing in KB cells with membrane-bound GNRs was determined to be 6 mW (a fluence of 24 J/cm2), whereas cells with internalized GNRs experienced blebbing only after exposure to a laser power of 60 mW, a difference of an order of magnitude. Furthermore, exposing tumor cells with membrane-bound GNRs to ultrashort NIR laser pulses resulted in bleb formation at a threshold power of 0.75 mW, or a fluence of only 3 J/cm2. The reason for this difference has been attributed to the ultrafast electron dynamics involved in plasmon-mediated heating [162], and the greater absorption efficiency of GNRs under pulsed conditions. The subcellular localization of internalized GNRs in the study above was not specifically addressed, but it is reasonable to assume standard cell uptake mechanisms such as receptor-mediated endocytosis. TPL imaging suggests the endosomal transport of unfunctionalized (CTAB-stabilized) GNRs toward the perinuclear region, and also their compartmentation into inclusion bodies several days after initial exposure (Figure 26.3) [30]. TEM analysis of internalized polyelectrolyte-coated GNRs also suggest their residency within endosomes [32, 158]. GNRs can also inflict photothermal damage on particular organelles by changing the targeting strategy, thereby enabling more selective forms of cell death. GNRs functionalized with cysteine-appended octaarginine (R8) peptides have been introduced into immunostimulated mice by intraperitoneal injection and found to be sequestered by activated macrophages, as indicated by TPL imaging and immunofluorescent staining using F4/80+ antibodies (Figure 26.10) [139]. The photothermally active GNRs could induce either necrosis or apoptosis, the latter by NIR irradiation at a low power density and fluence (2.2 W/cm2 and ∼1 J/cm2, respectively). The apoptotic response was correlated with Annexin-V labeling and also with the production of reactive oxygen species, suggestive of mitochondrial dysfunction. Mitochondrial damage was confirmed by the loss of fluorescence response to MitoTracker Red, a membrane-potential sensitive dye. It is worth mentioning that, unlike the receptor-mediated uptake of folate-conjugated GNRs, the phagocytosis of GNRs was very rapid, with high levels of uptake after only 30 minutes.

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(A) F4/80 antibody

523

(B) R8-NRs

Figure 26.10. In vivo uptake of R8-conjugated GNRs by macrophages, 30 min after intraperitoneal injection [139]: (A) immunofluorescence staining of F4/80+ macrophages (blue) and (B) TPL image of R8-GNRs (red) inside macrophages (bar = 10 μm). (Reproduced with permission of the authors.) (See color insert.)

26.5.3 Photothermally Activated Drug Release The photothermal properties of GNRs suggest another attractive therapeutic application: molecular release triggered by NIR light. The photoinduced release of DNA adsorbed onto GNR carriers has been reported by several groups [165–167]. The release mechanism has been attributed to the reshaping of GNRs [165, 167] or to the photoinduced desorption of chemisorbed DNA from the GNR surface [166]. In all cases the released DNA remains biologically active, as demonstrated by subsequent transfection studies resulting in gene expression [166]. Polyelectrolyte-coated GNRs are also candidates for photothermally activated drug delivery and release. Thermoresponsive polymers such as poly(Nisopropylacrylamide), or PNIPAAm, can be induced to contract in response to light in the presence of GNRs [168]. Photoresponsive GNR–PNIPAAm hydrogels are presently limited by the uneven distribution of GNRs within the hydrogel matrix, resulting in a nonuniform photothermal response. However, PNIPAAm polymer brushes can be grown directly from GNR via surfaceinitiated atom transfer radical polymerization [169]. These polymer-grafted GNRs have well-defined core–shell structures and overall dimensions below 100 nm, and may be appropriate materials for targeted intracellular delivery.

26.6

CONCLUSION

In the past few years, we have witnessed remarkable progress in the development of surface-functionalized GNRs for biophotonic applications. Surface chemistry will clearly continue to play a critical role in advancing the utility

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of optically and photothermally active GNRs for biological imaging, theranostics, and other biomedically relevant objectives. The targeted delivery and subcellular localization of multifunctional GNRs may also prove to be invaluable for fundamental studies in cell biology. In particular, GNRs guided by TPL imaging may also be useful as photothermal probes toward any number of intracellular processes, possibly resulting in new and unexpected discoveries in cell and developmental biology.

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CHAPTER 27

Quantum Dot Labeling for Assessment of Intracellular Trafficking of Therapeutically Active Molecules DIANE J. BURGESS and MAMTA KAPOOR School of Pharmacy, University of Connecticut, Storrs, Connecticut

27.1

INTRODUCTION

Apart from their applicability as photovoltaic and light emitting devices, quantum dots (QDs), have gained tremendous popularity in biology over the past couple of decades due to their photostability, wide absorption spectra, narrow emission spectra, and high quantum yield. Consequently, these have been utilized for visualizing therapeutically active molecules in vitro and in vivo for a better understanding of their localization and bio-distribution in cells and live animals, respectively. This chapter discusses QD assisted intracellular trafficking studies of various biomolecules. The topics included are: Introduction to intracellular trafficking and quantum dots, cellular barriers, QDs versus organic dyes, synthesis and properties of quantum dots, methods of tagging QDs to therapeutically active molecules, intracellular trafficking of QD-linked molecules, intracellular trafficking tools, in vivo QD imaging and, cytotoxicity of QDs. Therapeutically active molecules, such as peptides, genes (DNA/RNA), antibodies, nutrients, and drugs, are often are required to be delivered to a specific target site in the body within a certain concentration range. The target site can be an area, a specific organ, or specific cells and even an Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

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organelle or region within a cell. To understand the delivery of therapeutically active molecules at the cellular level, suitable markers are used to label the therapeutic agent as well as the cell or organelle. An in-depth knowledge of the cellular distribution (intracellular trafficking) is often crucial to estimate pharmacokinetic and pharmacodynamic behavior as well as to understand and hence improve delivery to the target site. Therapeutic agents encounter a number of barriers prior to reaching their target cell organelle. Uptake through the cellular membrane can occur via direct fusion or endocytosis. Most cells have an endocytic pathway and the therapeutic agent and its carrier system as appropriate (such as liposomes or nanoparticles) (the cargo) can enter the cells via endocytic vesicles (early endosomes). From the early endosomes, the cargo enters late endosomes and this progresses to highly acidic lysosomes where the cargo would be degraded. Accordingly, it is usually important that the cargo escape from endosomes in order to exhibit the therapeutic effect. Exceptions to this are agents that have their effect in the lysosomes such as the ones used for treatment of lysosomal storage disease. After release from the endosome, the molecule must reach the target organelle before it is destroyed by any degrading enzyme in the cytoplasm. Thus for a clear understanding of the intracellular barriers, there is a need for specific labels, which can aid in microscopic visualization of these biomolecules intracellularly. Until now, researchers have been using organic dyes for cell trafficking studies via organelle labeling. These dyes label the cell organelles with different colors to facilitate distinguishing these [1]. Accordingly, the cellular entry and the cellular path can be monitored. Several dyes have been used in the past to stain various membrane organelles such as the plasma membrane (FM 4–64 FX [2]), the cytoskeleton (Alex flour 568-phalloidin [3]), the endoplasmic reticulum (ER Tracker [4]), the nucleus (Hoechst 33342 [5]), the microtubules (Tubulin Tracker green [6]), the mitochondria (mitotracker [7]), the Golgi body (Golgi-EGFP [8]), the endosomes (FITC-dextran [9]) and the lysosomes (lysosensor green [10]). The use of organic dyes has not been proved to be very useful for long-term trafficking studies since these are prone to photobleaching. Consequently, long-term same-cell imaging that would otherwise allow tracking of various delivery vectors and their interaction with cellular organelles has not been possible. Instead, research has been compromised by imaging different groups of cells at different time points and therefore critical information in the trafficking process is lost. Although valuable insights have been achieved via PCR analysis of the various cellular organelle fractions, this method only provides information on the plasmid DNA and not the delivery vectors. In addition, PCR is performed on bulk cell populations and only reports the presence of the plasmid, but not its functionality. Consequently, there is a need for more reliable markers, such as quantum dots (QDs), in place of organic dyes, which are not subject to photobleaching

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and enable tagging of large molecules such as plasmid DNA as described by Srinivasan et al. [11]. Quantum dots (QDs) are semiconductor nanocrystals that have bright illumination, photostability, expansive absorption spectra, and size-tunable emission spectra. This makes these moieties widely applicable for intracellular trafficking [11–13], live cell imaging [14–16], and cancer targeting [17, 18]. For this purpose, QDs have been tagged to a variety of biomolecules such as transferrin [19–21], folate [22, 23], siRNA [24–26], DNA [27–32], peptide [17, 33–41], neurotransmitters [16, 42, 43], drugs [44, 45], and even pathogens [46, 47]. QD visualization has also been made possible by resonance transfer techniques such as FRET (fluorescence resonance energy transfer) [32, 48–55] and BRET (bioluminescence resonance energy transfer) [56, 57]. Quantification of QD fluorescence can provide valuable insight into the barriers to efficient delivery and facilitate development of more efficient delivery systems [56–59]. Also, numerous reports in the literature indicate that quantum dots are safe to use [60, 61]; however, we are still unsure about their use in in vivo. This chapter will summarize the properties of QDs, techniques for QD labeling of biomolecules, intracellular trafficking studies of QD assisted molecules, and in vivo QD imaging and toxicity issues. Due to their outstanding properties, the future of these tiny nanocrystals seem to be really bright.

27.2

BARRIERS TO CELLULAR ENTRY

Cell membranes pose the first and foremost barrier to molecules entering cells. There are two main pathways for cellular uptake—endocytic (phagocytosis, pinocytosis) and non endocytic pathways (microinjection, electroporation) [62, 63]. Phagocytosis exists in some specialized cells such as macrophages, neutrophiles, and monocytes, whereas pinocytosis occurs in all cells. Pinocytosis can be a caveole-dependent, clathrin-dependent, macropinocytosis, or clathrin- and caveolae-independent pathway. It is believed that the molecules adapt to the pathway depending on their size and composition. For example, large molecules of size 5 μm enter through macropinocytosis, while particles of 60 nm, 120 nm, and 90 nm adopt a caveolae-dependent, clathrin-dependent, or clathrin- and caveolae-independent pathway, respectively [64]. Caveolae have caveolin proteins toward which certain ligands such as SV40 have specific affinity. It is believed that substrates from caveosomes (caveolae endosomes) directly enter Golgi or endoplasmic reticulum, bypassing the lysosomal degradation [65]. The clathrin-dependent pathway is for receptor-mediated endocytosis for molecules such as transferrin (Tf) and folic acid, which have affinity for

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transferrin (Tf) and folate receptors, respectively, that are present excessively in tumor cells. These endosomes (called sorting endosomes) can either behave as recycling endosomes, where they go back to the membrane and exocytose the cargo, or transfer it to late endosomes and then to lysosomes, where it is degraded. The cargo needs to be released from the early or late endosome to prevent it from degradation (endosomal escape). For the same, helper lipids such as DOPE [66] are sometimes used in the cargo to help in endosome destabilization and therefore aid in their escape from the endosomes. Besides, other pH-sensitive lipids or polymers are used that get protonated and break the endosomal membrane, at acidic pH [67, 68]. The pH in the early endosome is 6.2 that decreases to 5.5 in late endosomes and to 4.5 in lysosomes due to the proton pump operating in the endosomal membrane. Microtubules have also been considered to be responsible for the transfer of molecules from the early to late endosomes [69]. Besides, interleukin-2 receptors on lymphocytes adopt clathrin- and caveolae-independent pathways [64]; the exact mechanism for the use of this particular pathway is still unclear. Unlike other biomolecules, plasmid DNA needs to enter the nucleus for transcription and translation processes to happen. Transfer of plasmid DNA

(A)

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Figure 27.1. Endocytic pathway for uptake of DNA lipoplex (cargo) from cells. The cargo can enter via (A) caveolae-dependent, (B) clathrin-dependent, and (C) macropinocytosis pathways. In either case, the DNA is either directly released from the early endosome/caveosome/macropinosome (unknown), or is transferred to late endosomes followed by lysosomes, in the absence of endosomal escape. The release of DNA can occur from late endosome as well (unknown). Besides, depending on the nature of the cargo, it can also be recycled back to the surface by recycling endosomes. (Reproduced with permission from: Ref. 73.)

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into the nucleus may occur via passive DNA entry during cell division and/or by active processes via pores in the nuclear membrane [62, 70]. Transport across the nuclear membrane via a nonnuclear pore pathway in cells that are not undergoing active cell division has also been reported [71] for DNA–lipid complexes following conjugation with transferrin. Although the mechanisms of nuclear uptake of plasmid DNA are known, it remains unclear why the uptake efficiency is low and varies with cell type as well as transfection conditions (e.g., in vitro versus in vivo) [68, 72]. Figure 27.1 illustrates various possible endocytotic pathways for uptake of liposome–plasmid DNA complex followed by its nuclear entry.

27.3

QUANTUM DOTS VERSUS ORGANIC DYES

During the past several decades, quantum dots have gained immense importance due to their photostability, extremely bright illumination compared to organic dyes, wide absorption spectra, and tunable emission spectra [74–77]. Consequently, visualization of multiple colors at the same time has been possible, which has made intracellular trafficking studies relatively uncomplicated [78]. Due to the wide absorption spectra, multiple color QDs can be excited by a single laser source, thus averting overheating of the cells in in vitro cell culture studies. Also, in the case of QDs, there is a vast gap between wavelength of excitation and emission, which makes QD-based visualization far more sensitive by cutting down the autofluorescence. As mentioned previously, QDs are less susceptible to photobleaching compared to the organic fluorophores, thus enabling their visualization from hours to as long as a week [34]. Our lab has performed QD imaging for intracellular trafficking for 24 hours at a stretch [11]. As shown in Figure 27.4 (later section), QD conjugated DNA can be visualized in the perinuclear region for 24 h. We also compared the photostability of rhodamine conjugated DNA with QD-DNA and from Figure 27.2 it can be seen that, unlike the QDs, the rhodamine stain vanishes in as little as 50 min. In another study by Watson et al, [79], QDs stained nicely for 3 min of continuous laser exposure while the organic dye photobleaches. Additionally, unlike some organic dyes, QDs are not pH and temperature sensitive [10]. These can be visualized at acidic, basic, and neutral pH. Again, organic dyes seem to be diffused when these are in the cytoplasm, which gives a lot of background fluorescence, thereby making the imaging of specific organelles difficult. The issue of autofluorescence also comes with well-known fluorophores such as GFP. QDs of different sizes (different colors) can have similar optical properties if they have the same core material shell as well as the same surface chemistry, which is not possible in the case of organic dyes, all bearing different functional groups on the surface. Finally, the full-width half-maximum (FWHM) of QDs is around 30 nm, which is much lower than that of organic dyes (50– 100 nm), thereby reducing the chances of spectral overlap [76].

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QD

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Figure 27.2. Comparative study of photostability of QD-DNA conjugated with rhodamine-PE-DNA conjugate (organic dye complex). QDs can be visualized even after 100 min of laser exposure while most of the rhodamine bleaches after 50 min, in CHOK1 cells. The graph (G) represents the fluorescence intensity time plot for the two conjugates after 100 min of continuous laser exposure at 543 nm. QD-DNA conjugates were added as 1.469 × 1013 particles per 2.5 × 106 cells per well. (Bar = 25 μm) (From Ref. 11.) (See color insert.)

27.4

QUANTUM DOT: PROPERTIES AND SYNTHESIS

27.4.1 Properties Quantum dots (QDs) are semiconductor nanocrystals belonging to II–VI (CdSe, CdS, CdTe), III–V (InSb, InAs, GaSb, GaAs), or IV–VI (PbS, PbSe,

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541

PbTe) column elements. QDs are usually 1–10 nm in radii, making these tiny crystals even smaller than the Bohr exciton radii (50 Å for CdSe) [80, 81]. Consequently the electrons of the bulk material (semiconductor quantum dot) are confined to a small three-dimensional (3D) space (quantum confinement) and the bandgap (gap between conduction and valence band) strictly depends on its size [82]. The smaller the size, the smaller is the gap, and the shorter is the emission wavelength. Therefore blue QDs are the smallest in size, while red ones are the largest. Additionally, QDs can be excited from the same laser source while they emit at different wavelengths depending on their size. QDs also are known for their “blinking” behavior (switch “on,” switch “off”) when continuously excited due to Auger ionization [83, 84]. This property might be useful to distinguish QD aggregates from single QDs since it is unlikely that the aggregates will clearly demonstrate the blinking behavior [85]. Blinking may be a disadvantage during quantification studies using confocal microscopy; however, this is not expected to interfere in flow studies. 27.4.2 Synthesis QDs with heavy metal (semiconductor) core shells are synthesized under conditions of very high temperature (250–400 oC) in organic solvents usually in the presence of surfactants (TOPO, trioctyl phosphine oxide) to obtain hydrophobic nanocrystals. The detailed methods have been described elsewhere [86–89]. The heavy metals, especially Cd, are prone to oxidation and Cd forms Cd2+ ions, which are toxic. Therefore these shells are commonly passively coated with ZnS to prevent the diffusion of oxygen [60]. Additionally, this coating improves the photoluminescence since it overcomes the negative effect of metal defects on the quantum yield [90].

27.5 QD LABELING TECHNIQUES FOR THERAPEUTICALLY ACTIVE MOLECULES The surface of the QDs can be modified in several ways, which usually serves either or all of the following purposes: (1) renders quantum dots water soluble, (2) introduces functional groups for conjugation to biomolecules, and (3) further passivates the surface to prevent leakage of heavy metal ions (Cd2+) and to improve biocompatibility. Quantum dots are sometimes capped with mercaptoacetic acid (MAA) or dihydrolipoic acid (DHLA), which involves a disulfide linkage, to make them water soluble [91]. Additionally, these moieties also introduce –COOH functional groups onto the surface, thus acquiring a negative charge under basic pH, which enables conjugation of these molecules with the cationic species through electrostatic interactions. The –COOH group can be also used to covalently conjugate QDs with biomolecules via a –NH2 linkage [53].

