Microencapsulation (Drugs and the Pharmaceutical Sciences)

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Methods and Industrial Abblicalions

edited by Simon Benita The Hebrew University of Jerusalem Jerusalem, Israel

Marcel Dekker, Inc. New York.

Basel Hong Kong

Library of Congress Cataloging-in-Publication Data Microencapsulation : methods and industrial applications/ edited by Simon Benita. p. cm. -(Drugs and the pharmaceutical sciences;v. 73) Includes index ISBN 0-8247-9703-5 (alk. paper) 1. Microencapsulation. 1. Benita,Simon. 11. Series. RS20 1.C3M27 1996 6 15'.1 9 d c 2 0

96- 1569 CIP

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This book is printed on acid-free paper. Copyright 0 1996 by Marcel Dekker, Inc. All Rights Reserved. Neither this book nor any part may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, microfilming, and recording, or by any information storage and retrieval system, without permission in writingfrom the publisher. Marcel Dekker,Inc. 270 Madison Avenue,New York, New York 10016 Current printing (last digit): l0987654321 PRINTED IN THE UNITED STATES OF AMERICA

Preface

Research, development, and sales of drug-delivery systems are increasing at a rapid pace throughout theworld. This worldwide trend will intensify in the next decade as cuts in public health expenses demand lowercosts and higher efficacy. To meet this demand, many efficient drugs currently in use will be reformulated within delivery systems that can be value-added for optimal molecular activity. In addition to the health sector, the cosmetic, agricultural, chemical, and foodindustries operate in an open marketplace where free and aggressive competition demands novel coating techniques with enhanced effectiveness at the lowest possible cost. Currently, microencapsulation techniques are most widely used in the development and production of improved drug- and food-delivery systems. These techniques frequently result in products containing numerous variably coated particles. The exact number of particles needed to form a single administered dose varies as a function of the final particle size and can lie in either the micro- or nanometer size range for micro- and nanoparticulate delivery systems, respectively. The microparticulate delivery systems include mainly pellets, microcapsules, microspheres, lipospheres, emulsions, and multiple emulsions. The nanoparticulate delivery systemsinclude mainly lipidor polymeric nanoparticles (nanocapsules and nanospheres), microemulsions, liposomes, and nonionic surfactant vesicles (niosomes). Generally, the microparticulate delivery systemsare intended for oral and topical use. Different types of coated particles can be obtained depending on the coating process used. The particles can be embedded within a polymeric or proteinic matrix network in either asolid aggregated state ora molecular dispersion, resulting in the formulation of microspheres. Alternatively, the particles can be coated by a solidified polymeric or proteinic envelope, leading to the formation of microcapsules. The profile and kinetic pattern governing the release rate of the entrapped active substance from the dosage form depend on the nature and morphology of the coated iii

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Preface

particles, which need to be established irrespective of the manufacturing method used. Microencapsulation techniques are normally used to enhance material stability, reduce adverse or toxic effects, or extend material release for different applications in various fields of manufacturing. Until now, the use of some interesting and promising therapeutic substances has been limited clinically because of their restrictive physicochemical properties, which have required frequent administration. It is possible that these substances may become more widely used in a clinical setting. if appropriate microencapsulation techniques can be designed to overcome theirintrinsic inconveniences. Investigators and pharmacologists have been trying to develop delivery systems that allow the fate of a drug to be controlled and the optimal drug dosage to arrive at the site of action in the body by means of novel microparticulate dosage forms. During the past two decades, researchers have succeededin part in controlling the drug-absorption process to sustain adequate andeffective plasma drug levels over a prolonged periodof time by designing delayed- or controlled-release microparticulate-delivery systems intended for either oral or parenteral administration. The ultimate objective of microparticulate-delivery systems is to control and extend the release of the active ingredient from the coated particle without attempting to modify the normalbiofate of the active molecules in the body after administration and absorption. The organ distribution and elimination of these molecules will not be modified and will depend only on their physicochemical properties. Ontheotherhand, nanoparticulatedelivery systems are usually intended for oral, parenteral,ocular, and topical use, with the ultimate objective being the alteration of the pharmacokinetic profile of the active molecule. In thepast decade, ongoing efforts have been made to develop systems or drug carriers capable of delivering the active moleculesspecifically to the intended target organ, while increasing the therapeutic efficacy. This approach involves modifyingthe pharmacokineticprofile of various therapeutic classes of drugs through their incorporation in colloidal nanoparticulate carriers in the submicron size range such as liposomes and nanoparticles. These site-specific delivery systems allow an effective drug concentration to be maintained for a longer interval in the target tissue and result in decreased side effects associated with lower plasma concentrations in the peripheral blood. Thus, theprinciple of drug targeting is to reduce the total amount of drug administered, while optimizingits activity. It should be mentioned that the scientific community was skeptical that such goals could be achieved, since huge investments of funds andpromising research studies have in many cases resulted in disappointing and nonlucrative results and havealso been

Preface

V

slow in yielding successfully marketed therapeutic nanoparticulate dosage forms. With the recent approval by health authorities of a few effective nanoparticulate products containing antifungal or cytotoxic drugs, interest in colloidal drug carriers has been renewed. A vast number of studies and reviews as well as several books have been devoted to thedevelopment, characterization, and potential applications of specific microparticulate- and nanoparticulate-deliverysystems. No encapsulation process developed to date has been able to produce the full range of capsules desired by potential capsule users. Few attempts have been made topresent anddiscuss in a single book the entire size range of particulate dosage forms covered in this book. The general theme and purpose here are toprovide the readerwith a current and general overview of the existing micro- and nanoparticulate-deliverysystems and to emphasize the various methods of preparation, characterization, evaluation, and potential applications in various areas such as medicine, pharmacy, cosmetology, and agriculture. The systematic approach used in presenting the various particulate systems should facilitate the comprehension of this increasingly complex field and clarify the main considerations involved in designing, manufacturing, characterizing, and evaluating a specific particulate-delivery system for a given application or purpose. Thus, the chapters, which have been contributed by leading authorities in the field, are arranged logically according to the methods of preparation, characterization, and applications of the various particulate-delivery systems. The first chapter is by C. Thies, a renowned scientist in the field of microencapsulation techniques. To provide an idea of which process is most appropriate for a specific application, the general principles of several microencapsulation processes are summarized and reviewed. This chapter focuses primarily on processes that have achieved significant commercial use. S . Magdassi and Y. Vinetsky present an interesting technique of oil-inwater emulsion microencapsulation by proteins following adsorption of the protein molecules onto the oil-water interface. J. P. Benoit and Drs. H. Marchais, H. Rolland, and V. Vande Velde have contributed a chapter on advances in the production technology of biodegradable microspheres. This chapter deals mainly with the preparation and use of microspheres. The potential o f the various technologies addressed is also discussed, with an emphasis on marketedproducts or those products currently underclinical evaluation. A. Markus demonstrates in his chapter the importance of applying microencapsulation techniques in the design of controlled-release pesticide formulations to meet the multifaceted demands of efficacy, suitability to mode of application, and minimal damage to the environment. The nanoparticulate-delivery systems are introduced by a chapter, authored by myself, B. Magenheim and P. WehrlC, that explains factorial

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design in the development of nanoparticulate systems. This chapter illustrates the application of the experim9ntal design technique not only for optimization but also for elucidation of the mechanistic aspects of nanoparticle formation by spontaneous emulsification. The second part of the book, which focuses on the evaluation and characterization of the various particulate-delivery systems, starts with an important chapter on microsphere morphology by J. P. Benoit and C. Thies. The chapter helps to clarify definitions and differences, which are very often confused. In addition, the chapter illustrates how morphology can becharacterized by usingdifferent techniques. C. Washington provides his valuable expertise in the presentation of the various kinetic models used to characterize drug-release profiles fromensembles or populations of microparticulate-delivery systems. It is worth noting that therelease mechanism of a drug from multiparticulate systems such as microcapsules or microspheres cannot be identified by a study of global release profiles, since it has been shown that overall or cumulative release profiles from ensembles of microcapsules are entirely different from those of single microcapsules. The discrepancy arises from the heterogeneous distribution of the parameters determiningrelease behavior in individual microcapsules, which isbeyond the scope of the present chapter. The following chapter, by P. Couvreur, G . Couarraze, JrP. Devissaguet, and F. Puisieux, presents a very detailed explanation of the preparation and characterization of nanoparticles. The authors first clearly define the morphology of nanocapsules and nanospheres, providing the background, information, and guidelines for choosing the appropriate methodfor a given drug to be encapsulated. K. Westesen and B. Siekmann havecontributed an important chapter on biodegradablecolloidal drug carrier systems based onsolid lipids. These new colloidal carriers differ from the otherwell-known and widely investigated lipidic colloidal carriers, including liposomes, lipoproteins, and lipid or submicron oil-in-water emulsions by exhibiting a solid physical state as opposed to theliquid or liquid crystallinestate of the above-mentionedand well-knownlipidiccolloidal carriers. The authors present the different methods of preparation and point out the advantagesof the novel dosage forms suchas biodegradability, biocompatibility, ease of manufacture, lack of drug leakage, and sustained drug release. Despite three decades of intensive research on liposomes as drug-delivery systems, the number of systems that have undergone clinical trials and become products on the market is quite modest. Even though there have been few successes with liposomes, the need for drug-delivery systems is as acute as ever, and the potential that liposomeshold, although somewhat tarnished, has not been substantially diminished according to R. Margalit and N. Yerushalmi. An interesting and original approach is presented in their chapter on thephar-

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Vii

maceutical aspects of liposomes. Propositions are presented on how at least in some of the hurdles in research and development can be overcome and furthering the substantial strides that have been made in advancing liposomes from the lab to the clinic. An ingenious solution on how the drawis presented by D. Lasic in the backs of liposomes in vivo can be overcome chapter onstealth liposomes. He explains how the stability of liposomes in liposomicidal environments of biological systems presented a great challenge, which was only recently solved by coupling polyethylene glycol to the lipid molecules. An example of the potential of niosomes (a colloidal vesicular system prepared fromnonionic surfactants) for the topical application of estradiol is contributed by D. A. van Hal and J. A. Bouwstra, and H. E. Junginger. Niosomes have beenshown to increase the penetrationof a drug through human stratum corneum by a factor of 50 as compared with estradiol saturated in phosphate buffer solution, making this colloidal carrier promising for thetransdermal delivery of drugs. In thethird part of the book, thepotential applications of the various particulate-delivery systems are presented. The methods of preparation of microcapsules by interfacial polymerization and interfacial complexation and their applications are discussedby T. Whateley, a scientist who is extremelyknowledgeable in this field. The fast-growingfield of lipid microparticulate-delivery systems, particularly lipospheres, is explained and discussed by A. J. Domb, L. Bergelson, and S. Amselem. Lipospheres represent a new type of fat-based encapsulation technology developed for the parenteraldelivery of drugs and vaccines and the topical administration of bioactive compounds. In their comprehensive and exhaustive chapter, N. Garti andA. Aserin underline the potential of pharmaceutical applications of emulsions, multiple emulsions, and microemulsions, and emphasize the progress made in the last 15 years in understanding mechanismsof stabilization of these promising liquid dispersed-deliverysystems that open new therapeutic possibilities. J.-C.Leroux and E. Doelker and R. Gurny in their chapter on the use of drug-loaded nanoparticles in cancer chemotherapy cover the developments and progress made in the delivery of anticancer drugs coupled to nanoparticles, and the interactions of the latter with neoplastic cells and tissues. This is probably themost promisingand encouragingapplication of nanoparticles and by far themost advanced in the process of development into a viable commercial pharmaceutical product. G. Redziniak and P. Perrier have contributed a chapter on the cosmetic applications of liposomes that have beensuccessfully exploited over the last decade. To complete thewhole rangeof applications of capsular products, a final chapter, by M. Seiller, M.-C. Martini, and myself, discusses cosmetic uses of vesicular particulate-delivery systems. Cosmetics are definitely the largest mar-

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ket, asmanufacturers have demonstrated that marketedcosmetic products containing these vesicular carriers and tested by dermatologists improve cutaneous hydration and skin texture, increase skin glow, and decrease wrinkle depth. It is now taken forgranted that liposomes and othervesicular carriers represent a major step in cosmetics formulations. However, this field requires numerous research studies coupled with strict controls. It is my hope that the scientific information contained herein will modestly contribute to a better understanding of the various particulate systems of all sizes that are now available and to an improved comprehension of their current andpotential applications.

Simon Benita

Contents Preface Contributors

...

111

xiii

Part I: Methods of Encapsulation and Advances in Production Technology 1. A Survey of Microencapsulation Processes Curt Thies

2. Microencapsulation of Oil-in-Water Emulsions by Proteins Shlomo Magdassi and Yelena Vinetsky 3. Biodegradable Microspheres: Advances in

Production Technology Jean-Pierre Benoit, Hervt Marchais, Hervt Rolland, and Vincent Vande Velde

4.

Advances in the Technology of Controlled-Release Pesticide Formulations Arie Markus

5. The Use of Factorial Design in the Development of Nanoparticulate Dosage Forms Baruch Magenheim, Pascal WehrlE, and Simon Benita

21

35

73

93

Part I I: Evaluation and Characterizationof Micro- and Nanoparticulate Drug Delivery Systems 6. Microsphere Morphology Jean-Pierre Benoit and Curt Thies

133 iX

Contents Drug Release from Microparticulate Systems

155

Clive Washington Nanoparticles: Preparation and Characterization

183

Patrick Couvreur, Guy Couarraze, Jean-Philippe Devissaguet, and Francis Puisieux Biodegradable Colloidal Drug Carrier Systems Based on Solid Lipids

213

Kirsten Westesen and Britta Siekmann Pharmaceutical Aspectsof Liposomes: Perspectives in, and Integration of, Academic and IndustrialResearch & Development

259

Rimona Margalit and Noga Yerushalmi Stealth Liposomes

297

Danilo D. Lasic Nonionic Surfactant Vesicles (NSVs) Containing Estradiol for Topical Application Don A. van Hal, Joke A. Bouwstra, and Hans E. Junginger

329

Part IIk Applications of Particulate Delivery Systems 13.

Microcapsules: Preparation by Interfacial Polymerization and Interfacial Complexation and Their Applications Tony L. Whateley

14.

Lipospheres for ControlledDelivery of Substances Abraham J. Domb, Lev Bergelson, and Shimon Amselem

15.