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L+ –O A O

S

R COOH Thiol NHS ester

MAA O

P O

P

O

P

P

R

E X-L

COVALENT INTERACTION

P

O

P

O O

P

O P

P

P

P TO O

P

BIOTIN

O O

O

P

BIOTINYLATED LIGAND

L D

SS

O

V

O

PHOSPHOLIPID (PL)

P

O

ST

NONCOVALENT INTERACTION

O

CdSe

O

L, LIGAND

P

O

TO P

R

O

O

O

L

VP STO

S

O

HYDROPHOBIC INTERACTION

O

P

PO

C

P

TO

P TO

X NH2 MALEIMIDE NH2

S S

DHLA

O O– + L

ELECTROSTATIC INTERACTION

B

Figure 27.3. Illustration of various techniques for bioconjugation of therapeutically active molecules (ligands) with CdSe quantum dots passively coated with ZnS (blue). Disulfide linkage of ZnS coating with (A) mercaptoacetic acid (MAA) or (B) dihydrolipoic acid (DHLA) enabling electrostatic interaction with cationic ligand (L+). TOPO (trioctyl phosphine oxide) coating over ZnS enables hydrophobic interaction with phospholipids (C). Phospholipids functionalized with STV can be linked to biotinylated ligand via noncovalent interaction (D). This bond is unaffected by pH and other environmental conditions. Carboxylated, thiolated, or NHS ester group functionalized lipids can conjugate with amine or maleimide containing ligands (E).

Solubilization in aqueous environments can also be achieved by QD micellization in phospholipids/block copolymers via hydrophobic interaction between TOPO and the lipids/polymers. This approach gives broad flexibility in surface functionalization of quantum dots. Various functional groups have been introduced on the surface of QD micelles by using appropriate lipids/ polymers. Groups such as amine [12], maleimide [11], streptavidin (STV) [31], or carboxyl acid [24] have already been attained on QD surfaces to facilitate conjugation with therapeutically active molecules. QD-streptavidin (QD-STV) has been used to conjugate to biotinylated molecules [41]. Figure 27.3 illustrates various ways of QD conjugation with therapeutically active molecules. The ligands can also be conjugated to QD surfaces via ligand exchange, wherein the passive coating material (usually ZnS) is displaced by the attaching ligand [21]. Although those techniques maintain the size of QDs, there is a question as to QD stability and safety, which is normally guarded by the passive coating.

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Conjugation to quantum dots must not affect the biodistribution of the attached active molecule. Several studies have shown that this does hold true [12, 30, 92, 93]; however, if care is not taken, QDs can impair functionality of the biomolecule [42], the probable reason being the steric hinderance by large sized QDs tagged to small molecules. This problem can be taken care of by using a PEG spacer [16]. The conjugation techniques mentioned above open the door to numerous ways that can be used to attach a wide variety of molecules to quantum dots. Based on functionalization, there can be a change in QD solubility that might lead to aggregation, thereby affecting cellular uptake. One way is to coat QDs with PEGylated lipids, which help to keep them dispersed in the medium. Our lab has formulated QD micelles via TOPO associated coating with PEG lipids. PEG-QDs were successfully prepared and tagged to DNA via SPDP-labeled PNA linker [11]. The in vitro studies showed that the QD labeling did not affect the transfection efficiency of plasmid DNA associated delivery systems. Invitrogen [94] and Evident technologies [95] provide quantum dots that are PEGylated terminally functionalized with carboxyl, streptavidin (STV), or amine groups that can easily be conjugated to biomolecules using simple chemistry steps. Additionally, commercially available near-infrared (NIR) dots are useful for visualization of QDs in deep tissue and in in vivo conditions.

27.6 QDS FOR INTRACELLULAR TRAFFICKING OF THERAPEUTICALLY ACTIVE MOLECULES Quantum dots have been an exciting tool for trafficking of active molecules intracellularly, the finest benefit being their photostability and high sensitivity. Therefore these have proved to be very useful to reveal information on the localization of biomolecules inside the cell, thereby providing information on biodistribution. This helps in estimating their effective dose and in gaining a clear understanding of their PK/PD. Several molecules such as DNA, siRNA, peptides, enzymes, hormones, antibodies, neurotransmitters, liposomes, as well as targeting ligands such as folate and transferrin (Tf), pathogens, and drugs (e.g., anticancer, anti-inflammatory) have been successfully tracked using quantum dot labels. Examples in each category of biomolecules are discussed below. 27.6.1 DNA Genetic disorders, whether single gene (cystic fibrosis, sickle cell anemia) or multiple gene defects (cancer, hypothyroidism, Alzheimer disease), need to be treated by gene replacement therapy. Here the defective gene (plasmid DNA) needs to be supplied from outside. The plasmid DNA is required to enter the nucleus in order to undergo transcription (DNA to RNA) and then

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translation (RNA to proteins). Naked DNA itself cannot be delivered into the cells and thus needs to be protected from the cytoplasmic nucleases with the help of delivery vectors, which can be viral or non viral. Due to immunogenicity, and potential toxicity, nonviral vectors such as cationic and anionic liposomes are the preferred means to deliver the DNA. Cationic liposomes, being positively charged, are toxic to the cells since they can bind to the negatively charged plasma proteins and eventually kill the cells [96]. Anionic liposomes are therefore safer to use. Our lab has successfully prepared anionic lipoplexes that have been shown to be safe and effective for DNA delivery [97–99]. DNA usually gets separated from the liposome before it enters the nucleus. Exactly where this step happens is still unknown. If this happens too early, much before the DNA enters the nucleus, the DNA might degrade. Therefore it is important to track the DNA until its entry into the nucleus. Our group has also successfully demonstrated DNA degradation on Chinese hamster ovarian cells (CHOK1 cells) using DNA double labeled with organic dyes [100]. Our future work will focus on demonstrating the same using quantum dots. Much work has been done on the intracellular trafficking studies of quantum dots tagged with plasmid DNA [11, 13, 27, 31, 49, 53]. More than a decade ago, Alivisatos et al. [82] prepared ssDNA tagged with gold particles. Shortly thereafter, Dubertret et al. [12] attached QDs to ssDNA, which was then hybridized to a complementary strand to form QD-duplex DNA conjugates, thereby demonstrating that QD conjugation does not affect the hybridization process. Mitchell and co-workers obtained a QD-DNA conjugate via disulfide linkage between 3-mercaptopropionic acid coating of CdSe core shell quantum dots and thiolated DNA [29]. Crut et al [27] detected single DNA molecules using multicolored quantum dots conjugated to two ends of the DNA sequence via biotin–STV or digoxigenin–anti-digoxigenin linkages. This study was also helpful in studying DNA–protein interactions [27]. Xiao and Barker [31] prepared QD-DNA via STV–biotin linkage and successfully imaged DNA in metaphase chromosomes via FISH (fluorescence in situ hybridization) [31]. Our group demonstrated plasmid DNA in the nuclear/ perinuclear region using red QDs in as early as 6 h using lipofectamine 2000 as the delivery vector (Figure 27.4). Unlike a physical mixture of QD and DNA, this effect was seen only when QDs were covalently conjugated to plasmid DNA via a PNA–SPDP (peptide nucleic acid (PNA)–N-succinimidyl3-(2-pyridylthio) propionate) linker [11]. 27.6.2 Antisense Oligonucleotides Like DNA, antisense oligonucleotides have also been visualized using QDs. For example, Jia et al. [101] used mercaptoacetic acid coated CdTe quantum dot conjugated antisense oligodeoxynucleotides, which were linked to carbon nanotubes through PEI via electrostatic interactions between positively charged PEI and negatively surfaced nucleotides and nanotubes.

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(A)

6h

(C)

10 h (B)

545 (E)

24 h (D)

(F)

Figure 27.4. Confocal images of Lipofectamine 2000 mediated transfection studies of QD-DNA conjugates (red) 6 h, 10 h and 24 h postincubation, delivered in CHOK1 cells stained with SYTO-16 nuclear stain (green). Yellow regions in the nuclear/perinuclear region determine the efficient transfection of QD-DNA conjugates added as 1.469 × 1013 particles per 2.5 × 106 cells per well. (Bar = 25 μm). (From Ref. 11.) (See color insert.)

27.6.3 siRNA siRNA (silencing RNA) is a recently explored area for the treatment of diseases such as cancer [102], hyperapolipoproteinemia [103], respiratory syncytial virus (RSV) [104], and Huntington disease [105]. Many companies have made remarkable progress in this area and consequently there are a number of products in clinical trials [106]. siRNA is usually delivered as doublestranded RNA (sense and antisense) consisting of 20–25 nucleotides. Inside the cell, they form an RISC (RNA-induced siRNA complex) with Ago-2 protein. Complex activation occurs, thereby unwinding the siRNA and the sense strand leaves. The antisense strand participating in the complex formation guides it to complementary mRNA, binds to it, and gets cleaved, thereby blocking the translation process and hence the protein expression. This is called the “gene knockdown effect.” siRNA, like DNA, is prone to degradation by enzymes in the cytoplasm and thus needs to be monitored for delivery in the cytoplasm, which can occur by using powerful markers such as quantum dots. Several groups have worked on tagging siRNA with quantum dots. Bakalov et al. [24] conjugated carboxylic group functionalized quantum dots with amine modified single-stranded 21–23 mer oligoribonucleotides (siRNA)

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via EDAC coupling (EDAC: 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide, hydrochloride) and used it as a hybridization probe to determine siRNA transfection efficiency. Chan et al. [107] prepared QD–siRNA conjugates via biotin–STV linkage and used different colored QDs to image different mRNA targets using FISH technology in mouse brain. Yezhelyev et al. [108] prepared siRNA conjugated to QDs, which not only helps in intracellular trafficking studies but also assists in endosomal escape of siRNA via the “proton sponge effect,” at low pH, which excludes the requirement of pH sensitive lipids or fusogenic peptides for endosome destabilization. Derfus et al. [26] conjugated amine modified QDs with thiolated siRNA using sulfo-SMCC cross linker and quantified QDs to account for the siRNA transfection efficacy. Qi and Gao [109] prepared cationic polymer coated QDs attached to siRNA via electrostatic interactions. With the help of QDs they could see that the complex entering the cytoplasm in 1 h and its separation began at around 1.5 h as indicated by the visibility of FITC dye attached to the siRNA [109]. 27.6.4 Proteins and Peptides Proteins such as albumin, biotin, and STV have been conjugated to quantum nanocrystals. Hanaki et al. [17] prepared QD/sheep serum albumin (SSA) complex that was seen to be internalized in the endosome in Vero cells. The fluorescence could be visualized for 5 days after incubation compared to only for 3 days when similar studies were performed with fluorescein dextran. Another report from the same group found the QDs to be visible for 7 days at a stretch [18]. Bonasio et al. [33] covalently labeled membrane proteins specifically, Integrin lymphocyte function-associated antigen-1 (LFA-1), with quantum dots without affecting the functionality of protein, in 293T cells. For the purpose, they first entrenched cutinase in the extracellular region of the protein, which has affinity for pNPP (p-nitrophenyl phosphonate), cutinase suicide inhibitor. QDs were linked to the protein via QD-pNPP formation using QD-maleimide with pNPP-SH [33]. Quantum dots via STV–biotin linkage have been conjugated to peptide such as neuronal growth factor, which targets neuronal cells, where it binds to tyrosine kinase receptors [38, 41]. Protease labile linkers between QD and gold NPs have been used for QD-based targeted delivery to specific cell types. For this purpose, QDs have been conjugated to gold NPs via a collagenase degradable linker to enable FRET (fluorescence resonance energy transfer) based quenching of QDs by the NPs, where the former is the donor and the latter is the acceptor [110]. QDs have also been utilized for live cell imaging where their endosomal internalization has been visualized using avidin conjugated QDs with prebiotinylated surfaces of HeLa cells [111, 112].

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27.6.5 Targeting Ligand Various peptides such as TAT [14, 15] and NLS [34, 39] have been used as targeting ligands for entering the cell surface and the nucleus. Quantum dots have been very helpful in tracking the path of these useful molecules. Viruses have an inherent ability to enter the nucleus. Nuclear localization sequence (NLS) is a sequence derived from viral proteins, which are responsible for nuclear entry. Quantum dots have been linked to NLS via a STV–biotin linkage and those have been observed in the nuclear/ perinuclear region in HeLa cells [34]. Cell surface targeting ligands such as folate [22, 23] and transferrin (Tf) [19, 113] have also been tagged to quantum dots. Bharali et al. [22] prepared InP-ZnS quantum nanocrystals with folate via DCC coupling which were transfected in A549 cells. Wang et al. [20] synthesized transferrin (Tf)-QD via EDC coupling for fixed cell labeling and this was detected for 5 days.

27.6.6 Liposomes/Polymers Intracellular delivery of a QD-biomolecule requires that the conjugate enters the plasma membrane and then escapes rapidly from the endosome to avoid degradation in the lysosomes. To facilitate endosomal escape, these biomolecule conjugates are often delivered via cationic liposomes [24, 114], pHsensitive polymer [35], or fusogenic peptides [26]. Quantum dots have been coated with PEG-PEI to achieve long-term circulation, cell penetration, and endosomal escape, by the ligand exchange mechanism. It was shown that two molecules of PEG with PEI have better ability to escape from endosomes compared to four molecules; however, the latter was less cytotoxic than the former. Despite the fact that PEG is required in the formulation for long circulation, it is important to optimize the amount of PEG [35]. QDs have also been encapsulated in the lipid bilayer obtained from DMPC : DOTAP : DSPE-PEG lipids without loss of fluorescence [115]. In another case, PEGylated QDs have been encapsulated in lipid vesicles with different lipid concentration to maximize the encapsulation efficacy [116]. Similarly, Chen and co-workers prepared QD and silica loaded liposomes [114]. Wi et al. [117] developed a model for the stability of such QD loaded liposomes depending on their size. Another way of achieving QD–liposome association is by complexing the two via electrostatic interactions [118]. The method of choice for QD–liposome conjugation is still unclear but surface conjugation seems to be less complicated than encapsulation, since the chances of fluorescence quenching are minimal in this case. QD tagged liposomes can be valuable for tracking the delivery vector intracellularly and detecting their separation from the biomolecule.

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27.6.7 Antibodies A relatively recent publication by Xing et al. [119] demonstrated several ways by which QDs can be conjugated to antibodies, namely, SMCC (succinimidyl 4-[N-maleimidomethyl]cyclohexane-1-carboxylate) coupling, EDC (1-ethyl-3(3-dimethylaminopropyl)-carbodiimide) coupling, biotin–STV linkage, hydrazide-oxidized carbohydrate linkage, and histidine–Ni-NTA coupling, where hydrazide and histidine couplings were found to be more efficient than other coupling forms. Such conjugates, if prepared with primary antibodies, can bind to secondary antibodies in the cells and thereby help in performing immunohistochemistry studies [119]. Geho et al. [36] have also prepared QD-STV conjugated to biotin–secondary antibody, useful to detect primary antibodies for immune-staining studies (QD microarrays). QDs when conjugated to IgG antibodies can help detect cancer markers such as Her2 in breast cancer cells [120].

27.6.8 Pathogens Pathogens such as Escherechia coli have been visualized using quantum dots as reporters. This has been achieved by labeling the cells with biotinylated antibodies and binding these to streptavidin-QDs. By using this technique, even a single cell could be detected for hours, compared to organic dye that vanished in a few seconds [46].

27.6.9 Neurotransmitters Quantum dots have also been conjugated with neurotransmitters. Rosenthal et al. [42] linked QDs to the neurotramitter serotonin (5HT) but, unfortunately, this binding impaired ligand binding affinity to its receptor. To resolve this problem, such conjugates were prepared with PEG spacers that had great affinity for the serotonin receptors [16].

27.6.10

Drugs

Besides genes, proteins, and neurotransmitters, quantum dots have also been used to label various small molecule drugs such as anticancer, anti-inflammatory [121–123], and anti-hypertensive [45] drugs. Bagalkot et al. [44] delivered doxorubicin (anticancer drug) to prostate cancer cells using QD-RNA aptamer conjugated to the drug and confirmed the efficacy of this system by using the FRET mechanism. The fluorescence of the QDs is better quenched when tagged with doxorubicin through aptamer, and dequenched once the conjugate breaks [44]. Table 27.1 summarizes numerous examples of QD tagged therapeutically active molecules for cell or in vivo studies.