Pharmaceutical Emulsions, Double Emulsions and Microemulsions

349

377

411

Nissim Garti and Abraham Aserin 16. The Use of Drug-Loaded Nanoparticles in Cancer Chemotherapy

Jean-Christophe Leroux, Eric Doelker, and Robert Gurny

535

Contents

Xi

17. CosmeticApplications of Liposomes G6rard Redziniak and Pierre Perrier

577

18. Cosmetic Applications of Vesicular Delivery Systems Monique Seiller, Mane-Claude Martini, and Simon Benita

587

Index

633

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Contributors Shimon Amselem, Ph.D. Director of Pharmaceutics, Department of Pharmaceutics, Pharmos Ltd., Weizmann Industrial Park, Rehovot, Israel Abraham Aserin, Ph.D. Casali Institute of Applied Chemistry, School of Applied Science and Technology, The Hebrew University of Jerusalem, Jerusalem, Israel SimonBenita,Ph.D.* Professor, Department of Pharmacy,School of Pharmacy, The Hebrew University of Jerusalem, Jerusalem, Israel Jean-PierreBenoit,Ph.D. Professor, Laboratoire de Pharmacie GalCnique et Biophysique Pharmaceutique, UniversitC d’Angers, and Centre de Microencapsulation, Angers, France Lev Bergelson, Ph.D. Professor, Department of Pharmaceutical Chemistry, School of Pharmacy, Faculty of Medicine, The Hebrew University of Jerusalem, Jerusalem, Israel Joke A. Bouwstra, Ph.D. Associate Professor, Department of Pharmaceutical Technology, LeidedAmsterdam Center for Drug Research, Leiden University, Leiden, The Netherlands GuyCouarraze,Ph.D. Professor, Centred’EtudesPharmaceutiques, URA CNRS 1218, Universitt de Paris-Sud, Chbtenay-Malabry, France PatrickCouvreur,Ph.D. Professor, Centred’EtudesPharmaceutiques, URA CNRS 1218, UniversitCde Paris-Sud, Chitenay-Malabry, France

*Affiliated with the David Bloom Centerfor Pharmacy, Jerusalem, Israel.

xiii

xiv

Contributors

Jean-PhilippeDevissaguet,Ph.D. Professor, Centred'EtudesPharmaceutiques, UR4 CNRS 1218, Universite de Paris-Sud, Chtltenay-Malaby, France EricDoelker,Ph.D. Professor, School of Pharmacy,University of Geneva, Geneva, Switzerland Abraham J. Domb,Ph.D.* Professor, Department of Pharmaceutical Chemistry, School of Pharmacy, Faculty of Medicine, The HebrewUniversity of Jerusalem, Jerusalem, Israel Nissim Garti,Ph.D. Professor, Casali Institute of Applied Chemistry, School of Applied Science and Technology, The Hebrew University of Jerusalem, Jerusalem, Israel Robert Gurny,Ph.D. Head Professor, Department of Pharmaceutics and Biopharmaceutics, School of Pharmacy, University of Geneva, Geneva, Switzerland Hans E.Junginger, Ph.D. Professor, Pharmaceutical Deparhnent, Leided Amsterdam Center for Drug Research, Leiden University, Leiden, The Netherlands Danilo D. Lasic, Ph.D.** Department of Research, Liposome Technology, Inc. ,Menlo Park,California Jean-ChristopheLeroux,Ph.D. neva, Geneva, Switzerland

School of Pharmacy, University of Ge-

ShlomoMagdassi,Ph.D. SeniorLecturer, Casali Institute of Applied Chemistry, School of Applied Science and Technology, The Hebrew University of Jerusalem, Jerusalem, Israel BaruchMagenheim,M.Sc.+ Department of Pharmacy, School of Pharmacy, The Hebrew University of Jerusalem, Jerusalem, Israel Hem6 Marchais, Ph.D. Laboratoire de Pharmacie GalCnique et Biophysique Pharmaceutique,UniversitC #Angers, Angers, France *Affiliated with the David Bloom Center for Pharmacy, Jerusalem, Israel. **Current affiliation: Senior Scientist, MegaBios Corporation, Burlingame, California +Current affiliation: Agis Industries Ltd., Yeruham, Israel

Contributors

xv

Rimona Margalit, Ph.D. Professor, Department of Biochemistry, Faculty of Life Sciences, Tel Aviv University, Tel Aviv,Israel ArieMarkus,Ph.D. Institute for Chemistry and Chemical Technology, The Institutes for Applied Research, Ben-GurionUniversity of the Negev, Beer-Sheva, Israel Marie-Claude Martini, Ph.D. Professor, Institut Pharmaceutiques et Biologiques, Lyon, France ,

des

Sciences

Pierre Perrier, Ph.D. Parfums Christian Dior, Saint Jean de Braye, France FrancisPuisieux,Ph.D. Head Professor, Centre d’Etudes Pharmaceutiques, URA CNRS 1218, UniversitC de Paris-Sud, Chitenay-Malabry, France G6rard Redziniak, Ph.D. Applied Research Manager, Parfums Christian Dior, Saint Jean de Braye, France Hew6 Rolland,Ph.D. France

Director, Centre de Microencapsulation,Angers,

Monique Seiller, Ph.D. Professor, Unit6 d’Enseignement et de Recherche des Sciences Pharmaceutiques, UniversitC de Caen, Caen, France Britta Siekmann, Ph.D. Associate Director, Department of Pharmaceutics, Astra Arcus AB, Sodertalje,Sweden CurtThies,Ph.D. Professor, Department of Chemical Engineering, Washington University, St. Louis, Missouri Don A. vanHal, Ph.D. LeidedAmsterdamCenter for Drug Research, Leiden University, Leiden,The Netherlands Vincent Vande Velde,Ph.D. Technical Director, Centre deMicroencapsulation, Angers, France Yelena Vietsky, M.Sc. Casali Institute of Applied Chemistry, School of Applied Science and Technology, The Hebrew University of Jerusalem, Jerusalem, Israel Clive Washington, Ph.D. Lecturer in Pharmaceutics, Department of Pharmaceutical Sciences, The University of Nottingham, Nottingham, England

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Contributors

PascalWehrl6, Ph.D. Professor, Laboratorie de Pharmacotechnie, Centre de Recherches Pharmaceutiques,UniversitC Louis Pasteur, Strasbourg, France Kirsten Westesen, Ph.D. Professor, Department of Pharmaceutical Technology, Institute of Pharmaceutics, Friedrich-Schiller University Jena, Jena, Germany Tony L. Whateley,Ph.D. Reader, Department of Pharmaceutical Sciences, University of Strathclyde, Glasgow, Scotland NogaYerushalmi,Ph.D. Department of Biochemistry, Faculty of Life Sciences, Tel Aviv University, Tel Aviv, Israel

1 A Survey o f Microencapsulation Processes Curt Thies Wizshington University, St. Louis, Missouri

I. Introduction 11. Processes for Preparing Microcapsules A.GeneralComments B. Type A Encapsulation Processes C. Type B Encapsulation Processes 111. Summary

1.

1 2 2 3 10 17

INTRODUCTION

Microcapsulesare small particles that contain an active agent or core material surrounded by a coating or shell. At present, there is no universally accepted size range that particles must have in order to be classified as microcapsules. However, many workers classify capsules smaller than 1 pm as nanocapsules andcapsules larger than 1000 pm as macrocapsules. Commercial microcapsules typicallyhave a diameter between3 and 800 pm and contain 10-90 wt. percent core. A wide range of core materials has been encapsulated, including adhesives, agrochemicals, live cells, active enzymes, flavors, fragrances, pharmaceuticals, and inks. Most capsule shell materials are organic polymers, but fats and waxes are also used. 1

Thies

2 Shell

Material

A

B

Fig. 1 Schematicdiagrams of tworepresentativetypes of microcapsules: (A) Continuous core/shell microcapsule. (B) Multinuclear microcapsule. (Courtesy of C. Thies.) Microcapsules can havea variety of structures. Some havea spherical geometry with a continuous coreregion surrounded by a continuous shell, as shown in Fig. 1A. Others have an irregular geometry and contain a number of small droplets or particles of core material, as shown in Fig.1B. Microcapsules are usedin a wide range of commercial products. They are thekey component of all carbonless copy papers and are present in several oraland injected drug formulations. Encapsulated adhesive resins coated on automotive fasteners are routinely used to assure that such fasteners are firmly set when installed. Microcapsules also are the basis for a number of long-actingcommercial pesticide and herbicide products. Improvement of these products and developmentof new ones is an ongoing process that involves a large number of development groups globally.

II.

PROCESSES FOR PREPARING MICROCAPSULES

A.GeneralComments

The concept of preparing small particles that carry a core material trapped within a shell material dates back at least to spray-drying work carried out in the 1930s. However, the first truly significant commercial productthat utilized such particles was carbonless copy paper. The microcapsules for this product were made by a process called complex coacervation. Since then many other methods forpreparing microcapsules have been developed and improved significantly. Some are based exclusivelyon physical phenomena. Some utilize polymerization reactions to produce a capsule shell. Others combine physical and chemical phenomena. Many investigators classify en-

Microencapsulation ofA Processes Survey

3

capsulation processes as either chemical or mechanical. I prefer toclassify them as type A or type B processes, since so-called mechanical processes may actually involve a chemical reaction and so-called chemical processes may rely exclusively on physical phenomena. Table 1 lists representative examples of both types of processes. Capsules produced by type A, or chemical processes, are formed entirely in a liquid-filled stirred tank or tubular reactor. Capsules produced by type B, or mechanical processes, utilize a gas phase at some stage of the encapsulation process. In such processes, capsules can be formed by either spraying droplets of coating material on acorematerial being encapsulated, solidifying liquid droplets sprayed into a gas phase, gelling droplets sprayed into a liquid bath, or carrying out apolymerization reaction at a solid-gas interface. No encapsulation process developed to dateis able to produce the full range of capsules desired by potential capsule users. Some readily produce small, liquid-filled capsules, whereas others produce relatively large capsules with a solid core material. In order to obtain a feeling for which process is most appropriate for a specific application, it is appropriate to summarize the general principles of several types A and B processes. This chapter focuses primarily on processes that have achieved significant commercial use. Several reviews giveadditional details[l-61. B. Type A Encapsulation Processes

l . Complex Coacervation Because of its original use in carbonless paper, the first type A process considered is complex coacervation. It is based on the ability of cat-

Table 1 List of Types A and B Encapsulation Processes Type A (chemical) processes

Type B (mechanical) processes

Complex coacervation Polymer-polymer incompatibility Interfacial polymerization in liquid media In situ polymerization In-liquid drying

Spray drying Spray chilling Fluidized bed Electrostatic deposition Centrifugal extrusion Spinning diskor rotational suspension separation Polymerization at liquid-gas or solid-gas interface Pressure extrusion or spraying into solvent extraction bath

Thermal andionic gelation in liquid media Desolvation in liquid media

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4

ionic and anionic water-soluble polymers to interact in water to form a liquid, polymer-rich phase called a complex coacervate. Gelatinis normally the cationicpolymer used. A variety of natural and synthetic anionic watersoluble polymers interact with gelatin to form complexcoacervates suitable for encapsulation.When the complex coacervate forms,it is in equilibrium with a dilute solution called the supernatant. Inthis two-phase system, the supernatant acts as the continuous phase,whereas the complex coacervate acts as the dispersed phase. If a water-insoluble core material is dispersed in the system and the complex coacervate wets this core material, each droplet or particle of dispersed core materialis spontaneously coated with a thin film of coacervate. When this liquid film is solidified, capsules are formed. Fig. 2 outlines one version of a complex coacervation encapsulation process [7]. The first step is to disperse the core material in qn aqueous gelatin solution. This is normally done at40-60°C, a temperature range at which the gelatin solution is melted and liquid. After a polyanion or negatively charged polymer like gum arabic is added to thesystem, the pH and

tielalln

4

Hyoo (40-60C)

H

Form Emulsion of Core In Solulion Gelalln (40-60°C)

1 Gel Shell (515°C)

l

Core Malerial (Llquicl)

Form Liquid Coacervale by Adjusling pH I Adding Polyanion and Dilulion Waler (40-60'C)

7Crosslink with Glutaraldehyde

Fig. 2 Schematic flow diagram of encapsulation process based on complex coacervation. (Courtesy of C. Thies.)

A Survey of Microencapsulation Processes

5

concentration of polymer are adjusted so that a liquid complex coacervate forms. The pH atwhich this occurs is typically between 4.0 and 4.5. Once the liquid coacervate forms, the system is cooled to room temperature. The gelatin in the coacervate gels thereby forming capsules with a very rubbery shell. In order to increase the strength of the water-swollen shell and create a gel structure that is not thermally reversible, the capsules normally are further cooled to approximately 10°C and treatedwith glutaraldehyde. The glutaraldehyde crosslinks the gelatin by reacting with amino groupslocated on the gelatin chain. Glutaraldehyde-treated capsules can be dried to a free-flow powder or coated on a substrate and dried. Complex coacervation has been used to encapsulate many waterimmiscible liquids and is used in a variety of products, including inks for carbonless paper, perfumes for advertising inserts, and liquid crystals for display devices. This technology routinely produces single capsules of 20800 pm diameter that contain 80-90 wt. percent core material. Capsules outside this size range can be made, but considerable know-how may be required. Most coacervate capsules have a continuous core/shell structure (see Fig. lA),although the shell is not of uniform thickness. The mechanical and barrier propertiesof dry capsules formed by complex coacervation generally are sensitive to moisture. When stored at humidities above 70% or immersed in water, the shell of a complex coacervate capsule swells greatly, thereby facilitating transport of material into or outof the capsule. Postreatment of such capsules with urea and formaldehyde under acidic conditions has been used to reduce their moisture sensitivity. The value of complex coacervation encapsulation technology for food and pharmaceutical applications would be greatly enhanced if such capsules could be isolated commercially without the use of a chemical crosslinking agent.

2. Polymer-PolymerIncompatibility Polymer-polymer incompatibility is a second type A process that has been commercialized. This technology utilizes a polymer phase-separation phenomenon quitedifferent from complex coacervation. In complex coacervation, two oppositely charged polymers, gelatin and a polyanion, join together to form the complex coacervate and both polymers become part of the final capsule shell. In contrast,polymer-polymer incompatibility occurs because two chemically different polymers dissolved in a common solvent are incompatible and do not mix in solution. They essentially repel each other and form two distinct liquid phases. One phase is rich in polymer designed to act as the capsule shell. The otheris richin the second, incompatible polymer. The incompatible polymer is present in the system to cause formation of two phases. It is not designed to be part of the final capsule shell, although small amounts may remain entrapped in the final capsule as

Thies

6

% l

0000

@ V

Phase Separated Solution of Ethylcellulose+ Polyethylene in Hot (80'C) Cyclohexane

Core Material (Small Particles)

t

Dispersion of Core Material in Hot, Phase-Separated Polymer Solution

+ Cool to 20-c (Ethylcellulose Coating Solidifies)

Elhylmllulose Shell

Wash and Harvest Capsules

Fig. 3 Schematic flow diagram of encapsulation process based on polymerpolymer phase separation. (Courtesy of C. Thies.)

an impurity. This type of encapsulation process generally does not involve any chemicalreaction. Fig. 3 illustrates a specific polymer-polymerphase-separation encapsulation process used to produce commercial pharmaceutical grade microcapsules [8]. The first step is to disperse the core material in ahot (80°C) solution of ethyl cellulose in cyclohexane. Lowmolecular weight polyethylene, a polymer soluble in hot cyclohexane and incompatible with ethyl cellulose, is added to thesystem. This induces phase separationwith formation of an ethyl cellulose-rich phase and a polyethylene-rich phase. The