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TABLE 27.1 Various Molecules Tagged to QDs Via Several Different Linkages for a Variety of Applications Category DNA

Antisense ODN RNA

Molecule Plasmid DNA Plasmid DNA Plasmid DNA

Interaction

Cells/Animal

Application

Reference

CHOK1 cells COS-7 cells Glass surface

DNA trafficking studies Cancer Single DNA molecure detection

11 53 27

Fluorescence imaging FRET FISH

13 49 31

Plasmid DNA Plasmid DNA Genomic DNA

Disulfide Amide STV–biotin, Digoxigenin– Antidigoxigenin Thioether STV–biotin STV–biotin

ODN ODN ss ODN ODN

Amide Carbamate Disulfide Amide

Glass surface HEK293 cells Human metaphase chromosome ND Human sperm cells Xenopus embryo HeLa cells

ssRNA ssRNA siRNA siRNA HER2 siRNA HER2 siRNA siRNA miRNA

Amide STV–biotin Electrostatic Disulfide Electrostatic Electrostatic Thioether STV–biotin

K-562 cells Mouse brain MDA-MB-231 cells HeLa cells SK-BR3 cells SK-BR3 cells 3T3-J2 cells Glass surface

FRET FISH DNA trafficking studies Nanotube mediated intracellular delivery Cancer FISH Cancer Cancer Breast cancer Breast cancer Cancer miRNA profiling microarray

32 30 12 101 24 107 108 26 109 132 25 129

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550 TABLE 27.1 Category Proteins

Continued Molecule SSA, BSA SSA BSA Membrane protein—LFA-1 antigen Nerve growth factor (NGF) WGA

Peptides

Interaction

Cells/Animal

Application

Reference

Surface adsorption Amide Avidin–biotin Amide

Vero cells

Cancer

17

EL-4 T lymphocyte cells HeLa cells 293T cells

Cancer Cancer Cancer

18 37 33

STV–biotin

PC12 cells

Neuronal growth determination

41

Amide

Microbiological labeling

93

Tricosanthin

Amide

PSMA TAT TAT CPP NLS NLS Phytochelatin related peptides Lung-targeting peptide

Amide Disulfide STV–biotin Disulfide STV–biotin STV–biotin Hydrophobic

Gram-positive bacteria—S.aureus JAR cells (human choriocarcinoma) Mice (IV) Cardiac myocytes HeLa cells HeLa cells HeLa cells HeLa cells HeLa cells

Amide

Mice (IV)

HIV infection

125

Prostrate cancer Live cell imaging Live cell imaging/ targeting Efficient intracellular delivery Nucleus targeting Nucleus localization Live cell imaging

136 14 15 40 34 39 130

Lung targeted delivery

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Category Antibody (Ab)

Drugs Pathogens

Neurotransmitters

Miscellaneous

Molecule

Interaction

Cells/Animal

Application

Reference

Anti-Pgp Polyclonal antimouse Ab (IgG) Anti staphylococcal enterotoxin B Doxorubicin Staphylococcus aureus Escherichia coli Serotonin transporter protein inhibitor (SERT) Dopamine

Avidin–biotin Amide

HeLa, AX2 cells MCF-7r breast adenocarcinoma cells

Live cell imaging Immunohistochemistry studies

111 131

Electrostatic

Fluoroimmunoassay

128

Electrostatic Amide

Staphylococcal enterotoxin B LNCaP cells Staphylococcus aureus

Cancer Recognition of pathogens

44 47

STV–biotin STV–biotin

Escherichia coli HEK293 cells

Recognition of pathogens Live cell imaging

46 16

Amide

Hyaluronic acid

Electrostatic

Mouse epithelial A9 cells HeLa, human dermal fibroblast (HDF) cells

Phototoxic drugs and redox specific fluorescence labeling Lymphatic vessel imaging

124 127

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27.7 QD ASSOCIATED TOOLS FOR INTRACELLULAR TRAFFICKING: FISH AND FRET Fluorescence in situ hybridization (FISH) is a valuable tool to study specificity between a probe and chromosomes. Therefore this is very useful for medical purposes. In this technique, labeled DNA sequences are thrown onto the chromosomes and washed after awhile. If the sequence gets hybridized by a complementary sequence on the chromosomes, it is indicated by the label. Due to extraordinary photostability and high quantum yield, quantum dots are the favorite markers for FISH studies. QD-oligonucleotide FISH probes have been used on human sperm cells wherein they hybridize to the Y chromosomes [30]. Xiao and Barker [31] prepared QD-based FISH probes for metaphase chromosomes. QD labeled DNA was used as a hybridization probe for human chromosomes. Similar studies were also performed using HER2, a cancer marker in breast cancer cells. FISH was used to successfully detect the match between the DNA sequence and the chromosomes. It has been shown that these probes are more sensitive than those prepared with organic dyes such as Texas Red [31]. FISH has the advantage that we can visualize multiple targets using a number of hybridization probes at the same time in the same specimen. Gerion et al. [78] prepared four different colored (green, yellow, orange, and red) QDs tagged with four different ss-DNA sequences of 21 bases each. As seen in Figure 27.5 the different colored probes can be seen with the same excitation source and particular emission filter [78]. If the four probes are mixed together, these can be sorted by using a sequence specifically complementary to one of the probes. In another study, two different QDs were conjugated to the oligonucleotides targeted for Vmat2 mRNA in mouse midbrain. Colocalization of both QDs was seen due to in situ hybridization of the oligonucleotide to the mRNA sequence [107]. Such broad applications of FISH makes it a very useful technique in pathological labs (for detection of specific bacteria) and for medical diagnosis to check for particular chromosomal abnormalities. FISH requires different emission filters to visualize different colors. It is limited to hybridization and cannot be used to obtain information on the stability of delivery of vector–biomolecule complexes, which is required to determine the efficacy of the delivery system. To overcome these issues, Forster resonance energy transfer or fluorescence resonance energy transfer (FRET) has been used. FRET is a mechanism by which energy transfer occurs between two fluorophores that are less than 10 nm apart (Forster distance) in the presence of an external excitation laser source. For FRET to happen three factors are of utmost importance:

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(A) 2.0

PL [a.u.]

1.6 1.2 0.8 0.4

(B)

PL [a.u.]

(C)

1.0 0.8 0.6 0.4 0.2 0.0

520 600 680 Wavelength [nm]

0 480 520 560 600 640 680 720 Wavelength [nm]

520 600 680 Wavelength [nm]

520 600 680 Wavelength [nm]

520 600 680 Wavelength [nm]

Figure 27.5. (A) Fluorescence spectra of the solution (black) obtained by superposition of the spectra at different emission filters for green, yellow, orange, and red QDlabeled oligonucleotides when hybridized to their respective complementary sequence (mechanism is called FISH). (B) Squares indicate presence of QD tagged hybridization probes at different emission wavelength. (C) Individual spectra for different oligonucleotides, each with the solution spectra. (Reproduced with permission from Ref. 78.) (See color insert.)

• Distance between the fluorophores (should be less than 10 nm) • Spectral overlap of the fluorophores’ spectra • Quantum yield of the donor In usual cases, quantum dots are used as the donor due to their high quantum yield, broad absorption, and narrow emission spectra. The second fluorophore used for QD-based FRET must have its excitation specta overlapping the QD emission spectra so that the photon emitted by the QDs excite the second fluorophore, which can be seen in the microscope in its emission range. If these fluorophores are tagged on different participants of a complex, we can study the separation of the complex by FRET. For example, Wang’s group has linked QDs to plasmid DNA via a biotin–streptavidin linkage, and Cy5 dye to chitosan, which was complexed to DNA. FRET between the two

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occurs only when the complex was intact and only the red QDs were seen when the complex was lysed with heparin and chitosanase. By using this technique, it was possible to visualize a single molecule of chitosan–DNA complex after 4 h postincubation [49]. Recently, Lee et al. [51] performed similar studies using QD-DNA complexed with Texas Red tagged chitosan in HEK293 cells and found that better transfection occurs with chitosan of high molecular weight (∼500 kDa). FRET has also been useful for single QD-based detection of very low concentrations of DNA molecules (≤50 copies) using QD-DNA with a Cy5 labeled target gene. FRET between QD-Cy5 occurred when DNA was bound to the target sequence [55]. Interestingly, Zhou et al. [32] utilized a double-stranded DNA intercalating agent, ethidium bromide (EB), to study DNA hybridization. DNA tagged with QD, when hybridized to a complementary strand linked to EB, showed FRET. Energy transfer was also seen using an Alexa Fluor 594 linked complementary DNA strand [32]. Another recent publication used intercalating dye BOBO-3 for QD-based FRET [52]. Bakalova et al. [24] linked QD (donor) to siRNA and Cy5 (acceptor) to an mRNA sequence and used FRET as a means of testing the affinity of siRNA to the target mRNA. If the sequences are complementary, FRET could be seen, otherwise only the green QDs could be seen with the excitation wavelength. FRET is an effective tool for trafficking studies and can facilitate detection of complex dissociation, which in turn helps to explain the observed transfection efficiency. Besides these applications, FRET can also be used to study protein–protein [54] and protein–DNA interactions [133].

27.8

QD FOR IN VIVO IMAGING IN LIVE ANIMALS

Commendable progress on QD work in vitro has led to its exploration in in vivo systems. Near-IR dots have been imaged in live cells since these are less absorbed by the deep tissues when compared to UV or visible light. Special in vivo systems with CCD (charged couple device) cameras have been designed for this purpose. IVIS and Maestro systems are the most common ones. These instruments have excitation-emission filters that can be set as per the experimental conditions and the image is captured using the CCD camera [134]. Such systems are highly sensitive and allow noninvasive imaging of quantum dots. Cai and Chen [135] used both systems for imaging targeting peptide linked QDs in tumor cells expressing integrin αvβ3 receptors. Thiolated RGD (Arg-Gly-Asp) peptide was linked to amine QDs via heterobifunctional PEG and those were injected into mice. It was shown that the Maestro system was more sensitive due to reduced background signal as compared to the IVIS system [135].

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The work on in vivo QD imaging picked up after the remarkable work done by Dubertret et al. [12] on delivery of quantum dot–DNA conjugates to blastomers of Xenopus embryos with 2–5 × 109 QDs/cell. QDs were imaged only an hour after postincubation, using a fluorescence microscope. These were observed for 4 days since QDs were being transferred to the progeny. QD toxicity was negligible at the concentration used, whereas higher concentrations caused abnormalities [12]. Multicolored QDs can be visualized in vivo in the same animal, which makes it useful for trafficking of different molecules in the body simultaneously. Gao et al. [136] imaged three different colored QD-coated microbeads injected at three different locations in mice, with the same excitation laser. Kobayashi and co-workers [137] delivered five different colored QDs intracutaneously in different sites to study the lymphatic drainage system specifically in the neck and upper trunk region of live mice, using the Maestro in vivo imaging system 5 min after injection. In another study, Lin et al. [138] subcutaneously injected murine embryonic stem cells labeled with six different QDs 525, 565, 605, 655, 705, and 800 at different sites, in nude mice. All the colors could be visualized using a single excitation laser immediately after injection. Previous examples illustrate the imaging of QDs when delivered locally. A better utilization of the possibility of QD imaging in deep tissues would be to use targeted QD-based delivery systems. For this purpose, the Nie group delivered PEGylated QD-tagged antibody (prostate-specific membrane antigen—PSMA) that reached the target site when injected intravenously [136]. Carboxylated QDs did not reach the target site while some of the PEGylated QDs did make it due to PEG-mediated long circulation (passive targeting). QD-PEG-PSMA showed a strong signal in the tumor region, indicating effecting active targeting. Moreover, QD-COOH was cleared off much faster than QD-PEG due to the absence of PEGylation on the surface (Figure 27.6). Another useful application of QDs for in vivo imaging comes from its use with bioluminescent luciferase enzyme, a phenomenon called bioluminescence resonance energy transfer (BRET), which is similar to FRET, the major difference being that it doesn’t need an external excitation source and QDs act as acceptors rather than donors (for FRET). The concept of BRET was initially utilized to study protein–protein interactions [139] but later was developed for other applications. Quantum dots have also been utilized with luciferase to study the QD–biomolecule complexes in live animals. In these cases luciferase acts as a donor whose emission spectra overlaps with the excitation spectra of quantum dots. Photons emitted from the luciferase reaction excite the dots, which can be imaged using a suitable in vivo imaging system. For example, So et al. [57] conjugated QD with luciferase from Renilla reniformis and used those together with coelantrazine in a mouse. Coelenterazine in the presence of luciferase oxidizes and releases a photon that excites quantum dots. Accordingly, these conjugates when injected subcutaneously could be imaged without any excitation source in live mice

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QD-COOH

QD-PEG

QD-PEG-PSMA

Tumor

Tumor

Tumor

Injection site

Figure 27.6. Live mouse models injected with C4-2 human prostrate tumors treated with (A) 6 nmol QDs only (QD-COOH), (B) 6 nmol QD-PEG, and (C) 0.4 nmol QDPEG-PSMA. QD-COOH could not be seen, indicating that it was cleared off very fast. QD-PEG can be visualized weakly in the tumor region while a strong signal is seen at the tumor site on injecting QD with targeting ligand (QD-PEG-PSMA), indicating the efficiency of targeted delivery systems. (Reproduced with permission from Ref. 136.) (See color insert.)

[57]. When another fluorophore is used with this system that excites in the emission range of the quantum dots used, BRET can be utilized to see multiple targets in vivo. Another interesting experiment based on BRET was performed by Yao and co-workers for detection of the protease enzyme MMP2 (matrix metalloproteinases). This endopeptidase enzyme is found in high concentration in cancer cells. Yao linked MMP-2 substrate with six histidine tags to Renilla luciferase (Luc8). When the MMP2substrate-6His-Luc8 was mixed with QDCOOH in the presence of Ni2+ ions, a complex was formed between the metal ion, the carboxylic group (QD-COOH), and the MMP2 substrate-6His (QD/ Ni/MMP2-6His-Luc8), bringing QD and Luc8 in close proximity to allow BRET to occur. In the presence of the MMP2 proteolytic enzyme, BRET does not occur since this enzyme cleaves the MMP2 substrate, consequently dissociating the QD/Ni/MMP2-6His-Luc8 complex and thereby pushing Luc8 away from QDs [140]. Therefore without any QD modifications, we can visualize biomolecules in vivo without any external laser source by simply having them in close proximity to bioluminescent proteins such as luciferase.

27.9

CYTOTOXICITY OF QUANTUM DOTS

Considering that QDs have a heavy metal core shell, several questions have been raised regarding the safety of using QDs for biological purposes. Lovric et al. [141] have shown that unmodified CdTe core quantum dots are toxic

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due to surface oxidation when exposed to UV light/aqueous environment, which results in the released Cd2+ ions. Therefore a passive coating (usually ZnS) is required on the surface of the heavy metal core, to prevent its direct interaction with UV/oxygen. Although ZnS protects from air oxidation, it is ineffective in the presence of UV. Derfus et al. [60] showed that an additional BSA (bovine serum albumin) coating protects from UV-mediated oxidation for up to 8 h. A similar effect is provided by a DHLA (dihydrolipoic acid) and a polyacrylate coating. It should be noted that all these results are valid for QD concentrations below 1 mg/mL [60]. It has also been shown that along with the need for coating, the QD concentration also defines the potential toxicity. Earlier, Dubertret et al. [12] showed that 2–5 × 109 QD/cell were nontoxic and Lin et al. [138] showed that a 10-nM concentration of six different colored QDs was safe in embryonic stem cells for 72 h. Recently, it has been shown that an N-acetyl cysteine modification of QDs makes them less cytotoxic as indicated by their reduced membrane lipid peroxidation in human neuroblastoma cells [142]. Kirchner et al. [143] showed that polymer coating is better than mercaptopropionic acid. In addition, it has been shown that QD aggregation has a deleterious effect on their safety [143]. PEG coating has been shown to improve the biocompatibility of quantum dots [12]. CdSe/ZnS quantum dots coated with functionalized PEGylated lipids and conjugated with DNA have been shown to be nontoxic compared to DNA alone [11]. By appropriate surface modifications, quantum dots have been used extensively in in vitro conditions. Additionally, as discussed in previous sections, QDs have also been used in live animals and embryos [12].

27.10 CONCLUSION Quantum dots are being used as a valuable label for therapeutically active molecules such as DNA, RNA, peptides, membrane proteins, antibodies, pathogens, and even drugs. This is due to their stability against long-term laser exposure, high sensitivity, as well as their wide absorption and tunable emission spectra. QDs can be conjugated to biomolecules in several ways and this has helped in understanding the distribution of molecules inside the cells. The safety of QDs has enabled their use even in live animals. QDs have enormous potential to further aid in understanding the distribution of molecules at a cellular level as well as cellular processes.

27.11 FUTURE PERSPECTIVES So far, successful use of QDs in cells and animals has helped reveal important information on the localization and biodistribution of several biomolecules. QD properties such as photostability, extreme sensitivity, broad absorption spectra, and tunable emission spectra make these an outstanding labeling

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moiety when compared to organic dyes. Their further utilization to monitor molecular delivery especially in vivo may provide a clear understanding of cellular uptake and processing of biomolecules, including therapeutic agents, and therefore help in the efficiency and safety of therapeutic agents. Consequently, QDs have an enormous potential as markers for uncovering unknown facts at the molecular level and to help us learn the intricate behavior of biomolecules at the molecular and cellular levels.

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138. Lin, S., Xie, X., Patel, M. R., Yang, Y. H., Li, Z., Cao, F., Gheysens, O., Zhang, Y., Gambhir, S. S., Rao, J. H., and Wu, J. C. Quantum dot imaging for embryonic stem cells. BMC Biotechnol. 7: 67 (2007). 139. Subramanian, C., Xu, Y., Johnson, C. H., and von Arnim, A. G. In vivo detection of protein–protein interaction in plant cells using BRET. Methods Mol. Biol. 284: 271–286 (2004). 140. Yao, H., Zhang, Y., Xiao, F., Xia, Z., and Rao, J. Quantum dot/bioluminescence resonance energy transfer based highly sensitive detection of proteases. Angew. Chem. Int. Ed. Engl. 46(23): 4346–4349 (2007). 141. Lovric, J., Cho, S. J., Winnik, F. M., and Maysinger, D. Unmodified cadmium telluride quantum dots induce reactive oxygen species formation leading to multiple organelle damage and cell death. Chem. Biol. 12(11): 1227–1234 (2005). 142. Choi, A. O., Cho, S. J., Desbarats, J., Lovric, J., and Maysinger, D. Quantum dotinduced cell death involves Fas upregulation and lipid peroxidation in human neuroblastoma cells. J. Nanobiotechnol. 5: 1 (2007). 143. Kirchner, C., Liedl, T., Kudera, S., Pellegrino, T., Munoz Javier, A., Gaub, H. E., Stolzle, S., Fertig, N., and Parak, W. J. Cytotoxicity of colloidal CdSe and CdSe/ ZnS nanoparticles. Nano Lett. 5(2): 331–338 (2005).

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INDEX

1,1-bis-(2-acryloyloxyethoxy)-[4methoxy-phenyl]methane), 133 2,2-bis(N-maleimidoethyloxy)propane, 133 3,3-dithiopropionimidate dihydrochloride, 133 3T3-L1-cells, 59 4-(4′-dimethylaminophenylazo) benzoic acid (Dabcyl), 273 5-(N-ethyl-N-isopropyl)amiloride (EIPA), 59 A431 cells, 130 A459 lung epithelial cells, 236 Accessory protein-2, 366 Accumulation, subcellular, 4 Acid sphingomyelinase (ASM), recombinant, 460 Acid/base strength, 195 Acidic endosome, 64 Acidity constant, 387 Actin disruption, 452 -driven invagination, 479 filaments, 290 flow, 339 polymerization, 36 and active transport, 339 Acute ischemic stroke, gold nanoparticle-based assessment of, 100 Acute lung injury, 457 AD. See Alzheimer disease Adeno-associated virus, 209 single particle tracking of, 169 Adenocarcinoma MCF7 cells, 78 Adenovirus, 209

Adsorptive endocytosis, 196 Albumin-mediated polyplex clearance, 216 Aldehyde fixation, 184, 187 Alex fluor 568-phalloidin dye, 536 Alexa 488-R8 peptide, 274 Alexa 488-streptavidin, 276 Alexa Fluor 488, 186 Alginate chitosan polyplexes, 126 Allotopic expression, 394 ALS. See Amyotrophic lateral sclerosis Alzheimer disease (AD), 105 Amebas, 21 Amiloride, 107, 266, 326, 452, 455, 460 Amines, protonable, 129 Amine-terminated G4 PAMAM, 235 Ammonium telluride, 78 Amphipatic peptide-based nanocarriers, 325 Amphiphilic index, 387 Amphiphilicity index, 195 Amyotrophic lateral sclerosis (ALS), 105 Anionic lipoplexes, 544 Anisotropy, time-resolved, 39 Antennapedia protein, 276 Antibodies, QD-tagged, 555 Anti-CAM, 454 CAM nanocarriers, epitope controlled subcellular destination of, 462 catalase conjugates, 458 elliptical disks, 465 nanocarriers, 456 Antigen delivery by cell penetrating peptides, 405 Antigenomic peptide nucleic acids (PNAs), 492

Organelle-Specific Pharmaceutical Nanotechnology, Edited by Volkmar Weissig and Gerard G. M. D’Souza Copyright © 2010 John Wiley & Sons, Inc.