A Survey of Microencapsulation Processes

7

core material, a solid unaffected by 80°C cyclohexane, is dispersed in this two-phase system. Since the ethyl cellulose is more polar than polyethylene, it adsorbs preferentially on the surface of the core material and thereby causes a thin coating of shell material solution to engulf the particles of core material. When the system iscooled to room temperature, the ethyl cellulose precipitates, thereby solidifying the ethyl cellulose solution and formingsolid microcapsules that can beharvested. Aspirin and potassium chloride are examples of commercial encapsulated pharmaceutical products producedin this manner [g]. Polymer-polymer phase-separation processes normally are carried out in organic solvents and are used to encapsulate solids with a finite degree of water solubility. Many of the capsules produced commercially have an ethyl cellulose shell, are irregularly shaped 200- to 800-pm particles, and are loaded with solid drug particles. They are used to provide taste masking or prolonged oral drug delivery [g, 101. Smaller capsules (e.g., 20-120 pm in diameter) with a biodegradable poly(d,l-lactideglycolide) shell have been preparedby polymer-polymer phase separation and used asinjectable drug-delivery devices [ll].

3. Interfacial Polymerization Interfacial polymerization (IFP) is a third type A encapsulation process that has been commercialized. A unique featureof this technology is that thecapsule shell is formed at or on the surface of a droplet orparticle by polymerization of reactive monomers. This approach to encapsulation has evolved into a versatile technology able to encapsulate a wide range of core materials, including aqueous solutions, water-immiscible liquids, and solids. Many types of IFP reactions are used. Fig. 4 is a schematic diagram that illustrates how the capsule shell forms when a capsule loaded with a water-immiscible liquid is prepared by IFP. In suchcases, a multifunctional monomer is dissolved in the liquid core material. This monomer often is a multifunctional isocyanate but could be a multifunctional acid chloride or a combination of isocyanates and acid chlorides. In all cases, the resulting solution is dispersed to a desired drop size in an aqueous phase that contains a dispersing agent. A coreactant, usually a multifunctional amine, is then added to the aqueous phase. Thisproduces a rapid polymerization reaction at the interface which generates the capsule shell[12]. A polyurea capsule shellis obtained if the IFP reaction involves an isocyanate and an amine. A polyamide or nylon capsule shell is obtained if the IFP reaction involves an acid chloride and an amine. Reaction of an isocyanate with a hydroxylcontaining monomer dissolved in the aqueous phase produces a polyurethane capsule shell.

Thies

8 Polyurea Polyurea Shell

-

Core Material ["oil")

'. H20

+

EMULSION STABILIZER

Fig. 4 Schematic illustrationof an interfacial reaction frequently usedto prepare microcapsules by interfacial polymerization. (Courtesyof C. Thies.)

If the core material being encapsulated is an aqueous solution, the abovereactantadditionsequence is reversed.Thecoreactant,e.g.,a water-soluble amine,is dissolved in the aqueous phase.The resulting mixture is dispersed in a water-immiscible solvent.unti1the desired drop sizeis obtained. A solvent-soluble reactant like an isocyanate or acid chloride is added to the organic phase, thereby initiating rapid polymerization at the interface to produce a capsule shell. A variety of core materialsof biological interest (e.g., active enzymes) have been producedin this manner [13]. Solids can also be encapsulated by interfacial polymerization reactions, although the polymerization chemistry typically differs from that used to encapsulate liquids. Vinyl monomers that polymerize by free radical reactions generally are used to encapsulate solids. In one process, the solid to be encapsulatedis dispersed in aqueousmedia along with a dispersing agent. Vinyl monomer is added to thesystem and freeradical polymerization is initiated by a redox initiation system [14]. Success of this type of encapsulation process depends on forcing polymer formation to occur at the solid-liquid interface rather than allowing it to occur throughout the entire aqueous media. Another type of interfacial polymerization reaction involves the polymerization of p-xylylene at a gas-solid interface [15]. In this case,polymerization of vaporized p-xylylene radicals occurs spontaneously on contactwith a solid surface. Although interfacial polymerization can be used to produce large capsules, most commercial uses of this technology have focused on the preparation of small capsules that are coated or sprayed onto a substrate rather thanbeing isolated as afree-flow powder. For example, IFP is routinely used to produce 20- to 30-pm diameter

icroencapsulation ofA Processes Survey

9

capsules loaded with pesticides and herbicides. It also is used toform 3-to 6-pm diameter capsules loaded with carbonless paper ink. Capsules with polyurea or polyamide shells have liquid cores or cores that melt below 60-70°C. Because one of the reactantsused to create thecapsule shell is dissolved in the core material and is free to react with any reactive groups located on molecules of core material to create new molecular species, IFP encapsulation technology is viewed suspiciously by workers concerned with the encapsulation of fragrances, drugs, and flavors. Many agrochemicals like pesticides and herbicides are relatively water-insoluble low-melting solids or high-boiling point liquids free of groups that could react with the reagent(s) dissolved in the core material. Thus, many current commercial encapsulated agrochemical formulations are prepared by IFP [12]. Capsules formed by IFP often have a continuous core-shell structure and a spherical geometry. Significantly, the exterior surface of many IFP capsules is smooth and uniform. This is not the case for the interior surface. Capsule fracture studies show that the interior surface of many IFF' capsule shells is cratered and irregular. That is, the capsule shell is not uniformly deposited around the core by an IFP process except for a comparatively thin outer region of the shell. In such cases, the shell of an IFP capsule is not a membrane of uniform thickness. It is a thin skin membrane resting on a thick, porous support. In Situ Polymerization In situ polymerization is a type A encapsulation technology closely related to IFP. Like IFP, capsule shell formation occurs because of polymerization of monomers added to theencapsulation reactor. However, with in situ encapsulation processes, no reactive agents are added to thecore material. Polymerization occurs exclusively in the continuous phase and on the continuous-phase side of the interface formed by the dispersed core material and continuous phase. Polymerization of reagents located there producesa relatively low molecular weight prepolymer. As this prepolymer grows in size, it deposits onto thesurface of the dispersed core materialbeing encapsulated where polymerization with crosslinking continues to occur thereby generating a solid capsule shell. The first example of this technology was the encapsulation of various water-immiscibleliquids with shells formed by the reaction at acid pH of urea with formaldehyde in aqueous media [14]. Similar encapsulation reactions occur between melamine and formaldehyde in aqueous media. Addition of anionic polymers to theaqueous phasecan have a significant effect on thein situ encapsulation process [16]. In situ polymerization is used extensively to produce small(3- to 6-pm diameter) capsules loaded with carbonless paper inks or perfume for scented strips. Larger capsules loaded with mineral oil are used for cos-

4.

.

10

Thies

metic applications, whereas capsules filled with epoxy resin are used for fasteners. In all of these cases, the capsules have a continuous core-shell structure. Thetechnology can also be adapted to encapsulatesolids.

5. Centrifugal Force and Submerged Nozzle Processes Several interesting type A processes use centrifugal force or two-fluid submerged nozzles to form microcapsules. In oneprocess, a cup perforated with a series of fine holes is immersed in an oil bath. It is rotated while immersed in the oil, thereby extruding into the oil phase a stream of droplets of an oil-in-water emulsion [17]. The water phase of this emulsion is a concentrated solution of a water-soluble polymer that gels on cooling. Gelatin is a specific example. By controlling the temperature of the oil bath, the external phase of the extruded emulsion droplets is gelled to create oil-loaded gel beadlets that can be isolated and dried. When isolated, the capsules consist of a number of small oil droplets dispersed throughout a matrix of shell material. This process was developed in 1942 in order to produce capsules that improved the oxidative stability of vitamins and fish oils [17]. Capsules can also be produced by coextruding an aqueous gelatin solution and an oil to be encapsulated through a two-fluid nozzle into a moving fluidstream of an oil solution. Fig. 5 schematically illustrates such a nozzle. The gelatin solution surrounds the oil drop to be encapsulated at the moment of extrusion and is gelled thermally before the particles are harvested and dried. In the case shown in Fig. 5 , continuous shell-core capsules are obtained.

C. Type B EncapsulationProcesses Centrifugal force, extrusion,coextrusion, and formation of sprays are the principal means by which type B capsules are made. Type B methods of encapsulation predate type A encapsulation processes, since spray drying, a type B process, was developed in the 1930s. Because spray drying is the oldest type B encapsulation process, it is appropriate to discuss it first.

1. SprayDrying The first step in a spray-dry encapsulation processes is to emulsify or disperse the core material in a concentrated (40-60 wt. percent solids) solution of shell material.The core materialgenerally is a water-immiscible oil such as a fragrance, flavor, or vitamin. It is emulsified in a solution of shell material until 1-to 3-pmoil droplets are obtained.The shell material normally is a water-soluble polymer like gum arabic or a modified starch. These materialsdo not form high-viscosity solutions at thehigh-shell material concentrations favored for spray drying. Mixtures of these shell materi-

A Survey of Microencapsulation Processes

-

Core Material (e.g., Vitamin O i l )

Shell Material (e.g.. Warm Gelatin Solution)

+ 2-Fluid2-Fluid. Nozzle

Warm Carrier Phase (e.g.,Vegetable Oil)

@

l =l

Coil

@

z

@ /

Gel Beads that Core can be harvested Material and dried

Fig. 5 Schematicdiagram of a submergedtwo-fluidnozzleused microcapsules. (Courtesyof C. Thies.)

to prepare

als with hydrolyzed starches (maltodextrins) or hydrolyzed gelatins have been used. Water is the preferred solvent for most spray-drying encapsulations. Concerns about solvent flammability and toxicity severely restrict the use of organic solvents for conventional spray-drying encapsulation applications. However, several groups are exploring the use of spray drying from organic media to produce pharmaceutical capsules from biodegradable polymers. Once a suitable dispersion of core material in a solution of shell material has been prepared, the resulting emulsion is fed as droplets into the heated chamberof a spray drier. The droplets may be sprayed intothis chamber or spunoff a rotating disk. In either case, they are rapidly dehy-

72

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drated in a heated chamber, thereby producing dry capsules. The capsules fall to the bottom of the spray-drying chamber where they are harvested. Capsules produced in this manner typically fall between 10 and 300 pm in diameter. They tend to have an irregular geometry and may be aggregates of a number of small particles. Each spray-dried capsule has a number of small droplets of core material dispersed throughout it. Spray-dry encapsulation has a number of advantages. It is a wellestablished technology, involves readily available equipment, andis able to produce large amounts of capsules. Many shell materials preferred for spray drying are approved for food use. Furthermore, these materials are water soluble and notchemically crosslinked. Thus, capsules prepared from them dissolve in water andrelease core material without leaving any capsule shell debris. In contrast, the shell of complex coacervate capsules crosslinked with glutaraldehyde swells greatly in water but does notdissolve. Such capsules can be ruptured, therebyreleasing their contents, butcapsule shell debris is left behind. For some applications, this is objectionable. Spray-dry encapsulation has several problems and limitations. For example, if water is the preferred solvent, spray-dry encapsulation is limited to shell materials soluble or dispersible in water. The list of candidate water-soluble shell materials is limited, because many candidate materials form aqueous solutions that are tooviscous to spray in spray-dry encapsulation protocols. Another limitation is the 20-30% core loading carried by most spray-dried capsules. Spray-drying protocols that allow core loading up to 50-60% have been reported [18], but current spray-dried capsules have lower loadings. A persistent problem with spray-dried capsules is free or surface core material. Because water evaporation from a capsule in the chamber of a spray drier occurs rapidly, it is not uncommon to harvest spray-dried capsules that have a finite amount of free or unencapsulated oil. The higher the core loading, the more pronounced this problem can become. This is undesirable, since free fragrance or flavor oil is susceptible to oxidation and the development of an off-odor or off-taste. Brenner [l81 noted that maximizing core loading and minimizing free core content involves a judicious choice of coating material, emulsifying agent, andspraydryeroperating conditions. Finally, it is important to stress that lowboiling point compounds with a finite degree of water solubility have posed a persistent problem to spray-dry encapsulation. Such compounds volatilize from the capsules in the spray-drying chamber. In summary, spray drying is a viable commercial method of forming microcapsules. It is an established, comparatively low-cost encapsulation technology that continues to develop. To date, spray drying has primarily been used to encapsulate fragrances and flavors.

A Survey of Microencapsulation Processes

13

2. Fluidized Bed Coaters Fluidized bed coaters are another type B encapsulation technology. They are limited to encapsulating solid particles or porous particles into which a liquid has been absorbed. Nevertheless, they are used extensively to encapsulate many different solids. Fluidized bed coaters function by suspending a bed or column of solid particles in a moving gas stream, usually air. A liquid coating formulation is sprayed onto the individual particles, and the freshly coated particles are cycled into a zone where the coating formulation is dried either by solvent evaporation or cooling. This coating and drying sequence is repeated until a desired coating thickness has been applied. Many variables affect the fluidized coating process, including dew point of the incoming air. Lehmann [l91 reviewed the operation of fluidized bed coaters. They represent a major capital investment, but are widely usedto produce encapsulated solids, especially for the pharmaceutical industry. A major advantage of fluidized bed coaters is their ability to handle an extremely wide range of coating formulations. They have been used to apply hot melts, aqueous latex dispersions, organic solvent solutions of shell material, and aqueous solutions of shell material. Enteric polymer solutions have been of particular interest, since they are used by the pharmaceutical industry to produce drug-dosage formulations that survive passage through the stomach. Enteric polymers are insoluble in gastric fluid (pH 2) and soluble in intestinal fluid (pH 7), so capsules or tablets coated with them can pass intact through the stomach and not disintegrate until they reach the intestine. Three types of fluidized beds are available: top-spray, tangentialspray, and bottom-spray. These units differ in location of the nozzle or nozzles used to apply the coating formulation. Fig. 6 offers schematic diagrams of a top-spray and a bottom-spray unit. In the top-spray unit (Fig. 6A), the coating formulation is sprayed into the fluidized bed. The droplets of spray leaving the nozzle move countercurrent to the gas stream until they impact the particles being coated. If the spray formulation contains a volatile solvent, evaporation of this solvent from the spray droplets occurs, thereby increasing the solids content, perhaps to such a degree that the spray droplets cannot spread on the particles being coated. This spraydrying effect has been used to explain why solvent-based coating formulations applied in top-spray fluidized bed coaters oftenyield coated particles with a degree of internal void volume and porous coatings. Enteric coatings applied as aqueous latex dispersions are an exception. Such coatings applied in a top-spray fluidized bed coater form a continuous coating analogous to that obtained with tangential-spray and bottom-spray units. Fats

Thies

14

A

Being Coated

Gas Flow

Expansion Chamber

Core Material Being Coated Formulation Gas Flow

Fig. 6 Schematic diagrams of two types of fludized bed coaters: (A) Top-spray unit. (B) Bottom-spray, or Wurster unit. (Courtesy of C. Thies.) are also applied by top-spray fluidized bed coaters. Top-spray coaters have limitations, but they are more simple to operate and have higher production capacity than the othertypes of units. In tangential-spray and bottom-spray units, the droplets of coating formulation move in the same direction as the particles being coated. The droplets of coating formulation travel a shorterdistance before theyimpact