569

bindex.indd 569

7/30/2010 2:16:31 PM

570

INDEX

Anti-ICAM, 454 ASM/carriers, 460 nanoconjugates, 457 Antioxidants, 457 replacement enzymes, 457–459 Anti-PECAM, 454 catalase, 458 monoclonal, 465 SOD, 458 Anti-Stokes scattering, 517 Antiviral immune response, 375 Apoptosis, 6 and mitochondria, 385 Archaeal lipids, 390 Arenavirus, eGP defined receptor specifity and cell tropism of, 363 Arginine, role for CPP, 23 -rich peptides, 483 -rich protein transduction domains, 23 role for CPP, 23 Artificial chaperones, 104–109 Asialoglycoproteins, 482 ASM. See Acid sphingomyelinase, recombinant Astrocytes, 95 ATPase-H+ pumps, 248 Au nanostructures, TPL activities of, 516 Auger ionization, 541 AuNP. See Gold nanoparticles Autofluorescence, 95 Avidin-biotin affinity, 276 interactions, 495 Azomethine ylides, 147 Bacteriochlorin, 313 Bafilomycin A1, 129 Bioactivity, 2 Bioavailability, 2 Biodegradable esters, 133 Biodistribution, 2 Bioluminescence resonance energy transfer (BRET), 537, 555 BioPORTER®, 324 Block copolymers, micellization of QDs, 542 Blood-brain barrier, 105 Bodipy-LacCer, 58

bindex.indd 570

Bola-amphiphiles, 390 Bola-lipids, 390 Branched PEI, 215 BRET. See Bioluminescence resonance energy transfer Brownian motion, 168 Brownian particles, 39 Brust method, 83 Brust-Schiffrin one- and two- phase systems, 76 Bulk viscosity, 38 Bunyavirus, eGP defined receptor specifity and cell tropism of, 363 BV-2 microglial cells, 242 C60 fullerene derivatives, localization of, 200 Caco-2 cells, 44 and dendrimere, 239 Calcium channels, L-type, 442 CAM. See Cell adhesion molecule CAM-mediated endocytosis, 454–457 CAM-targeted carriers, 460 CAM-targeted drug delivery, 454 Cancer treatment, 85 Capillary endothelium, 438 Capping ligand, 74 Carbon nanotubes (CNT), 143–159 functionalization of, 145–147 intracellular trafficking of, 143–159 membrane receptor-mediated uptake of, 143–159 subcellular distribution of, 151–153 water-soluble, 151 Carboxyl-terminated G1.5 PAMAM dendrimers, permeability of, 239–240 Cardiac capillary barrier, 438 Cardiac myocytes, 433–447 delivery of cerium oxide nanoparticles to, 438 Cardiac Troponin I, 443 Cardiac vascular permeability, 437–439 Cardiolipin, 388 Carrier degradation, 218 Carrier geometry controlled intracellular trafficking, 463–466 Catalase, 110, 458 conjugated to anti-CAM, 458 immunoconjugates, 459

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INDEX

Cathepsin D, 183 Cationic dendrimere, 123 Cationic G2 PAMAM dendrimers, permeability of, 239–240 Cationic lipids, 480 in virosomes, 372 Cationic polymer coated QDs, 546 Cationic polymers, 480 Caveolae, 56, 265, 479 -dependent endocytosis, 40 -dependent pinocytosis, 537 -mediated endocytosis, 98, 124, 366– 369, 439, 451–453, 479 -mediated process, 198 Caveolin proteins, 537 Caveolin-mediated endocytosis, 366-369 Caveolins, 479 Caveosomes, 126, 479 CBN. See Conjugated bond number CCI. See Clathrin, and caveolinindependent endocytosis CCP-induced lactate dehydrogenase leakage, 330 CCVs. See Clathrin, coated vesicles C-dots, superbright, 18 CdSe quantum dots passively coated with ZnS, 542 CdTe quantum dots, mercaptoacetic acid coated, 544 Cell(s) apical pole of, 59 basolateral pole of, 59 dividing, 42 non-dividing, 42 nonmitotic, 48 surface markers, 28 surface ruffling, 479 viability assays, 27 Cell adhesion molecule (CAM), 5 cytosolic domain, tyrosine phosphorylation of, 455 endocytosis, 454–457 epitopes, carrier binding of, 461–463 targeted carriers, 460 targeted drug delivery, 454 Cell entry mechanism, 5 Cell fixing, artifacts, 270 Cell-penetrating peptides (CPP), 22, 263–268, 324, 403–432 antigen delivery, 405

bindex.indd 571

571

attachment to proteins, 274–278 carriers for cargo, 404–410 carriers for nucleic acids, 407–410 chimeras with targeting ligands, 410 combination with drug delivery systems, 404–432 definition of, 267 derivatization with acyl chains, 419 disulfide-linked conjugates of, 409 endocytosis of, 414 endosomal escape of, 414–417 fluorophores, 278–279, 405 fusion proteins, 278–279 intracellular trafficking of, 417–418 liposomes, 410–412 magnetic nanoparticles, 410–412 mechanism of cell internalization, 412–418 -mediated toxicity, 277 mimetics, 275–276 particulate drug delivery systems, 410–412 peptide-based immunotherapy, 407 peptide and protein delivery, 406 peptide-protein chelates, 278 peptides and proteins, 406 polymeric micelles, 410–412 protein complexes, 277–278 quantum dots, 410–412 side effects of, 421–422 siRNA, 408 toxicity of, 421–422 transduction of, 417 Cellular distribution, assessment of by quantum dot labeling, 535–567 Cellular entry, barriers to, 537–539 Central nervous system, 95 Ceramide Gb3 receptor, 64 incorporated in STPP liposomes, 395 Cerebral ischemia, 110 Cerium oxide nanoparticles, cardioprotective effect of, 443 Cetyltrimethylammonium bromide (CTAB), 508 -coated GNRs, 510 -depleted GNRs, 514 toxicity of, 512

7/30/2010 2:16:31 PM

572

INDEX

Chaperones, 186 artificial, 104–109 polymeric artificial, 93–121 Charge density oscillations, 99 Charge-reducing polycations, 220 Chariot™, 327 Chemical libraries, 2 Chemisorptive coatings, 511–512 CHEMS. See Cholesteryl hemisuccinate Chitosan, 123 -based carriers, packaging efficiency of, 215 Chlorin, 313 Chloroquine, 129 Chlorpromazine, 59, 266, 481 CHO cells, proteoglycan-deficient mutant, 124 CHO-K1 cells, 106 Cholera toxin B, 5 Cholesterol -bearing pullulans (CHPs), 105 -dependent CME, 481 depletion experiments, 126 -GALA peptide, 489 Cholesteryl hemisuccinate (CHEMS), 490 Chondroitin sulfate, 124 Chondroitinase ABC, 345 CHP nanogels, 106 cationic derivatives of, 106 HER2 tumor antigen incorporation, 108 CHP-HER2 CD4+ and CD8+ T-cell response, 108 CHPNH2. See CHP nanogels, cationic derivatives of CHPNH2-FITC-Bovine serum albumin, 106 CHP-quantum dots, 106 CHPs. See Cholesterol, -bearing pullulans Chromosomal DNA, role in unpacking, 217 Ciliates, 21 Cis-acting nuclear DNA import, 135 Cis-Golgi, 185 Clathrin, 56 and caveolae-independent (CCI) endocytosis, 124, 440, 451–453

bindex.indd 572

-coated pits, 57, 149, 366 -coated structures, 264 -coated vesicles (CCV), 265, 478 -dependent pinocytosis, 537 -dependent endocytosis, 148 -independent endocytosis, 148, 440 -mediated endocytosis, 98, 124, 167, 366–369, 439, 451–453, 477–478 -mediated endocytosis, inhibition of, 107 Click bioconjugation, 509 CMC. See Critical micelle concentration CME. See Endocytosis, clathrin-mediated CNT. See Carbon nanotubes Coated pits, 478 Coelenterazine, 555 Collagenase degradable linker, 546 Collisional interaction, 39 Colloid chemistry, 194 Colloidal gold, treatment of rheumatoid arthritis, 100 Compartments, subcellular, 4 Confocal reflectance microscopy, 515 Congestive cardiomyopathy, 441 Conjugated bond number (CBN), 195, 387 Convection currents, 35. See also Cytoplasmic diffusion Convectional drift, 165 Copper acetate, 77 Core-shell particles, 18 Coronavirus, eGP defined receptor specifity and cell tropism of, 363 Cos-7 cells, 106 CPP fluorophores, 278–279 CPP fusion proteins, 278–279 CPP-mediated toxicity, 277 CPPs. See Cell-penetrating peptides Cremophor EL, 392 Critical micelle concentration (CMC), 251, 371 Crosslinker degradation, 221 CTAB. See cetyltrimethylammonium bromide Cutinase suicide inhibitor, 546 Cutinase, 546 Cyclodextrin, 452 Cylindrical polymer particles, 464

7/30/2010 2:16:32 PM

INDEX

Cysteine capping of quantum dots, 61 Cytochalasin D, 25, 59, 266, 326, 343, 452, 455 Cytoplasm, 4 Cytoplasmic diffusion, 35 coefficients, 290 Cytoplasmic dynein, 292 Cytoplasmic meshwork, 49 Cytoplasmic radius, 41 Cytoplasmic viscosity, 386 Cytoprotection, 111 Cytoskeletal filaments, 292 Cytoskeletal motors, 292–293 Cytosolic delivery of biomacromolecules, 403–432 Cytosolic mobility of gene delivery vehicles, 213 Cytostatica, specific targeting of, 64 Cytotoxic T lymphocytes, 405 Dabcyl, 273 Darkfield microscopy, 515 Delivery technique, mechanical, 19 technique, membranal, 19 intracellular, 19 liposomal, 22 nanosensor, 19 systems using virus envelope glycoproteins, 369 Delocalized lipophilic cations, 388 Dendrimer(s), 35–54 actin interaction, 44 adhesion, 42 cationic, 123 cellular entry dynamics, 233 cellular trafficking of, 231–246 DNA complex, see Dendriplexes generation number, 231 PCI-mediated drug delivery, 317 photosensitizers, 317 polydispersity indices of, 231 polylysine sixth generation, 37 rate of uptake, 236 self-fluorescent, 36 surface-engineered, 40 therapeutics, 233 Dendrimer structures carbohydrates, 231

bindex.indd 573

573

nucleic acids, 231 poly(aryl ethers), 231 polyamidoamines, 231 polyamines, 231 polyesters, 231 Dendriplexes, 35–54 hydrodynamic diameter, 47 zeta potential, 36, 47 Dendritic architecture, 231–233 Dendritic cells, 405 Dendritic poly(L-lysine), 217 Dequalinium chloride, 6 Dequalinium derivatives, 201 Dextran, 18 DHPE, 254 Dichotomous decision rules, 195 Didodecyldimethylammonium bromide, 77 Diethylaminomodified dextran, 123 Diffusion coefficient, 38, 41 hindered, 4, 386 intracellular, 4, 386 net solute translational, 38 Dihydrolipoic acid capping of quantum dots, 61 Dihydrolipoic acid, disulfide linkage with ZnS coating, 542 Dimethyl amiloride hydrochloride, 349 Diphtheria toxin, 64 Distribution, intracellular, 4 DNA carrying nanodelivery systems, intracellular trafficking of, 475–506 complexation, 209 delivery to mitochondria via DQAsomes, 6, 390–394 polyplex dissociation, 132–134 tagged with QDs, 537 transport, barriers to, scheme, 211 Dodecanethiol, 77 DOPE/CHEMS, 491 DOTAP, 58 DOX release from VM nanogels, 256 DOX-induced multidrug-resistant ovarian carcinoma cell lines, 254 Doxorubicin release from mixed micelles, 251

7/30/2010 2:16:32 PM

574

INDEX

Doxycycline, 442 DQAplexes, 492. See also DQAsome/ DNA complexes DQAsome/DNA complexes, 391–392 DQAsomes, 6, 201, 390–394, 492 conjugated with folic acid, 393 Monte Carlo computer simulations of, 391 Drosophila Antennapedia homeoprotein, 22 Drug carriers, intracellular delivery of, 449–451 Drug delivery to mitochondria via DQAsomes, 390–394 nanoparticle-based, 85 role of carrier geometry, 463–466 Drug disposition, 3, 4 Drug release, photothermally activated, 523 Drug targeting, 3 Drug therapy, 1 Drugs, mitochondriotropic, 6 Drugs, tagged with QDs, 537 Dual-color total internal reflection fluorescence (TIRF) microscopy, 63 Dynamin, 452 Dynein binding peptides, 338 dominant negative mutant, 59 mediated transport, 171 motors, 292, 338 single motor parameters of, 297 Early endosomal antigen 1 (EEA1), 181, 368 immunolabeling of, 182 Early endosomal marker, 64 Early endosome(s), 181–182, 248, 368, 478 subpopulations of, 367 Ebola virus, EM of, 358 EEA1. See Early endosomal antigen 1 EGF receptor. See Epidermal growth factor receptor EGF-PEG-PEI, cell binding, cell entry and intracellular trafficking, scheme of, 128 EGF-QDs, 63

bindex.indd 574

eGP. See Envelope glycoproteins eGP-coated nanoparticles, 370 eGP-mediated delivery, 361–362 Ehrlich, Paul, 1 EIPA, 59 Electric charge Z, 195 Electroporation, 20 Ellipsoidal dendrimers, 232 Elliptical disks, 464 Endocytic markers, 277 Endocytic organelles, markers of, 180–186 Endocytic pathways, 177–192, 264–266, 537–539 inhibitors of, 266 Endocytosis, 5, 148, 232, 248, 264–266, 365–369 adsorptive, 196 Arf6-mediated, 56 CAM-mediated, 450 caveolin-assisted, 5 caveolin-dependent, 40 caveolin-mediated (CME), 98 Cdc-42-mediated, 56 cell adhesion molecule-mediated, 450 clathrin-dependent, 5, 40, 64 clathrin-independent, 56, 57 clathrin-mediated, 56, 98 lipid raft associated, 5 nonselective, 480 peptide-mediated, 65 Rho-A-mediated, 56 Endocytotic pathways, scheme of, 478 Endolysosomal network, escape from, 272–273 Endolysosomal system, 5 Endolysosomolytic materials, 248 Endolysosomolytically active nanotechnology, 247–262 Endoplasmic reticulum, 186 Endoplasmatic reticulum to Golgi intermediate compartment (ERGIC), 185 Endosomal acidification, 365 Endosomal compartments, destabilization of, 248–250 Endosomal disruptors, 415 Endosomal escape, 213, 488–491, 538

7/30/2010 2:16:32 PM

INDEX

Endosomal release, photoinduced, 130–131 Endosomal rupture, 248 Endosome(s), 5 acidic, 64 lysosome fusion, 25 movement of, 294 transport of along microtubule, 128 Endosomolytic polymeric gene nanocomplexes, 258–259 Endothelial CAM-mediated endocytosis, 451 Endothelial cells, 56, 64, 450 Endothelial junction, inflammation of, 438 Endothelial targeting of drugs, 454 Endothelial transmembrane glycoproteins, 454–457 Endothelium capillary, 438 venular, 438 Enhanced permeability and retention (EPR) effect, 3, 212 Envelope glycoproteins (eGP), 358–360 conformation changes of, 364–365 delivery systems, 369 as environmental sensors, 364–365 targeting to endocytic pathways, 365–366 Enveloped viruses, 358–359 Environment, intracellular, 15 Ephrin receptors EphA3, 67 Epidermal growth factor, 63 Epidermal growth factor (EGF) receptor, 368, 511 biotinylated, 63 Epilepsy, 110 Epithelial cells, 56 Epitope controlled subcellular destination of anti-CAM nanocarriers, 462 EPR effect. See Enhanced permeability and retention effect ER tracker, 536 ERGIC. See Endoplasmatic reticulum to Golgi intermediate compartment Escort, 22 Ethidium bromide, 82 Ethidium thiol, structure of, 81

bindex.indd 575

575

Excitation energy, dissipation of, 85 Ezrin, 456 Ezrin/radixin/moesin protein family, 456 f-CNT. See Functionalized carbon nanotubes FCS. See Fluorescence correlation spectroscopy FG motif. See Phenylalanine glycine motif Fibroblasts, 56 Fick’s law of diffusion, 195 Fick–Nernst–Planck model, 388 Filipin, 58, 148, 452 Filopodia, 337 Filovirus, eGP defined receptor specifity and cell tropism of, 359, 363 FISH. See Fluorescence in situ hybridization FITC. See Fluorescein isothiocyanate FITC-dextran, 536 Fixation of cells, 186–189 Flavivirus, eGP defined receptor specifity and cell tropism of, 363 Flotillin, 57 Flotillin-1-dependent pathway, 265 Flow cytometry, 27 Fluid phase endocytic capture, 184 endocytic pathways, 349 endocytosis, 196 uptake, 265 viscosity, 4, 38, 386 Fluorescein diacetate, 27 Fluorescein isothiocyanate (FITC), 106 Fluorescence correlation spectroscopy (FCS), 162 Fluorescence energy transfer (FRET), 273, 546 Fluorescence in situ hybridization (FISH), 134, 544 and QDs, 552–554 Fluorescence recovery after photobleaching (FRAP), 44, 161 Fluorescence resonance energy transfer, 537 and QDs, 552–554 Fluorescent dyes, 15

7/30/2010 2:16:32 PM

576

INDEX

Fluorescent gold nanoparticles, 79 Fluorescent polystyrene beads, intracellular trafficking of, 167–168 Fluorophores, 4, 18 FM 4–64 FX dye, 536 f-MWNTs. See Functionalized multiwalled nanotubes Folate conjugated gold nanorods, 519 conjugated InP-ZnS quantum nanocrystals, 547 folate receptor-mediated pathway, 149 receptors, over expression of, 85 tagged with QDs, 537 Folic acid, 5 Folic acid modified DQAsomes, 393 Formaldehyde, 187 Formulation charge ratio of lipoplexes, 209 Forster distance, 552 Four-wave mixing, 517 FRAP. See Fluorescence recovery after photobleaching FRET. See Fluorescence energy transfer FRET based quenching of QDs, 546 Frictional force, molecular motors, 293 f-SWNTs. See Functionalized singlewalled nanotubes Fugene, 22 Functionalized carbon nanotubes (f-CNT), 146 Functionalized multiwalled nanotubes, 146 Functionalized single-walled nanotubes (f-SWNTs), 146 Fusogenic lipids, 415, 488 Fusogenic peptides, 415, 488–489 G3 PAMAM dendrimers, cellular uptake mechanisms, 236 dendrimers, intracellular trafficking of, 238 PEG modification of, 235 GAGs. See Glycosaminoglycans GALA, 131, 267, 438 Gap junctions, 196 Gene carriers, single particle tracking of, 169–171

bindex.indd 576

Gene delivery, 123 Gene guns, 856 bombardment, 19 Gene packing, methods of, 208–210 Gene therapy, 475 mitochondrial, 394 Genistein, 58, 126 GFAP. See Glial fibrillary acidic protein GFP. See Green fluorescence protein GFP protein, 7 Glial cells, 95 Glial fibrillary acidic protein (GFAP), 96 Glutathione glutaredoxin, 111 peroxidase, 110 S-transferase tags, 110, 270 Glycine receptors, lateral dynamic of, 63 Glycosaminoglycans (GAGs), 124, 209 GNR-based SERS, 519 GNR-mediated photothermolysis, 521 GNR-PNIPPAAm hydrogels, 523 GNRs. See Gold nanorods Gold colloids, 73 nanocrystals, 83 nanodisks, 100 Gold nanoparticles, 75, 93–121 alkanethiolate-coated, 80 assessment of hepatic function, 100 biocompatibility of, 83, 100–103 biodistribution of, 100–103 blood-brain barrier, 102 cellular internalization of, 102 interaction with living cells, 100–103 PEG-stabilized, 85 phosphonium-capped, 82 phosphonium-functionalized, 83 radioactive, treatment of liver cancer, 100 ricin binding, 67 rod-shaped, 99 size and toxicity, 101 tiopronin-coated, 80 toxicity toward red blood cells, 100 tris(hydroxymethyl)phosphine(THP)capped, 83 Gold nanoprisms, 100 Gold nanorods (GNRs), 100, 507–533 anisotropic functionalization of, 511

7/30/2010 2:16:32 PM

INDEX

antibody-functionalized, 511 antibody-labeled, 515 -based SERS, 519 biocompatibility of, 512–514 biosensing applications of, 511 cellular imaging, 514–520 cellular uptake of, 512–514 coating, 510–512 core-shell formation of, 512 CTAB-stabilized, 510 detoxification of, 514 folate-conjugated, 519 -mediated photothermolysis, 521 membrane-bound, 522 multiplex SERRS imaging with, 520 NIR-absorbing, 508 nonlinear optical properties, 516–519 PEG-conjugated, 518 photothermal agents, 520–522 polyelectrolyte-coated, 522 -PNIPPAAm hydrogels, 523 silica-coated, 512 subcellular localization of, 522–523 surface chemistry of, 508–510 TPL contrast agents, 518 TPL intensity of, 516 Gold nanoshells, 99 Gold nanostars, 100 Gold nanourchins, 99, 100 Gold nanowires, 100 Gold(III) chloride, reduction of, 75 Gold-labeled DNA, 481 Golgi apparatus, 185, 478 Golgi ribbon, 185 Golgi-EGFP, 536 gp-120, 268 Green fluorescence protein (GFP), mitochondrial delivery of, 493 Groove binding, 82 GTPase dynamin, 56 GTPase Rab5, 181, 367 GTPases, rho family, 367 Heat-shock protein, 186 HEK293T cells, 126 HeLa cells, 106 digitonin permeabilized, 135 Hemagglutinin peptide HA2, 131 Heme oxygenase 1, 110

bindex.indd 577

577

Hemolysis, for evaluation of endolysosomolysis, 251 Hemophilia, gene therapy of, 208 Henderson-Hasselbach equation, 195 HEp-2 cells, 64 Hepadnavirus, eGP defined receptor specifity and cell tropism of, 363 Heparin sulfate, 124 Heparan sulfate proteoglycan, 210, 345, 480 Heparinases, 345 Hepatic function, gold nanoparticlebased assessment of, 100 HepG2 cells, 130 Herpes simplex virus, 209 Herpes simplex virus VP-22 protein, 22 Herpesvirus, eGP defined receptor specifity and cell tropism of, 363 HGH6 peptide, 330 HGH6/caspase-3 complex, 330 Hierarchical oxidative stress model, 110 HIV-Tat, 22, 263, 267–268, 272 protein, cellular dynamics of, 268–269 protein, nuclear entry of, 273 HIV-Tat-enhanced green fluorescent protein fusion protein, 269 hMSCs. See Stem cells, human embryonic mesenchymal Hoechst 33342, 42, 536 HUH 7 cells, 127, 130 Human adenocarcinoma cells, 127 Human embryonic kidney HEK293 cells, 258 Human hepatoma HepG2 cells, 258 Hyaluronic acid, 124 Hydrodynamic injection, 247 Hydrogen peroxide, 457 Hydrophobically modified glycol Chitosan polyplexes, 126 Hydroxyl-terminated G4 PAMAM, 235 Hydroxyl-terminated G2 PAMAM dendrimers, permeability of, 239–240 Hyper-Rayleigh scattering, 516 ICAM-1 antibodies, 5, 454 ICAM-1. See Intercellular adhesion molecule-1

7/30/2010 2:16:32 PM

578

INDEX

ICP-MS. See Inductively coupled plasma mass spectrometry ICR mice, 87 IgG-opsonized particles, 465 IgorPro software, 41 Ikarugamycin, 107 IL-12, 108, Imaging technology, 8 Immunoadhesins, 364 Immunofluorescent microscopy, 179 Immunotoxin, intracellular delivery of, 65 Importins, 135 In vivo imaging with multicolored QDs, 555 India ink, 198 Inducible nitric oxidase synthase (iNOS), 109 Inductively coupled plasma mass spectrometry (ICP-MS), 101 Inflammation caused by metallic nanoparticles, 109 suppression of, 454 Inflammatory mediators, 439 Influenza A hemagluttinin, 272 Influenza X-13 virus, single particle tracking of, 170 iNOS. See Inducible nitric oxidase synthase InP-ZnS quantum nanocrystals, folate conjugated, 547 Integrins, 124, 480 binding domain, 5 Intercalation, 82 Intercellular adhesion molecule-1 (ICAM-1), 453 antibodies, 5, 454 Interendothelial junctions, 438 Interleukin-2 receptors, 538 Intermitochondrial contacts, 441 Interstitial extracellular matrix, 212 Intracellular barriers, 386–390 delivery of biologically active proteins, 323–336 distribution, QSAR-based prediction of, 387 drug delivery, 449–474

bindex.indd 578

localization of nanoparticles, prediction of, 193 polymer localization, 177–192 trafficking of DNA, uptake pathways dependent, 475–506 trafficking of viruses, 290–291 trafficking, assessment of by quantum dot labeling, 535–567 trafficking, control by carrier geometry, 463–466 transport, regulation of, 295 Intranuclear microinjection, 134 Intranuclear transfer, 213 Ionophores, optically silent, 16 IR light, 85 IRQ-Lip, 487 Ischemia reperfusion injury, 457 Isopolar microtubule cytoskeleton, 294 Jembrane disease virus, 272 jetPEI, 217 Jurkat T lymphocytes, 270 K562 cells, 127 K8. See octalysine K8-MEND, 490 KALA, 131, 267 Karyoplasm, 135 Kinase, p21-activated, 58 Kinesin -mediated transport, 171 motors, 292, 338 single motor parameters of, 297 Kinesin-1, 292 LabelIT® Tracker™ Fluorescein kit, 41 LacCer. See Lactocyl-ceramide Lactate dehydrogenase leakage, CCPinduced, 330 Lactocyl-ceramide (LacCer), 58 LAMPs, 183 LAMP1. See Lysosome-associated membrane protein Largest conjugated fragment, 195, 387 Laser ablation, 78 Late endosomes, 182–183, 213, 248, 368, 478 Latex beads, 58 Latrunculin A, 455

7/30/2010 2:16:32 PM

INDEX

Layered double hydroxide (LDH) nanoparticles, 7 LCG. See Largest conjugated fragment LDH nanoparticles. See Layered double hydroxide nanoparticles LDL receptors, 477 Leader peptide, mitochondrial, 5 Leader sequence, 4 Lentivirus, 209 Lesion development, gold nanoparticlebased assessment of, 100 Leupeptin, 183, 188 Libraries, chemical, 2 Ligand exchange, schematic representation of, 79 mitochondriotropic, 5 mediated internalization, 482–483 nanoparticle bioconjugates, 67 subcellular targeting, 4 Linear PEI, 215 Linear photoluminescence, 515–516 Linkers cleavable, 3 photocleavable, 26 Lipfectamin, 214 Lipid rafts, 124, 148, 271, 367, 452 Lipid-raft-dependent pathway, 59 Lipofectamine 2000, 22 comparison to dendrimers, 241 comparison to R8-MEND, 486 Lipofectin, comparison to R8-MEND, 486 Lipoplex(es), 209, 480–482 anionic, 544u uptake, 481, 538 Lipopoly-L-lysine lipoplexes, 481 Liposomal delivery, 21 Liposome(s) delivery by CPPs, 410–412 double-labeled, 484 long-circulating, 3 mitochondriotropic, 394–395 pH-responsive, Tat-modified, 412 QD loaded, 547 R8-modified, 483 silica-loaded, 547 targeted to selectins, 451 Load force, molecular motors, 293

bindex.indd 579

579

Localized surface plasmons, 99 logP. See Octanol-water partition coefficient Longitudinal plasmon resonance, 508 Long-term same-cell imaging, 536 Long-term trafficking studies, 536 Low-density lipoprotein, 5, 477 Luciferase -expressing mice, 96–97 QD-conjugated, 555 transfection, 58 Lymphocyte function-associated antigen, quantum dot labeled, 546 Lymphocytic choreomeningitis virus, 369 Lysosensor green, 536 Lysosomal enzyme replacement therapy, 459–460 Lysosomal markers, 64 Lysosomal storage disease, 278 Lysosome-associated membrane protein (LAMP) 1, 456 Lysosomes, 5, 21, 183–185, 478 targeting of, 278 LysoTracker®, 23, 184. 254 M918 peptide, 329 Macromolecular crowding, 38 Macrophages, 56 Macropinocytosis, 7, 57, 98. 124, 199, 265, 348–349, 368, 439, 451–453, 479–480 Macropinosomes, 58, 479 Magic Bullet, 1, 2, 3 Magnetic nanoparticles, delivery by CPPs, 410–412 Major histocompatibility complex classes I and II, 405 MALDI-MS, 84 Mannose-6-phosphate receptor(s), 5, 182, 278, 460 Marina Blue-labeled WGA, 186 Masked thiolate, 83 Mastoparan, 267 mCherry fluorescent protein, 273 Mean squared displacement, 163 Medial-Golgi. See Golgi ribbon Melanosomes, traffic of, 295 Melittin, 131, 267

7/30/2010 2:16:32 PM

580

INDEX

Membrane -active peptides, table of, 268 curvature, 200 disruptive peptides, 131–132 disruptive polymers, 416, 489 fusion, 199 invagination, 56 perinuclear, 50 permeation, QSAR-based prediction of, 387 ruffling, 57, 349 MEND. See Multifunctional envelopetype nanodevice Mercaptoacetic acid, disulfide linkage with ZnS coating, 542 Mercaptobenzoic acid, 75 Meso-tetraphenylporphine, disulfonated, 130 Metal nanoparticle biological response to, 109–111 DNA assemblies, 80 photoluminescence of, 515 synthesis of 74 therapeutic agents, 85 toxicology of, 86 Methotrexate (MTX), 242 -functionalized magnetic Fe3O4 nanoparticles, 85 interaction with folate receptor, 85 -resistant Chinese hamster ovary cells, 242 -resistant human acute lymphoblastoid leukemia cells, 242 -sensitive Chinese hamster ovary cells, 242 -sensitive human acute lymphoblastoid leukemia cells, 242 Methylene blue, 27 Methylene glycol, 187 Methyltriphenylphosphonium cations 394 Methyl-β-cyclodextrin, 61, 266 Micelles, 8 delivery by CPPs, 410–412 Microdomains, 63 Microglia, 95 Microtubule -dependent active transport, 168, 213 filaments, 290 minus end, 293

bindex.indd 580

organization of, 294 organizing center, 290 plus end, 293 transport system, 128 Microvilli, 337 apical, 349 Middle Ages, 73 Mie theory, 508 Mitchondria-junctional SR contacts, 441 Mitochondria mammalian, 5 movement of, 294 targeting of, 83 Mitochondrial accumulation complex formation based, 388 electric potential based, 388 ion-trapping based, 388 of xenobiotics, physicochemical features, 389 Mitochondrial delivery of green fluorescence protein, 493 via membrane fusion, 395–396 MITO-Porter, 494 Mitochondrial diseases, 491 DNA, 385 DNA delivery, 6 DNA metabolism, 390 DNA mutations, 491 gene therapy, 6, 394 intermembrane space, 441 leader peptide, 5, 391 localization, 201 network, 385 targeting, 491–494 targeting signal (MTS) peptide, 492 Mitochondria-targeted drug and DNA delivery, DQAsomes, 390–394 nanocarriers, 385–401 polyethylene imine, 394 therapies, 385 Mitochondriotropic compounds, 388 drugs, 6 liposomes, 394–395 MITO-Porter, 7, 395–396, 492–493 mitochondrial delivery, 492 Mitotracker, 536

7/30/2010 2:16:32 PM

INDEX

Mixed pH-sensitive micelles containing targeting folate (PHSM/f), 253 MLP. See Mitochondrial leader peptide Moesin, 456 Molecular crowding, 386 Molecular motors, 291–293 antagonistic motors, 300 building blocks for nanotechnological applications, 306 cargo transport, 302–303 frictional forces, 299 hydrodynamic friction, 299 intracellular drug delivery, 289–309 motility states, 301 patterns of movements, 301–302 receptor proteins for, 305 spatial displacement of cargo particles, 302 Tug-of-War between, 300–301 Monensin, 455, 461 Monocytes, 56 Monolayer protected clusters (MPC), 74 Monte Carlo computer simulations of DQAsomes, 391 Monte Carlo simulations, 166 Motor protein-guided active transport, 170 Motor proteins, 338 Motor velocity, 297 MPC. See Monolayer protected clusters MSD. See Mean squared displacement MSNs. See Nanoparticles, mesoporous silica mtDNA replication, selective inhibition of, 492 mtDNA, 491 MTOC. See Microtubule-organizing center MTS. See mitochondrial targeting signal peptide MTS-conjugated PNA, 492 MTX. See Methotrexate MTX-resistant Chinese hamster ovary cells, 242 MTX-resistant human acute lymphoblastoid leukemia cells, 242 MTX-sensitive Chinese hamster ovary cells, 242 MTX-sensitive human acute lymphoblastoid leukemia cells, 242

bindex.indd 581

581

Multicolored QDs, in vivo imaging. 555 Multifunctional envelope-type nanodevice (MEND), 476, 486 IRQ-modified, 487 tetralamellar, 488 Murin cadherin, 276 Murine leukemia virus, eGP of, 367 Murine leukemia virus, EM of, 358 Myosin motors, 292, 338 Myosin V, 295 myotubes, 440 N-(2-mercaptophoropionyl)glycine, 82 N-(fluorescein-5-thiocarbamoyl)-1,2dihexadecanoyl-sn-glycero-3phosphoethanolamine (DHPE), 254 N9 cells, 107 N-acetyl cysteine, 457 NADPH quinine oxyreductase, 110 NaN3, 148 Nanocantilevers, 437 Nanocapacitors, 437 Nanocarriers amphipatic peptide-based, 325 anti-CAM conjugated, 456 cell-penetrating peptide-based, 329–330 distribution, 4 mitochondria-targeted, 6 peptide-based, 323–336 pharmaceuticals, 3 targeted, 4 Nanodelivery systems, subcellular fate of, 93–121 Nanodevices, 437 Nanoepigenetics, 97 Nanogels, 104–109 artificial chaperoning properties of, 105–106 blood-brain barrier, 105 drug delivery system, 107 interaction with Aβ, 107 intracellular delivery and fate, 106–107 neutral, 107 protein aggregation prevention, 105 protein refolding by, 105 tetramethyl rhodamine isothiocyanatelabeled, 107 Nanomedicine, cytosolic delivery of, 311–322