A Survey of Microencapsulation Processes

15

the particles being coated. Solvent evaporation is minimized, and a more uniform film of coating material is deposited. The bottom-spray, or Wurster coater, shown in Fig.6B has becomea standard unit used to produce encapsulated solids, especially solidpharmaceuticals. The partition and gas-distribution plates of such units are important components, becausethey direct the particles being coated in a cyclic path past the nozzle that supplies the coating formulation. Particles move past this nozzle, receive a spray of coating material, and move up into the upper section of the unit. Here the coating is solidified either by solvent evaporation or cooling of a hot melt. The coated particles then fall back down to thebottom of the unit where a fresh coating is applied. This cyclic process is repeated until the desired weight of shell material has been applied. Application of the coating in a series of cyclic steps is both an advantage anda disadvantage. The disadvantage is that stepwise deposition takes time. A one-step coating process is much quicker, but with a multistep coating process, defects can beminimized or perhaps eveneliminated. This makes it possible to use the Wurster process to produce capsules with a shell that approaches the samebamer properties as a free-standing solvent-cast film. In a one-step coating process, capsule shell defects are not healed and, hence, can havea profound effect on capsule release behavior.

3. Centrifugal Extrusion Fig. 7 is a schematic diagram of a centrifugal extrusion process [20]. The core and shell material, two mutually immiscible liquids, are pumped through a spinning two-fluid nozzle. This produces a continuous two-fluid column or rod of liquid that spontaneously breaks up into a stream of spherical droplets immediately after it emerges fromthe nozzle. Each droplet contains a continuous core region surrounded by a liquid shell. How these droplets are convertedinto capsules is determined by the nature of the shell material. If the shell material is a relatively low-viscosity hot melt that crystallizes rapidly on cooling (e.g., a wax or wax toughened with a polymer), the droplets are converted into solid particles as they fall away from the nozzle. Suitable core materials typically are polar liquids like water or aqueous solutions, since they are immiscible with a range of hot melt shell materials like waxes. Alternatively, droplets emerging from the nozzle may have a shell that is an aqueous polymer solution able to be gelled rapidly. In this case, the droplets fall into a gelling bath wherethey are converted intogel beads. A specific example is the gelatin of an aqueous sodiumalginate shell by an aqueous CaCl, gelling bath. The gel beads produced can be dried to give capsules with a solid shell. Core materials suitable for this type of capsules shell are water-immiscible oils.

Thies

16 Core Material Shell

Shell

AIR ,Liquid Core

Shell gelled b y immersion in gelling bath

Shell solidifies

by cooling in air

Fig. 7 Schematicdiagram of centrifugaltwo-fluidnozzleused crocapsules. (Courtesyof C. Thies.)

to

produce rni-

Capsules prepared by centrifugal extrusion tend to be large with diameters typically ranging from over 250 p up to several millimeters. 4. Rotational Suspension Separation Fig. 8 is a schematic diagram of the rotational suspension separation process for preparing microcapsules. In this process, core material dispersed in a liquid shell formulation is fed onto a rotating disk. A flat disk is shown, but conical or bowl-shaped disks can be used. Individual core particles coated with a film of shell formulation are flung off the edge of the rotating disk along with droplets of pure coating material. When the shell

A Survey of Microencapsulation Processes "Spherical" Material

Coating Core Phase (e.g.. molten lot)

!

17

Rotate

Spinning Disc

0 h

t

Free Coating Material

Fig. 8 Schematicdiagram

t

Capsules

of rotational suspension separation encapsulation

process.

formulation is solidified, e.g., by cooling, discrete microcapsules are produced. The droplets of pure coating material also solidify, but they are said to collect in a discrete zone away from the capsules, as shown in Fig. 8. This technology is claimed to be a fast, low-cost, high-volume method of encapsulating a variety of materials that act like solids on the rotating disk, including particles below 150 pm in diameter [21]. In order toobtain optimal results, the core material must have a spherical geometry. Formation of this geometry may require a granulation step before encapsulation is attempted. A variety of hot-melt shell materials can be applied, butmelt viscosities below 5000 CP are favored. Capsule shell formulations that do not solidify rapidly are said to pose problems. Since solidification of the coating formmulation occurs rapidly and the coating formulations applied are crystalline materials susceptible to polymorphic transformations or changes in percent crystallinity on storage, it is reasonable to suggest that capsule shells produced by rotational suspension separation are notat thermodynamic equilibrium immediately after capsule formation is complete. 111.

SUMMARY

Most microcapsules of commercial significance are produced by the encapsulation processes described in this chapter. The commercialization of products that contain microcapsules is progressing as is the development of encapsulation processes. Today, a number of capsule-based products exist.

Thies

18

Several have significant sales. The biggest application, carbonless paper, consumes thousands of tons of capsules per year [5].Such capsules are produced by complex coacervation, IFP, or in situ polymerization. Large volumes of commercial capsules loaded with agrochemicals, especially herbicides, are madeby IFF. Although a numberof encapsulated food ingredients arebeing marketed [22,23],the volume of such capsules remains small relative to the huge potential that exists. The development of encapsulation technology and microcapsule-based products continues tobe pursued actively by a number of groups globally. Accordingly, it is reasonable to project that this field will experience steady growth for the foreseeable future. It will be interesting to see when the next high-volume capsulebased product appears and what it will be.

REFERENCES 1. P. B. Deasy, Microencapsulation and Related Drug Processes, Marcel Dekker, New York, 1984. 2. A. Kondo, Microcapsule Processing and Technology, Marcel Dekker, New York, 1979. 3. C. Thies, Kirk-Othmer Encyclopedia of Chemical Technology, Vol. 16, 4th ed., Wiley, 1995, pp. 628-651. 4. W. Slwika, Angew. Chem. Internat Edit., 14539-550 (1975). 5. C. A. Finch, Ullmann’s Encyclopedia of Industrial Chemistry, Vol. A16, 5th ed., VCH Publishers, New York, 1990, pp. 575-588. 6. I. C. Jacobs, and N. S. Mason, in Polymeric Delivery Systems, ACS Sympo-

sium Series520 (M. A. El-Nokaly, D. M. Piatt, and B. A. Charpentier,eds.), American Chemical Society, Washington, DC,1993, pp. 1-17. 7. B. K. Green, and L. Schleicher,U.S. Patent 2,800,457,1957. 8. J. L. Anderson, G. L. Gardner, and N. H. Yoshida, U.S. Patent 3,341,416, 9.

1967.

J. A. Bakan, in The Theory and Practice of Industrial Pharmacy, 2nd ed. (L. Lachman, H. A. Lieberman, and J. L. Kanig, eds.), Marcel Dekker, New York, 1986. 10. C. Thies, CRC Crit. Rev. Biomed. Eng.,1983,8:335-383 (1983). 1 1 . B. J. Floy, G . C. Visor, and L. M. Sanders in Polymeric Delivery Systems, ACSSymposiumSeries 520 (M. A. El-Nokaly, D. M. Piatt,and B. A. Charpentier, eds.), American Chemical Society, Washington, DC,1993, pp.

154-167. 12. H. B. Scher, in Pesticide Chemistry-Human Welfare and the Environment, Vol. 4 (J. Miyamoto and P. C. Kearny, eds.), Pergamon Press, Oxford, UK, 1983, pp. 295-300. 13. T. M. S. Chang, Artificial Kidney, Artificial Liver, and Artificial Cells, Plenum Press, New York,1978.