7/30/2010 2:16:32 PM

582

INDEX

Nanomedicine, subcellular, 1–13 Nano-object(s) barriers for their delivery, 436 delivery to cardiac myocytes, 433–447 diffusion pathways inside myocytes, 434 internalization by transmembrane pathways, 439–440 Nanoparticle(s), 3 anionic polymeric, 59 based drug delivery, 85 cationic, 59 cellular redox state, 109 charged polystyrene, 59 distribution in brain, 95 DNA assemblies, 81 DNA interactions, 80 electrochemical synthesis of, 78 endocytosis of, 55–72 fibrillation of β-amyloid, 106 functionalized for biological detection, 84 induced injury, 109 intracellular active, directed transport, 128 K8-modified, 490 long-term cumulative exposure to, 93 mesoporous silica, 59 noble metal, 78 QSAR models, 193–206 R8-modified, 486 size dependence of uptake, 103, 126 trafficking of, 55–72 ultrabright, nonbleaching, 99 uptake by cells, mechanism of, 98 virus-envelope coated, 357–383 water-soluble cadmium telluride, 78 Nanopores, 437 Nanopores, transient, 47 Nanorods, 7 Nanoscale particles, active cellular transport of, 337–356 Nanosensor(s), 15–33 Calcium-sensitive, 24 matrix, 18 PEBBLE, 16 schematic figure, 17. See also color insert

bindex.indd 582

TAT-conjugated, 24, 24 technology, optical, 15 Nanotherapeutics, intracellular transport of, 161–175 Nanotoxicology, 97 Nanotrucks, cellular, 290 Naphthalocyanine, 313 NCE pathway, 368 Near-infrared (NIR) wavelengths, 507 Near-infrared laser irradiation, 100 Near-infrared QDs, in vivo imaging, 554–556 Nernst-Planck equation, 195 Neurotransmission, 110 Neurotransmitters, tagged with QDs, 537 Neutral red, 27 Nicotinic acid, 5 Niemann-Pick disease, 460 NIH-3T3 cells, 106 NIR. See near-infrared wavelengths NIR-emitting quantum dots, 95 Nitric oxide, 110 NLS. See Nuclear localization sequence NO. See Nitric oxide Nocodazole, 171, 455, 460 Non endocytic pathways, 537–539 Non-clathrin-noncaveolar endocytosis, 366 Non-Newtonian fluid, 38 Non-specific macropinocytosis, 212 Nonspecific particle uptake, 198 Nonspherical carriers, 463 Non-viral gene delivery vehicles, 209–210 Nonviral gene vectors, trajectory of, 164 Nonviral vectors, 475 NP. See nanoparticles NPC. See nuclear pore complex Nuclear localization sequence (NLS), 5, 135, 273, 494 Nuclear membrane barrier, 495 Nuclear morphology assay, 393 Nuclear pore complex, 47, 134, 441 Nuclear transfer of DNA, 494–497 Nucleic acid delivery by cell penetrating peptides, 408 Nucleoplasm, 4 Nucleus, particle uptake, 47 Nystatin, 326

7/30/2010 2:16:32 PM

INDEX

Octaarginine (R8), 7, 274 nitriliotriiacetic derivatized, 278 role in endosomal escape, 489–490 Octalysine, 490 Octanol-water partition coefficient, 195 Oligomeric propylacrylic acid, 259 Oligonucleotide-CPP complexes, 409 Optical traps, 296 Optochemical sensors, 16 Optodes, 16 Optoporation of tumor cell membranes, 521 Oregon green-labeled poly(ethylene glycol), 189 Organelle-specific ligand, 395 Organic dyes, vs. quantum dots, 539–540 Organic fluorophores, limitations of, 99 Organometallic complexes, reduction of, 77 Ornithine transcarbamylase, 6 Orthomyxovirus, eGP defined receptor specifity and cell tropism of, 363 Oxidative stress, 457 hierarchical model of, 110 induction of, 93 P10C peptide, 328 P10C/β-gal complex, 328 Paclitaxel, 392 Paclitaxel, DQAsomal encapsulation of, 392–393 Paclitaxel-induced apoptosis, 392 PAMAM dendrimere conjugates, ibuprofen, 238 dendrimere conjugates, methylprednisolon, 238 surface modification, 238 methotrexate conjugates, 242 125 I-labeled, 240 tacking with fluoroisothiocyanate, 236 transepithelial transport of, 239 PAMAM. See Polyamidoamine dendrimere PAMAM-oligonucleotide complexes, 241 Pancreatic cancer cells, 515 Paracellular transport, 232 Paraformaldehyde, 187

bindex.indd 583

583

Paramyxovirus, eGP defined receptor specifity and cell tropism of, 363 Parkinson disease (PD), 105 Particle uptake, nonspecific, 198 Partition coefficient, 387 Parvo viruses, 359 Passive diffusion, 232 Patch clamp, 20 Pathogens, tagged with QDs, 537 PCI. See Photochemical internalization PCI-mediated gene delivery, 316 PD. See Parkinson disease pDNA decondensation, 132 PECAM-1 antibodies, 454 epitopes, 461 PECAM-1. See Platelet-endothelial adhesion molecule-1 PEG-coated quantum dots, 168 PEGylated polyplexes, 216 PEGylated QDs, encapsulation in lipid vesicles, 547 PEGylated QD-tagged antibody, 555 PEI. See Polyethylene imine PEI/DNA nanocomplexes active transport of, 338 retrograde transport of, 338 Penetration, 23, 276, 325, 404 Pep-1-/cargo complexes, 326 Pep-1-mediated protein delivery, 326 Peptide(s) biomaterials, 323 cargo interactions, 323 mediated endocytosis, 65 mediated internalization, 483–487 membrane disruptive, 131–132 tagged with QDs, 537 Peptide and protein delivery by CPPs, 406 P-glycoprotein, 241 pH targeted delivery, 247 Phage display library, 328 Phagocytosis, 21, 56, 98, 124, 196, 232, 451–453, 537–539 Pharmacokinetic study of QD705 quantum dots, 87 pH-dependent eGPs, 365 pH-dependent transmittance transition of PHSM, 255

7/30/2010 2:16:32 PM

584

INDEX

Phenyl arsine oxide, 59 Phenylalanine glycine (FG) motif, 50 pH-insensitive enveloped viruses, 365 Phosphine oxides, 83 Phosphines, 83 tertiary, 74 Phosphonioalkylthiosulfate zwitter ions, structure of, 81 Phosphoroamidate morpholino oligomers, covalent conjugates with CPPs, 410 Photoacoustic tomography, 99 Photobleaching, 536 Photochemical delivery of macromolecules, 314–317 Photochemical internalization (PCI), 130, 311–322 of dendrimer-based drug delivery, 317 -mediated gene delivery, 316 Photodynamic therapy, 312–313 Photosensitizer, 130, 312–313 Photostability, 18, 540 Photothermal therapy, 100 Photothermally activated drug release, 523 Photothermolysis, 520 Phototoxicity, 317 pH-sensitive copolymers of His and Phe (PHP), 254 linear poly(amido amines), 219 polymeric nanotechnology, 247–262 polymers, 132–133 PHSM/f. See Mixed pH-sensitive micelles containing targeting folate Phthalocyanine, 313 di- and tetrasulfonated aluminium, 314 pH-triggered micelle destabilization, 251 Physiosorptive coatings, 510–511 Pick disease, 110 Picoinjection, 20 Picornaviruses, 359 Pinocytosis, 21, 56, 232, 537–539 Plasma membrane, labeling of, 181 Plasmadesmata, 196 Plasmalemmal sodium-proton exchanger NHE1, 455 Plasmid DNA delivery by CPPs, 408

bindex.indd 584

Plasmid DNA polyplexes, intracellular fate of, 123–142 Plasmodium falciparum, 6, 390 Plasmon resonant light-induced heat release, 100 Plasmon-enhanced emission, 516 Plasmon-resonant gold nanorods, 507–533 Plasmon-resonant light scattering, 515 Plasmon-resonant nanoparticles, photothermal action of, 520–522 Platelet-endothelial adhesion molecule-1 (PECAM-1), 453 Platinum chloride, 77 PLL. See Poly-L-lysine PMSA, QD-PEG conjugated, 556 PNA, 492 PNIPAAm. See Poly(N-isopropylacrylamide) p-Nitrophenyl phopshonate, 546 pNPP. See p-Nitrophenyl phopshonate Point spread function (PSF), 162 Poly(His-co-Phe), 254 Poly(L-histidine), 250 -based micelles, 250–254 Poly(L-histidine-co-L-phenylalanine)based micelles, 254–256 Poly(N-isopropylacrylamide), 523 Poly(N-vinylpyrollidone-co-maleic anhydride) copolymere, 240 Poly( -aminoesters), 219 Polyacrylamide, 15 Polyamidoamine dendrimere (PAMAM) -oligonucleotide complexes, 241 superfect, 126 Polyamidoamine dendrimers, 232 Polyarginine (R9), cholesterolconjugated, 419 Polycations, NLS-modified, 495 Polydispersity indices of dendrimers, 231 Polyethylene imine (PEI), 6, 58, 123, 213, 482 branched, 125 /DNA nanocomplexes, 338 linear, 125 single particle tracking of, 170 PolyHis, acid-base titration curves of, 252–253 PolyHis. See Poly(L-histidine)

7/30/2010 2:16:32 PM

INDEX

PolyHis-b-poly(ethylene glycol), 250 acid-base titration curves of, 252–253 fusogenic characteristics of, 253 proton buffering characteristics of, 253 Poly-L-lysine, 123, 482 histidylated, 482 Polymeric delivery systems, 104 Polymeric nanoparticles, 93 Polymers, pH-sensitive, 132–133 Poly-N-isopropylacrylamide-coacrylamide co-polymer, 78 Polyplex(es), 123, 209, 482 adhesion to syndecans, 124 cell entry of, 123 cytoplasmic trafficking of, 123 disassembly, 214 endolysosomal escape of, 123, 129–132 extracellular unpacking, 216–217 intracellular reducible, 133–134 intracellular routing, 127–129 intracellular unpacking, 217 nuclear import of, 134–136 packing, 215 self-assembly, scheme of, 208 targeting ligands for, 127 uptake pathways, schematic illustration of, 125 Polystyrenesulfonate (PSS), 510 Polyvinylpyrrolidone copolymer, 240 nanoparticles, 372 Pore slit model, 47 Porphin, 313 Porphyrins, 312 Potassium depletion, 483 Poxviruses, 359 Pre-fixation methodologies, 188 Proapoptotic peptide, His6 tagged, 278 Probes encapsulated by biologically localized embedding, 16 Probes, intracellular, 18 Pro-Jet™, 324 Prostate specific antigen, 511 Protamine, 496 Protease labile linkers, 546 Protein delivery into cells, 266–267 Protein kinase C, 58 Protein sponge effect, 126 Protein transduction, 22

bindex.indd 585

585

Protein transduction domain (PTD), 23, 267, 324, 403, 477 -mediated nonendocytic delivery, 477 Proteins, conjugated to QDs, 546 intracellular delivery of, 323–336 Proteoglycans, 124 -deficient mutant cells, 481 Protists, 21 Proton sponge effect, 129, 248, 489 Protonable polymers, 249 Proton-sponge polymers, 213 PSF. See Point spread function PSS. See Polystyrenesulfonate PTD. See Protein-transduction domain Pulmonary endothelium, 454 PULSin™, 324 Pyrenebutyrate, 199 QD (quantum dot(s)), 8, 93–121 anionic dihydrolipoic acid-capped CdSe/ZnS, 62 astrocyte response to, 96 bioconjugation techniques, 542 biodistribution in the whole body, 94 CdSe, 60 CdTe, 96 coating, toxicity, 556–557 conjugated Luciferase, 555 cytotoxicity of, 556–557 delivery by cell penetrating peptides, 410–412 DNA conjugates, delivery to blastomers, 555 DNA, FISH, 544 DNA, Lipofectamine 2000 mediated transfection, 545 duplex DNA conjugates, 544 encapsulation in lipid bilayers, 547 epidermal growth factor receptor, 63 epigenetic changes, 97 fluorescence in situ hybridization, 552–554 fluorescent, 61 FRET, 553 full-width half-maximum of, 539 histone interference, 97 in vivo imaging in live animals, 554–556

7/30/2010 2:16:32 PM

586

INDEX

QD (quantum dot(s)) (cont’d) induced cellular damage, 94 induction of anti-apoptotic genes, 110 induction of pro-apoptotic genes, 110 intracellular trafficking of, 168–169 intravenous administration, 95 labeling, 535–567 labeling techniques for therapeutically active molecules, 541–543 ligand bioconjugates, 55–72 ligand-bound, 63 lipophilic, 61 loaded liposomes, 547 maleimide, 546 micellization, 542 model drug carriers in SPT studies, 168 modification with thiolated siRNA, 546 multivalent ligand, 67 naked, 94 near-infrared, in vivo imaging, 537 near-infrared emission, 95 nPP, 546 oligonucleotide FISH probes, 552 PEG-capped, 65 PEG-PMSA, 556 properties and synthesis of, 540–541 renal clearance of, 95 ricin B, 65 streptavidin-coupled, 63 subcellular distribution of, 94 tagged with plasmid DNA, 544 tagging, 537 targeted delivery, 546 TAT-peptide-coupled DHLA, 65 thioglycolic acid-capped CdTe, 63 Toxin-conjugates, 64 versus organic dyes, 539–540 visualization, 539–540 QD-based intracellular trafficking of antibodies, 548–551 DNA, 543–544 drugs, 548–551 liposomes and polymers, 547 neurotransmitters, 548–551 oligonuclotides, 544–545 pathogens, 548–551 proteins and peptides, 546

bindex.indd 586

siRNA, 545–546 targeting ligands, 547 QD-siRNA conjugates, 546 QD-streptavidin, 542 Quantitative structure-activity relationship (QSAR) models, 386–390 application to nanoparticles, 200 correlational models, 195 decision rule models, 195 Quantum confinement, 541 Quantum dots. See QD R8. See Octaarginine R8 peptides, complexation to DNA, 483 R8-EGFP, 274 R8-lip. See R8-modified liposomes R8-Lip-HD, 485–486 R8-MEND, 486 sugar-modified, 496 R8-modified liposomes, 483 Rab5, 182, 368 Radiofrequency radiation, 86 Radixin, 456 Rafts, 439 Raman scattering, surface-enhanced, 5 Ras-superfamily, 57 Rat insulinoma RINm5F cells, 258 Rath peptide, 327 Rath/cargo complex, 328 Reactive nitrogen species (RNS), 109 generation by metallic nanoparticles, 109 Reactive oxygen species (ROS), 94 generation by metallic nanoparticles, 109 Real-time particle tracking, 161–175 Receptor internalization, 56 Receptor-mediated DNA delivery, 482 Receptor-mediated endocytosis (RME), 149, 212, 482 Receptors bound by eGPs, 362–364 Recombinant single chain antibody-virus eGP fusion, 364 Recycling endosomes, 182 Redox-sensitive transcription factors, 109 Reducible polyplexes, 133–134 Reductive degradation, 219

7/30/2010 2:16:32 PM

INDEX

Resolution, spatial, 17 Resonant light scattering, 515 Retrograde motion, 343–344 Retrograde recruitment system, 337 Retrograde transport, 64 Retrovirus, 209 eGP defined receptor specifity and cell tropism of, 363 RGD, targeting ligand, 483 tripeptide, 5 Rhabdovirus, eGP defined receptor specifity and cell tropism of, 363 Rho-dependent kinase (ROCK), 455 Riboflavin, 5 Ribosome inactivating proteins, 185 Ricin, 64 Ricin B subunit, 64 Ricin B:QDs, 65 RISC. See RNA-induced siRNA complex RME. See Receptor-mediated endocytosis RNA-induced siRNA complex (RISC), 545 RNS. See Reactive nitrogen species ROCK. See Rho-dependent kinase ROS. See Reactive oxygen species RRPs. See Arginine-rich peptides Ruffling, cell surface, 479 SAINT-2/DOPE lipoplexes, 481 Saponin, 188 SAR. See Structure-activity relationship Scanning ion conductance microscopy (SICM), 20 Schizophrenia, 110 Scrape loading, 21 Second harmonic generation, 516 Secretory organelles, markers of, 180–186 Secretory pathways, 177–192 Selectins, turnover in endothelium, 451 Self-unpacking gene delivery scaffolds, 207–230 Self-unpacking materials, 218–221 Semiconductor quantum dots, 18, 84. See also QD

bindex.indd 587

587

SE-model. See Stretched-exponential model Sendai virus, 369 Senile plaques, 107 Sensors fiber-optic, 16 intracellular, 18 optochemical, 16 pulled fiber, 16 SERRS. See Surface enhanced resonance Raman spectroscopy SERS-based imaging, 519 Serum endonucleases, 216 Serum stability of gene delivery vehicles, 210–212 Severe combined immunodeficiency, gene therapy of, 208 Sheets, hexagonal, 7 Shielded polyplexes, 127 Shiga toxin, 57, 64, 417 Short single wall nanotubes, 463–464 SICM. See Scanning ion conductance microscopy Signal peptides, 26 Silica, functionalization of, 512 Silica sol-gel, 15 Silica-loaded liposomes, 547 Silver acetate, 77 Silver nanoparticles, 76 tiopronin-coated, 80 Single motor parameters for kinesin and dynein, 297 Single-particle tracking (SPT) analysis, 127, 161, 513 of carbon nanotubes, 153–155 Singlet oxygen, 130, 313 SiO2 shell, 512 siRNA delivery by cell penetrating peptides, 408 delivery into cytosol, 266 tagging with QDs, 537, 545 therapeutics, 241 si-RNA-TAT conjugates, 409 SKMES-1 cells, 44 Small GTPase Tab5, 181 Small-molecule QSAR models, application to nanoparticles, 200

7/30/2010 2:16:32 PM

588

INDEX

Small-molecule xenobiotics pattern of localization in live cells, 197 QSAR models, 193–206 Smallpox, 359 Smart stimuli-responsive nanocarriers, 412 SNARE, 181 SOD. See Superoxide dismutase Solute binding, 38 Solvent fixation, 188 Sonication, 20 Sorting endosome, 181–182 Spatiotemporal image correlation spectroscopy, 128 Spheres, IgG-coated, 463 Spherical dendrimers, 232 Spot photobleaching, 386 SPT. See Single-particle tracking Squamous cell carcinoma of head and neck, gene therapy of, 208 ssDNA tagged with gold particles, 544 Stains, subcellular, 4 Starling forces. See Transcapillary filtration pressure Stearyl triphenylphosphonium (STPP) liposomes, 394 Stearylated R8, 483 Stem cells, human embryonic mesenchymal (hMSCs), 27 Stoeber process, 512 Stokes-Einstein equation, 37 STPP liposomes. See Stearyl triphenylphosphonium liposomes STPP-PEG5000 liposomes, 395 Streptolysin O, 188 Stretched-exponential (SE) model, 39 Stroke, 457 STR-R8. See Stearylated R8 Structure-activity relationship, 3. See also Quantitative structure-activity relationship models Subplasmalemmal vesicles, 453 Sulfadiazine (SDZ), 258 Sulfadimethoxine (SDM), 258 Sulfamerazine (SMZ), 258 Sulfamethizole (SMT), 258 Sulfonamide oligomers, 258 Superoxide anion, 457

bindex.indd 588

Superoxide dismutase (SOD), 110, 458 conjugated to anti-CAM, 458 Superresolution microscopy, 29 Surface chemistry, multi-layer multifunction, 17, 26 Surface electric charge, 199 Surface enhanced resonance Raman spectroscopy (SERRS), 76 Surface plasmon resonance, 76, 83 Surface-enhanced Raman scattering, 99, 519 SV40, 57 internalization of, 479 T antigen, 274, 494 SW480 cells, 64 SWNT. See Short single wall nanotubes Synaptic plasticity, 110 Syndecans, 347, 481 Syndecan-1 clustering, 348 Syntaxin, 179, 181 Syntenin, 347 Synthetic viruses, 124 T7 RNA polymerase, inhibition of, 82 Target, molecular, 2, 4 Targeting, 2 Targeting ligands, chimeras with CPPs, 410 Targeting moieties, endocytic, 5 TAT antigen-transduced dendritic cells, 407 biotinylated, 276 complexation to DNA, 483 Cre recombinase assay, 271 fused to tumor antigen, 407 fused β-galactosidase, 4–5 peptide motif, 65 peptide-coupled DHLA-QD conjugates, 65 peptide-siRNA conjugates, 409 protein, 22 TAT-S-S-asparaginase, FITC-labeled, 275 T-cell response and CHP-HER2, 108 Teleostean gelatin, 186 Tetradocylammonium bromide, 77 Tetralamellar multifunctional envelopetype nano device (T- MEND), 7, 496

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INDEX

Tetraphenylporphines, sulfonated, 314 Texas Red, 186 Texas Red-labeled wheat germ agglutinin, 181 Theranostics, 29 Thermal injury, gold nanoparticle-based assessment of, 100 Thioglycolic acid, 60 Thiolates, organic, 74 Thioredoxin, 111 Ti : sapphire laser, 517 Time-lapse multicolor fluorescence imaging, 347 Tiopronin (N-(2-mercaptopropionyl) glycine), 82 gold nanoparticles, 104 TIRF. See Dual-color total internal reflection fluorescence microscopy TMA. See Trimethyl(mercaptoundecyl) ammonium T-MEND. See Tetralamellar multifunctional envelope-type nano device TNFα. See Tumor necrosis factor alpha Togavirus, eGP defined receptor specifity and cell tropism of, 363 TOPO coating. See Trioctyl phosphine oxide coating Toxicology of metal nanoparticles, 86 Toxin-quantum dot conjugates, 64 TPL. See Two-photon excited luminescence TPP. See Triphenyl phosphonium cations Trafficking, cellular, 4 Trans-acting nuclear DNA import, 135 Transcapillary filtration pressure, 437 Transendothelial channels, 439 Transfection efficiency, 123 cell cycle dependence of, 134–135 Transferrin, 5, 127, 477 receptor, 63, 182 tagged with QDs, 537, 547 Transgene carrying capacity, 209 Transgenic reporter mice, 97 Trans-Golgi network, 185 Translational diffusion coefficient, 386 Transmembrane pathways, 439–440

bindex.indd 589

589

Transport bidirectional, 294 facilitated, 47 passive, 47 retrograde, 64 Transportan, 276, 325, 404 Transverse plasmon resonance, 508 Trimethyl(mercaptoundecyl)ammonium (TMA), 82 Trimethylammonium alkylthiol, structure of, 81 Trioctyl phosphine oxide (TOPO) coating, 542 Triphenyl phosphonium cations (TPP), 394 surface-coated liposomes, 394 Triphenylphosphonium, 4, 201 Trojan horses, 403 Trypan blue, 27 T-tubules, 440 Tubulin tracker green, 536 Tumor border, sensitivity to PDT, 314 Tumor cell membranes, optoporation of, 521 Tumor cells, hypoxia conditions, 63 Tumor immunotherapy, 106 Tumor necrosis, gold nanoparticle-based assessment of, 100 Tumor necrosis factor alpha (TNF ), 85 Two-dimensional (2D) particle tracking, 162 Two-photon excited luminescence (TPL), 99, 513 activities of Au nanostructures, 516 Tyrosine phosphorylation of CAM cytosolic domain, 455 Ultrashort laser pulses, 516 Vaccinia virus, 209 Vascular cell adhesion molecule-1 (VCAM-1), 453 epitopes, 453 Vascular extravasation, 212 Vasculature, leaky, 3 Vasodilation, 110 VCAM-1. See Vascular cell adhesion molecule-1 VDAC, 441

7/30/2010 2:16:32 PM

590

INDEX

Vehicle unpacking, 214 Ventricular cell structure of, 434 ultrastructure, changes associated with heart failure, 441 Ventricular myocytes, 440 Venular endothelium, 438 Vero cells, 185 Vesicle(s) associated membrane proteins, 181 cell membrane-derived, 7 endosomal, 7 acidified, 22 Vesicular endocytic pathways, 451–453 Vesicular fusion, 181 Vesicular stomatitis virus, EM of, 358 Viral gene delivery vehicles, 208–209 Viral gene vectors, comparison to polyplexes, 123 Viral spike proteins, 359 Viral vectors, drawbacks of, 475 Virosomes, 370–371 FITC-Dx loaded, 372 loading of, 371–372 reconstitution of, 371–372 Virus(es) classification, 358–359 eGP, preparation of, 371 envelope-coated nanoparticles, 357–383

bindex.indd 590

envelope glycoprotein-coated polymers, 372–374 envelope glycoproteins, recombinant sources of, 374 fluorescent labeling of, 169 inactivated, 369–370 interaction with cell-surface receptors, 362–364 intracellular trafficking of, 290–291 mimetic (VM) nanogels, 256–257 pseudotypes, 369 Viscosity, 38 bulk, 38 cytoplasmic, 5 fluid-phase cytoplasmic, 38 microscopic, 38 Visible light, 85 VM nanogels. See Virus-mimetic nanogels VTW-based amphipatic peptide, 329 Wheat germ agglutinin (WGA), 181 Wortmannin, 455 Wrapping time, 103 Wr-T peptide, 327 YTA2 peptide, 330 Zinc sulfide, 77 ZnS coating, disulfide linkage of, 542 ω-thioacetylalkylphosphonium salts, structure of, 81

7/30/2010 2:16:32 PM

O NH

2

O

O

O

N

N H

O N H

O

R functional matrix, chemical groups

lon+ polymer matrix, (silica sol-gel, polyacrylamide, liquid polymer)

NH

O

O

O

O 2 NH

N O NO H

O O

O N

N HO

S

NH 2

O

ionic matrix, comonomer additives +/–

SP* Lipid Multi-layer multifunction surface chemistry

CPP attachment

S

Fluorophore(s) (incl. FRET pairing), ionophores, enzymes hv

O

S

hv

O 2

S

O

N

Nanosensor function

Encapsulation in liposomes

PEG CPP *Signal Peptide

Figure 2.1. Nanosensor schematic.

Figure 2.2. Confocal fluorescence microscopy images of Tat-conjugated nanosensors delivered to (from left): CHO cells incubated with Ca2+-sensitive nanosensors (green) and DRAQ5 nuclear stain (blue); GH4 pituitary lactotrope cells loaded with Tat conjugated Ca2+-sensitive nanosensors; A172 human glioblastoma cell loaded with fluorescein nanosensors; human mesenchymal stem cells (hMSCs) loaded with rhodamine B nanosensors (red) and co-stained with CD105:FITC membrane stain (green) and DRAQ5 nuclear stain (blue).

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Figure 2.3. Confocal fluorescence microscopy images of Chinese hamster ovary K1 (CHO-K1) cells containing: (left) Tat-FITC conjugate (green) which do not show colocalized fluorescence with endosomes/lysosomes stained with LysoTracker Red® when loaded into CHO-K1 cells; (middle) Tat-conjugated nanosensors and LysoTracker Red® showing colocalized fluorescence (orange color) indicating that the nanosensors are residing in acidified endosomes; (right) Tat-FITC conjugate (green) and Tat-TRITC-nanosensors (red) loaded into a single cell. The overlay of these two channels shows no colocalization of fluorescence of the two different cargo sizes mediated by Tat peptide delivery.

104

104

104

A

B

C PE

103

PE

103

PE

103

102

102

102

101

24.0 73.4

0 1.77

100 0 10

101

nic

102

103

104

101

20.3 60.5

0.01 1.43

100 0 10

101

2 nic 10

103

104

101

94.4 0.50

4.01 0.17

100 0 10

101

2 nic 10

103

104

Figure 2.4. Use of cell surface markers to confirm cellular identity. (Top) Simultaneous detection of CD29 expression on hMSC containing Tat-functionalized FITCnanosensors. (A) Isotype control showing only FITC fluorescence coming from internalized nanosensors. (B) CD29 second antibody control showing only FITC fluorescence coming from internalized nanosensors. (C) hMSC containing Tat-functionalized FITC nanosensors and stained with CD29-PE. Both signals are detected, as evidenced by the shift along the y-axis of the cell population. (Bottom) hMSC cells loaded with rhodamine B nanosensors (red) and counterstained with mouse anti-human CD105:FITC antibody (green).

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A

Lys Lys Lys Lys Lys Lys Lys Lys Lys Lys

Lys Lys

Lys

Lys Lys

Lys Lys

Lys

Lys Lys

Lys

Lys

B

Lys

Lys

Lys

Lys Lys

Lys Lys

Lys Lys

Lys Lys

Lys

Lys

Gly

Lys

NH2

Lys

Lys Lys

Lys

Lys Lys Lys Lys

Lys

Core Lys Lys Branch Surface

Lys

Lys Lys

Lys Lys

Lys Lys

Lys Lys

Lys Lys

Lys Lys Lys

Lys

Control D

t=30min

10 μm

E

10 μm

G

t=2h

C

Lys

t=1h

t=2h

10 μm

F

10 μm

H

10 μm

t=15min

t=1h

10 μm

I

10 μm

t=4h

10 μm

Figure 3.1. The structure (A) of the sixth generation polylysine dendrimer used in this study and a time-dependent study of the interaction between the dendrimer (green fluorescence) and Caco-2 cells (B–I): the cells were incubated with complete media containing 0.1% (w/v) of dendrimers for 15 min (C), 30 min (D), 1 h (E, F), 2 h (G, H), and 4 h (I). The cells were observed under confocal microscopy. In the control (A), the cells were exposed to the complete medium without dendrimer. The dendrimer is found to attach to the cell membrane periphery (red arrows) as a result of electrostatic attraction (C, D). At 1 h the dendrimer distributes toward the cell nuclei in all regions of the cells (E, F). At longer incubation times (G–I), the dendrimer appears to be concentrated on the surface of the cells. This technique cannot prove internalization.

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Fluorescent Intensity

60 50 40 † Boxes in both channels are regions of interest (ROIs)

30

ROI 5 ROI 6 ROI 7 ROI 8 ROI 9 ROI 10

20 10 0 0

20

40 60 Time (min)

80

100

Figure 3.2. An overlay of the compressed z-stack images of the dendrimer uptake process in living Caco-2 cells. The cells were incubated with medium containing 0.1% dendrimer (represented in green) and images collected every 5 min before (A) and after addition of dendrimer (B–F). Images presented are at time t = 5 (B), 15 (C), 30 (D), 40 (E), and 90 (F) min of incubation. Data were analyzed over the time and plotted in (G); the inserts show the positions of regions of interest (ROIs, white boxes) for data analysis. The mean fluorescence signal in each ROI was measured from the projected z-stack images at different time points with background subtraction. The dendrimer was found in the cytoplasmic compartment 5–35 min after exposure to the dendrimer (ROI 9 and 10). The concentration of the dendrimer declined after 30 min, whereas the relative fluorescent intensity of the dendrimer inside the nucleus was detected (ROI 5–8). The cell nuclei were stained with Hoechst33342 represented in blue. The scale bar is 20 µm.

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A

B

C

D

Relative fluorescent intensity (F)

E

90 80 70 60 50 40 30 20 10 0 0

20

40

60

Time (min)

80

100

ROI 2 ROI 3 ROI 4 ROI 5 ROI 6 ROI 7 ROI 8 ROI 9 ROI 10 ROI 11 ROI 12 ROI 13

White boxes are regions of interest (ROIs)

Figure 3.3. An overlay of the compressed z-stack images of the dendriplex uptake process in living Caco-2 cells. The cells were incubated with complete medium containing dendriplexes (represented in green) and images were collected every 5 min before (A) and after adding the dendrimer (B–D). Images presented here are at time t = 5 (B), 30 (C), and 60 (D) min of incubation period. Data were analyzed and plotted in (E) and the insert pictures show the positions of regions of interest (ROIs, white boxes) for data analysis in both channels. The mean fluorescent signal in each ROI was measured from projected z-stack images taken at different time points, with background subtraction. The dendriplexes were found to be taken up into the cytoplasmic compartment (ROI 2–3, 7, 9, 12–13) but there is no uptake of the dendriplexes into the nuclear compartment within 2 h of incubation (ROI 4, 8, 10).

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A

Cell radius (h) B 4 1

Dendriplexes Dendrimer

2 3

Cell membrane

Nuclear membrane

FG surface

C 1

Cytoplasmic filaments Threading area

2

Passive transport

Selectivity filter Nuclear filaments

3

Active transport

Figure 3.4. An uptake model for dendrimer and dendriplexes: (A) demonstrates the sieving effect of the cytoplasmic meshwork with a cutoff size of 100 nm in diameter. The crowded condition and the obstruction effect cause the slow diffusion of the dendrimer and dendriplexes after endocytosis (X–Y) or passive diffusion (Y) through transient nanoholes, 1–10 nm in diameter, illustrated in (B). The dendrimer and dendriplexes have to detour in the aqueous compartment excluding crowded volumes. Some of the dendrimer and dendriplexes may interact with cell organelles or cytoplasmic proteins (Z). Passing through the NPC, the DNA released from the endosome ([) may cross the NPC by passive diffusion (X). As small as 6.5 nm, the dendrimer can diffuse through the central pore having a cutoff size of 8–10 nm (Y), whereas the larger of the dendriplexes, 107 nm, may employ active transport (Z). (Part CX–Z is modified from Peters [50]; other parts are original.)

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DHLA-QDs 1h:

EEA1

CD63 merged

EEA1

CD63 merged

10 μm

DHLA-QDs

10 μm

10 μm

4h:

10 μm

10 μm

10 μm

Figure 4.2. Intracellular location of dihydrolipoic acid (DHLA)-capped quantum dots (QDs, Em. 585 nm) in HeLa cells. The DHLA-QDs (30 nM) in complete growth medium were added to the HeLa cells and incubated at 37 °C for 1 and 4 h, respectively. Yellow and magenta colors in the merged images indicate colocalization with the early endosomal marker EEA1 and with the lysosomal marker CD63, respectively. Bars, 10 µm.

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Figure 4.3. Intracellular location of ricinB : QD655 bioconjugates in HeLa cells (left panel) and HEp-2 cells (right panel). (See text for full caption.)

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Figure 6.3. Two-photon luminescence (TPL) imaging of primary cortical neurons. TPL imaging of primary cortical neurons treated 21 days in vitro (DiV 21) with gold nanorods (rod GNP) for 4 hours prior to imaging (CTAB—cetyl-trimethylammonium bromide). Neurons were also stained with mitotracker green FM. Scale bar in the control panel is representative for both panels.

Figure 6.4. Cellular internalization of GNPs. Microglial cells (N9) were treated for 4 hours with two different functionalized rod GNPs (rod GNP-CTAB, rod GNP-INS) and urchin-like GNPs (GNP-URCHIN). Cells were fixed with paraformaldehyde and subsequently analyzed for the internalization of GNPs. (*Nanoscale hyperspectral images, where red signals denote a positive match with the spectral profile of the respective GNP, were performed using Cytoviva Image Analysis Software.)

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CHP

CHPNH2

0

1h

2h

3h

Figure 6.5. Cellular uptake of TRITC-labelled CHP-nanoparticles. N9 microglial cells were treated with tetramethyl rhodamine isothiocyanate (TRITC)-labeled CHP nanogels (12 nM) for 3 hours. Fluorescent images were taken at each time point: before treatment (T = 0 min) and at 3 time points therafter (T = 1, 2, 3 hours post-treatment). Scale bar in top left panel (20 µm) is representative for all images.

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3h

5h

+folate blocking

Fluor channel

FA-SWCNTs 1h

20 μm

20 μm

20 μm

20 μm

20 μm

20 μm

20 μm

20 μm

20 μm

20 μm

20 μm

Merged

TD channel

20 μm

Figure 8.3. Multichannel confocal images of Hep G2 cells after incubation with FA-SWCNTs for 1 h (left, first column), 3 h (left, second column), and 5 h (left third column). The green fluorescence from the outside to the inside of cells shows the uptake pathway of FA-SWCNTs into cancer cells. Right-hand column: multichannel confocal images of Hep G2 cells incubated in FA-SWCNTs for 3 h with pretreatment of free FA blocking. The concentrations of FA-SWCNTs in all groups are 20 μg/mL. (Copyright IOP. Reproduced with permission from Ref. 57.)

Figure 8.4. The cell penetration of FITC-PEG-SWCNTs and its nuclear accumulation in mammalian cells. (See text for full caption.)

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Figure 9.1. Trajectories of nonviral gene vectors in a live cell tracked using SPT. The observed trajectories are highly heterogeneous, which suggests that different transport mechanisms may be involved. (Reproduced from Ref. 53 with permission.).

100

%Cell entry

80 60 40 COOH OH NH

20 0 0

1

2 Time (h)

3

4

Figure 13.3. A549 cell entry profile of G4-NH2, G4-OH, and G3.5-COOH PAMAM dendrimers [14]. Rate of cell entry is determined as percent of (cell entry at time t/cell entry at 3 h). The cellular uptake was quantified by flow cytometry using fluorescence. A comparison of the time-dependent fluorescence intensity levels indicates that rate of cell uptake was highest for the cationic dendrimer, followed by anionic and neutral dendrimers. The cell uptake for cationic dendrimer plateaus off after 1 hour, unlike with the other two dendrimers where the cell entry increased more or less linearly with the treatment time, as shown in the graph.

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(A)

(B)

(C)

Figure 13.4. Confocal microscopic images (63×) of A549 lung epithelial cells after treatment with different dendrimers and LysoTracker for 5 min followed by chase for 30 min in serum-free medium devoid of any dendrimers. Images indicate cells treated with (A) FITC-labeled PAMAM-G4-NH2 dendrimer and LysoTracker. (B) FITC-labeled PAMAM-G4-OH dendrimer and LysoTracker. (C) FITC-labeled PAMAM-G3.5-COOH dendrimer and LysoTracker. The left-hand panel shows the green fluorescence from the dendrimer, the middle panel shows the red fluorescence from the LysoTracker, and the right panel shows the yellow fluorescence due to the colocalization of green dendrimer–FITC and red LysoTracker in the lysosomes. LysoTracker dye preferentially partitions to the lysosome, giving an intense red color in the acidic environment of the lysosomes. From the colocalization studies, it is evident that the dendrimers are in the lysosomes within 30 min of their transport from the cell membrane [14].

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(a) 14

(b) 120

NaCl polyHis5kDa-b-PEG2kDa polyHis5kDa polyHis3kDa-b-PEG2kDa polyHis3kDa

10

100 Hemolysis (%)

12

pH

8 6 4

PLLA3kDa-b-PEG2kDa

80 60 40 20

2 0

polyHis5kDa-b-PEG2kDa

0

200

400

600

800

0 7.4

7

1 N HCl (mL) (c)

6.75

6.5

pH

polyHis-b-PEG PLLA-b-PEG Proton

pH

pH

pH 7.4–7.0

pH 6.8–6.5

(d)

pH 6.0 (e)

7.6 DHPE

7.4

Lysotracker dye

Merged

7.2 pHt

PHIM-f

7.0 6.8 6.6 6.4

PHSM-f 0

10 20 30 40 50 PLLA3kDa-b-PEG2kDa fraction (wt.%) (g) Tumor volume (mm3)

(f) 120

Cell viability (%)

100 80 60 40 20 0 0.0001 0.001 0.01 0.1 1 DOX concentration (μg/ml)

3500 3000 2500 2000 1500 1000 500 0 0 3 6 9 12 15 18 21 24 27 30 Days after 1st i. v. injection

10

Figure 14.2. (A) Acid–base titration curves of polyHis and polyHis-b-PEG. (Reproduced from Ref. 29. Copyright © 2003, with permission from Elsevier B.V.). (See text for full caption.)

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pH 7.4 pH 6.4 DOX released F: folate F F F PEG Inner shell F F F F F F F pH F F F F F F F F F FF FF F F F BSA outer shell F FF 355 nm 55 nm

(A)

F F F F

F

F

(B) pH 7.4

pH 6.8

0 100

pH 7.4

0 100

pH 7.4

pH 6.4

pH 6.8

0 100

0 100

pH 7.4

pH 6.4

0 100 0 200 400 600 800 0 100 0 200 400 600 800

Particle size (nm)

(D) 20

FITC

Lyso Tracker

Merged

Control NPs VM-nanogels

pH pulses relative rate of DOX (μg/hour)

(C) 15 10 5 0

pH 7.4 pH 6.8 pH 6.4

: appled pH for 1 hour each

Cell viability / %

80

**

**

60 40 20 0

**

**

A2780 A2780/ A2780 A2780/ DOXR DOXR MDR cells MDR cells

Blank DOX-loaded DOX-loaded VM-nanogels Free DOX control NPs VM-nanogels

(F) pH 6.8

pH 7.4

(E) 100

Infected

not Infected

Subsequent Infections

A

B-0

B-1

B-2

B-3

A

B-0

B-1

B-2

B-3

A

B-0

B-1

B-2

B-3

A

B-0

B-1

B-2

B-3

Figure 14.4. VM nanogel: (A) Schematic presentation. (B) pH-modulating size change: A (pH 7.4), B (pH 6.8), and C (pH 6.4). (C) pH-dependent DOX release rate from DOX-loaded VM nanogels. (D) In vitro endosomal escaping activity. (E) pHdependent cytotoxicity of DOX-loaded VM nanogel (white), DOX-loaded control nanoparticles (gray), and free DOX (black) against A2780 wild-type cells and A2780/ DOXR MDR cells after 48-h incubation (DOX dose = 1 µg/mL). (F) Migration of DOX-loaded VM nanogels in A2780/DOXR MDR cells. (Reproduced with permission from Ref. 38. Copyright © 2008 Wiley-VCH Verlag GmbH & Co.KGaA.)

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(A)

(B) entry

egress cargo dynein

tubule micro actin

MTOC

kinesin

nucleus 25nm microtubule

delivery

v(F)

π0

ε(F)

Vf

vf(1–F/Fs)

0.5 Fs 0 0

(A)

vb(1–F/Fs)

(B)

2 4 6 8 10 12 load force F [pN]

unbind. rate ε [1/s]

F

velocity v [μm/s]

Figure 16.1. Intracellular trafficking. (A) Microtubule filaments (black) form long “highways” from the microtubule-organizing center (MTOC) near the nucleus to the cell periphery, while a dense meshwork of actin filaments (cyan) provides “side roads” for short-range traffic. Plus-end directed (blue) and minus-end directed (yellow) molecular motors pull various cellular cargoes along these filaments. Viruses hijack the cellular transport machinery: after entry into the cell, a virus particle (red) recruits motors from the cytoplasm to travel along microtubules toward the nucleus, where it delivers its genetic material (white). In order to leave the cell, the newly assembled virus uses motors to travel back to the plasma membrane. (B) Cargo particle (red) that is transported along a microtubule by one kinesin-1 (blue) and one cytoplasmic dynein (yellow) motor.

(C)

8

ε0exp[F/Fd]

6 4

ε0

2 0

0

2 4 6 load force F [pN]

Figure 16.3. Transport properties of a single motor. (A) A single motor walks along the filament with velocity v, unbinds from the filament with rate ε, and rebinds to it with rate π = π0. If the cargo experiences a load force F, both the velocity v and the unbinding rate ε depend on F. (B) Motor velocity v as a function of load force F as parameterized in Equations 16.2 and 16.3. The velocity decreases with increasing F until it vanishes at the stall force Fs. For superstall forces F > Fs, the motor steps backwards with a relatively small velocity. (C) Motor unbinding rate ε as a function of F as in Equation 16.4. The unbinding rate increases exponentially with increasing F.

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Figure 16.4. Transport by one team of identical motors. Each motor is characterized by a long stalk (grey), by which it is firmly attached to the cargo particle (red), and by two motor heads (blue), which can bind to the filament and then actively pull on the cargo. In this example, the total number of motors is N = 3 but the actual number n of crosslinking motors fluctuates between n = 0 and n = 3 because of thermally induced motor unbinding and rebinding.

(–)

(0)

(0)

(–)

(0)

(0)

(0)

(+)

(+)

Figure 16.6. Transport by two antagonistic teams of motors. The motors with blue heads are plus motors that pull to the right, the ones with yellow heads are minus motors that pull to the left. Both types of motors are firmly attached to the cargo particle (grey) via their long stalks. The total number of blue plus motors and yellow minus motors is N+ = 2 and N− = 2, respectively. The cargo particle exhibits (N+ + 1)(N− + 1) = 9 different states and undergoes transitions between these states arising from the unbinding and rebinding of single motors.

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Cargo Myosin

Kinesin

-

Distance [μm]

6

+

4

2

0 0

(A)

Microtubule

(B)

3

6

9

12

Time t [s]

Figure 16.8. Cargo transport by one active and one “passive” motor. (A) A cargo particle (grey) is crosslinked to a microtubule by one kinesin-1 (blue) and one myosin V (red). The kinesin-1 motor is actively stepping whereas the myosin V motor is passively diffusing along the filament. (B) The trajectory of such a cargo exhibits fast plus motion interrupted by diffusive events. For this example, the total binding time was about 11 s, much longer than in the absence of the “passive” myosin V motor [14].

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Figure 19.2. A sequence of DIC images, selected from a time series [62, Movie 1], demonstrates the directed retrograde motion of positively charged 500-nm amorphous silica particles along the elongated microvilli of alveolar type II epithelial cells in culture. The particles are indicated by color-coded arrows as they appear along the time series, and the cumulative time relative to the first image in the series is shown in the lower right corner of each image. Particles are tracked from one image to the next in the series and their trajectories are plotted in the last frame, where the cell body from which the elongated microvilli originate is outlined with the dashed line. (Adapted with permission from Ref. 62.)

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(A)

(B)

Figure 19.4. (A) Alveolar type II epithelial cells that are transfected with GFP-actin form fluorescent filopodia and microvilli that often contain small fluorescent clusters, which are distinguished from the diffused signal. Following the clusters over time reveals their retrograde motion toward the cell body, as demonstrated by the cluster that is indicated by the arrow in the first two images, which are selected from a time series [62, Movie 6b]. Treating the cells with cytochalasin D leads to fragmented filaments and the arrest of the retrograde flow, as demonstrated in the third and fourth images where the cumulative time relative to the time of applying the drug is shown in the lower right corner. (B) Images of a 500-nm fluorescent particle (red, white arrow) and a GFP-actin cluster (green, yellow arrow) demonstrate that the particle and the cluster travel toward the cell body at the same rate. (Adapted with permission from Ref. 62.)

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(A) 2

1

3

4

5

8

9

10

0 sec 6

7

400 sec (B)

+ + + + + + docking + + + + protein

syndecan-1

(−) (+) actin monomers

actin binding protein

Figure 19.6. (A) A DIC image of the elongated microvillus and a sequence of fluorescence images, selected from a time series [72, Movie 1], demonstrate the engagement of the particle (red), indicated by the arrow, and syndecan-1 molecules (green) in the motion along the microvillus as they become tightly associated (yellow) with each other. (B) The directed motion along the microvillus unravels the coupling of syndecan-1 and the particle with actin filaments across the cell membrane via docking and actin binding proteins. See discussion in the text. (Adapted with permission from Ref. 78.)

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Figure 20.2. Virus anatomy and the three stages of eGP-mediated delivery into cells. Virus particles and eGP-coated nanoparticles each interact with cells identically. The top image indicates the parts of a virus and show interaction with cells through eGP binding to specific receptors. eGPs are shown as trimers, which is typical for most Class 1 and 3 viruses. Actin is often involved in recruiting receptor. The particle is then taken into endosomes (for pH-dependent viruses), where the pH of the vesicle is made progressively more acidic as it moves through the cell, maturing from an early to late endosome. The eGP functions as an environmental sensor and when triggered undergoes a large conformational change to expose previously buried fusion peptides. These embed in the cell membrane and catalyze membrane fusion of the virus and endosomal membrane. Since this occurs at a specific pH, the core is delivered to a specific subcellular location within the cell. For pH-independent viruses, like HIV, membrane fusion may occur at the cell surface but this has recently been challenged and it too may use endosomes but is triggered by other stimuli [50].

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Figure 20.4. eGP-coated nanoparticles enter cells through endocytosis. The set of images shows ecotropic murine leukemia virus eGP-coated nanoparticles (blue) interacting with cells expressing the virus receptor tagged with mStrawberry (red) and GFP tagged Rab5 (green, a marker of early endosomes). Each image is a plane from a z-stack and was taken using a Zeiss LSM 510 confocal microscope. The z-axis depth is indicated at upper left. A combined projection of all z-axis planes is shown at lower right. Arrowheads indicated nanoparticles interacting with receptor (violet), rab5 (cyan), or receptor and rab5 together (white). CPP

CPP-cargo covalent attachment noncovalent complexation CPP modified nanoparticle – +

Clathrin- & Caveolin-independent endocytosis Inverted micelle Cytosol

Direct penetration

Golgi apparatus

Early endosome

Caveolin-mediated endocytosis

Late endosome

Clathrin-mediated endocytosis ER Nucleus

Lysosome

Macropinocytosis

Figure 22.1. Mechanisms of CPP-assisted cytoplasmic drug delivery.

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Clathrin-mediated Endocytosis Dynamin

Caveolae Caveolin

Clathrin coat

Receptor Clathrin-coated pit

Macropinocytosis

Recycling

Caveosome

Clathrin-coated vesicle (CCV)

H+ H+

Ruffling

Actin Recycling

Ligand

?

Macropinosome

H+ ?

Leakiness H+

Late endosome (Low pH) Endosomal release Nuclear delivery

Cytosolic release H+

Lysosome

Lysosome

Nucleus

Degradation

Figure 25.1. Different endocytic pathways. Clathrin-mediated endocytosis (CME) is the major and best-characterized endocytic pathway. Caveolae and macropinocytosis are two clathrin-independent endocytic pathways.

K8-MEND

R8-MEND Fusiondependent Cytosolic release

Macropinosome

Acidification

Acidification

Degradation Recycling

Degradation Recycling Stop cytosolic release

Continue cytosolic release

Figure 25.2. A schematic diagram of endosomal escape of R8- and K8-MEND. After internalization, membrane fusion occurs at a neutral pH in both cases. Upon acidification, the membrane fusion stops in the case of K8-MEND while it continues in the case of R8-MEND. Extensive interactions between endosomal and liposomal membranes occur in both neutral and acidic pH in the case of R8-MEND.

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Figure 25.3. A schematic diagram of mitochondrial delivery via membrane fusion by MITO-Porter. R8, octaarginine; OM, outer membrane; IMS, intermembrane space; IM, inner membrane.

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(A) Direct modification

(B) Insertion of transcription factorbinding sequence

(D) NLS-modified envelope-type nanoparticle

(C) Complex formation with NLS-modified polycation or NLS-peptides

(E) Envelope type nano particle encapsulated with endosome- and nuclear membrane-fusogenic membrane

Figure 25.4. Strategies to overcome a nuclear member barrier. (A) Direct modification of NLS peptides and/or nucleus-targeting proteins to pDNA by chemical linkage or avidin–biotin interaction. (B) Insertion of nuclear-transcription factor-binding sequences (i.e., NF-kB) in the pDNA. (C) Condensation of pDNA with NLS itself or NLS-modified polymers. (D) Surface modification of NLS on the surface of lipid-based particle. (E) Encapsulation of pDNA in NLS-fusiogenic lipid envelope.

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(B) Speed (μm/s)

(A)

(C)

0.1 0.0

–0.1 0 10 20 30 40 50 60 Time (s)

Figure 26.3. Nonspecific cell uptake of CTAB-stabilized GNRs [30]. (A) Overlay of transmission and TPL images of GNRs (green) internalized by KB cell, with nucleus outlined in red. (B) Trajectory of an internalized GNR exhibiting bidirectional motion, with overall displacement toward the cell nucleus. (C) TPL image of GNRs (yellow) segregated within inclusion bodies, 5 days after initial exposure; bar = 20 µm. (Reproduced with permission of the American Chemical Society.)

(A)

(B)

(C)

Figure 26.7. Targeted adsorption and uptake of folate-conjugated GNRs (red) by KB cells overexpressing folate receptors (imaged in transmission mode, grey) [99]. (A) Folate-conjugated GNRs were observed on the surface of KB cells with high density, after 6-h incubation at 37 °C. (B) GNRs were internalized into KB cells and delivered to the perinuclear region after 17-h incubation. (C) No GNR binding or uptake was observed by NIH-3T3 cells, which express folate receptors at a low level. Bar = 10 µm. (Reproduced with permission of Wiley-VCH.)

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(A)

before

(B)

after

(C)

Figure 26.9. “Optoporation” of tumor cell membranes by photothermally active GNRs [99]: (A, B) folate-conjugated GNRs (red) targeted to the membranes of KB cells, before and after a 1-min exposure to a scanning NIR laser (12 J/cm2); (C) ethidium bromide (red) and a Ca-sensitive dye (green) indicate a loss of membrane integrity and high levels of intracellular Ca2+, respectively. (Reproduced with permission of Wiley-VCH Publishing.)

(A) F4/80 antibody

(B) R8-NRs

Figure 26.10. In vivo uptake of R8-conjugated GNRs by macrophages, 30 min after intraperitoneal injection [139]: (A) immunofluorescence staining of F4/80+ macrophages (blue) and (B) TPL image of R8-GNRs (red) inside macrophages (bar = 10 µm). (Reproduced with permission of the authors.)

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QD

Rh

(A)

(B)

(C)

(D)

(E)

(F)

0 min

50 min

100 min

Fluorescence Intensity

(G) 1.2 1 0.8 0.6 RhPE/DNA QD-DNA conjugate

0.4 0.2 0 0

10

20

30

40

50

60

70

80

90

100

Time (min)

Figure 27.2. Comparative study of photostability of QD-DNA conjugated with rhodamine-PE-DNA conjugate (organic dye complex). QDs can be visualized even after 100 min of laser exposure while most of the rhodamine bleaches after 50 min, in CHOK1 cells. The graph (G) represents the fluorescence intensity time plot for the two conjugates after 100 min of continuous laser exposure at 543 nm. QD-DNA conjugates were added as 1.469 × 1013 particles per 2.5 × 106 cells per well. (Bar-25 µm) (From Ref. 11.)

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(A)

6h

(C)

10 h (B)

(E)

24 h (D)

(F)

Figure 27.4. Confocal images of Lipofectamine 2000 mediated transfection studies of QD-DNA conjugates (red) 6 h, 10 h and 24 h postincubation, delivered in CHOK1 cells stained with SYTO-16 nuclear stain (green). Yellow regions in the nuclear/perinuclear region determine the efficient transfection of QD-DNA conjugates added as 1.469 × 1013 particles per 2.5 × 106 cells per well. (Bar = 25 μm). (From Ref. 11.)

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(A) 2.0

PL [a.u.]

1.6 1.2 0.8 0.4

(B)

PL [a.u.]

(C)

1.0 0.8 0.6 0.4 0.2 0.0

520 600 680 Wavelength [nm]

0 480 520 560 600 640 680 720 Wavelength [nm]

520 600 680 Wavelength [nm]

520 600 680 Wavelength [nm]

520 600 680 Wavelength [nm]

Figure 27.5. (A) Fluorescence spectra of the solution (black) obtained by superposition of the spectra at different emission filters for green, yellow, orange, and red QD-labeled oligonucleotides when hybridized to their respective complementary sequence (mechanism is called FISH). (B) Squares indicate presence of QD tagged hybridization probes at different emission wavelength. (C) Individual spectra for different oligonucleotides, each with the solution spectra. (Reproduced with permission from Ref. 78.)

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QD-COOH

QD-PEG

QD-PEG-PSMA

Tumor

Tumor

Tumor

Injection site

Figure 27.6. Live mouse models injected with C4-2 human prostrate tumors treated with (A) 6 nmol QDs only (QD-COOH), (B) 6 nmol QD-PEG, and (C) 0.4 nmol QD-PEG-PSMA. QD-COOH could not be seen, indicating that it was cleared off very fast. QD-PEG can be visualized weakly in the tumor region while a strong signal is seen at the tumor site on injecting QD with targeting ligand (QD-PEG-PSMA), indicating the efficiency of targeted delivery systems. (Reproduced with permission from Ref. 136.)

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