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14. M. Cakhshaee, R. A. Pethrick, H. Rashid, and D. C. Shemngton, Polymer Commun., 26:185-192 (1985). E. 15. W. D.Jayne,inMicroencapsulation:ProcessesandApplications(J. Vandegaer, ed.), Plenum Press, New York, 1974, p. 103. 16. K. Dietrich, H. Herma, R. Nastke, E. Bonatz, and W. Tiege, Acta Polymerica, 40:243-251 (1989). 17. J. A. Reynolds, U.S.Patent 2,299,929, 1942. 8:40-44 (1983). 18. J. Brenner, Perfumer and Flavorist, 19. K.Lehmann, in Microcapsules and Nanoparticles in Medicine and Pharmacy (M. Donbrow, ed.), CRC Press, Boca Raton, FL, 1992, pp. 73-97. 20. J. T. Goodwin, and G . R. Somerville, Chemtech, 4:623 (1974). 21. R. E. Sparks, I. C. Jacobs, andN. S. Mason, in Polymeric Delivery Systems, ACSSymposiumSeries520(M. A. El-Nokaly,D.M. Piatt,andB.C. Charpentier, eds.), American Chemical Society, Washington, DC, 1993, pp. 145-153. 22. S. Hagenbart, Food Product Design, April:28-38 (1993). 23. R. Arshady, J. Microencapsulation, 10:413-436 (1993).

2 Microencapsulation of Oil-in-Water Emulsions by Proteins Shlomo Magdassi and Yelena Vinetsky Casali Institute ofApplied Chemisty, School ofApplied Science and Technology, The Hebrew University of Jerusalem, Jerusalem, Israel

Introduction I.

21

11. Methods of Encapsulation A. Phase Separation B. Spray Drying C. HeatDenaturation

23 23 25 25

1 1 1 . Surface Activity of Proteins

25

IV.

27

1.

PossibleFilm-Forming Proteins

INTRODUCTION

The formation of microcapsules is a widely used process that combines both basic chemical aspects and technological aspects. This process is usually very complex, since it involves various reactions and steps which take place atdifferent phases. For example, the initial step of a simple coacervation-based process isbased on adsorption of one of the wallforming components onto the core material-continuous phase interface. Therefore, the migration of the molecule from the bulk continuous phase to theinterface and the kinetics and reversibility of the adsorption process are of great importance.The second step is usuallybased on conversion of 21

22

and

Magdassi

Vinetsky

the adsorbed layer into a semisolid wall, with a defined and measurable thickness and permeation properties. This process involves the addition of other molecules, which lead to precipitation or coacervation onto the core material. Usually, this process is performed by adding an electrolyte (simple coacervation) or an oppositely charged polymer (complex coacervation). Since the first wall component may also be present in the bulk phase, and not only adsorbed at the core-continuous phase interface, it could also participate in the coacervation or precipitation process. It’is clear that such a situation is undesirable, as it results in waste of material, which could otherwise be used for the formation of the wall. Moreover, the free molecules in the continuous phase may also lead to aggregation of the individually dispersed particles or droplets of the core material, leading to formation of very large particles, which may sediment or float, and which are often unsuitable for most applications. The next step is usually based on hardening of the wall material by various crosslinking agents, which are added to the continuous phase. It is obvious that such a process should be carefully controlled, because, in addition to the hardening of the wall material of each microcapsule, it may also lead to crosslinking between several microcapsules, thus changing the physicochemical properties of the microcapsules. In most applications, it is desirable to obtain, a dry, microcapsules powder as a final product, which can be further redispersed in various liquid phases. Therefore, removal of the liquid continuous phaseis oftenrequiredand is achieved by a process such as spray drying or lyophylization. Such a process could often lead to agglomeration of the microcapsules, which should be prevented if a redispersion of the microcapsules is desired. These problems are often solved by including various additives in the liquid phase in a way that the removal of the liquid phase would cause the formation of an additional “barrier” between the individual microcapsules, which will disappear on redispersion. Obtaining dry, redispersible microcapsules is still not the end process. A caking phenomena that occurs as a result of agglomeration caused by the pressure gradient present in the storage containers may also be observed. This problem usually requires the use of anticaking agents, which can be added to the dry powder as well as to the final steps of removal of the liquid phase. The complexity of the process is more severeif the core material is also a liquid: In such cases, the microencapsulation process should also include an emulsificationstep in which a stable oil-in-water or water-in-oil emulsion could be obtained. The emulsification process is based on various parameters, such as interfacial tension, adsorption of the surfactant, or homogenization equipment. The use of a liquid core may also lead to manufacturing problems at the endof the process, during the removal of the continuous

Microencapsulation of Oil-inEmulsions Water

23

liquid phase, as in the case of encapsulation of volatile oils (such as orange oil), which are widely used as fragrance and flavor additives. This chapter deals with the formation of microcapsules in which the core material is a water immiscible oil and the continuous phase is an aqueous solution. The wall material described here is composed of various proteins and, in particular, gelatin. The principles for obtaining the microcapsules are similar, and therefore we will discuss in detail the various steps of the microencapsulation process with emphasis on the proteinbased microcapsules. II. METHODS OF ENCAPSULATION

Many different techniques have been proposed for the production of microcapsules, and it was suggested(1) that more than200 methods could be identified in the patent literature. Three methods which are significantly relevant to microcapsules that are based on proteins or polysaccharides, especially when the core material is an oil, are briefly discussed here. A.PhaseSeparation

As described in a publication by Finch [l], in the phase separation-based process, the core contents, in a solvent, are first suspended in a solution of the wall material. The wall polymer is induced to separate as a new, viscous, polymerrich phase by adding a nonsolvent, by lowering the temperature,by changing the pH,by adding a second polymer, or by changing other environmental conditions. It is essential that such changes would cause the polymer to come out of the solution and to aggregate around a core dropletto form a continuous encapsulating wall [2]. This process is shown schematically in Fig. 1, which includes the hardening of the coacervate layer, usually by a crosslinking agent. The coacervation of the polymer can be recognized bythe appearance of turbidity, droplets, or separation of the solution into two layers, which contain high and low concentrations of the polymer. Coacervation may be simple or complex: Simple coacervation is obtained by adding a water-miscible nonsolvent to theaqueous polymersolution (e.g., ethanol) or by adding an electrolyte, which causes polymer separation due to a “salting-out” mechanism. A typical example is the addition of sodium sulfate to an oil-in-water ( O M ) emulsion, whichis formed by using gelatin [3,4]. The system may be stabilized by altering the pH or the temperature by or addition of a crosslinking agent. Complex coacervation occurs with the mutual neutralization of two oppositely charged polymers. A widely used method is based on coacer-

Magdassi and Vinetsky

24

dissolved in water -

0

Emulsification

7I Film forming material, oppositely charged

Hardening of coacervate-capsule wall by chemical or physical means

-

V

Removal of water

Fig. 1 Schematic presentation of a formation of microcapsules of oil droplets in water by complex coacervation.

Microencapsulation of Oil-inEmulsions Water

25

vation of cationically charged gelatin (at pH below 8) by a negatively charged polymer such as gum arabic. This process was originallydeveloped for the formationof carbonless copying paper [5]. B. SprayDrying

Microencapsulationby spray drying is based on two steps: emulsification of the coreoil (or dispersion of particles) in the polymer solution followed by removal of the solvent by a hot stream of air. This method, in spite its limitations, is the most prevalent process used to encapsulate flavors [6]. The main problems relatedto this process are the nonuniformedsize of the microcapsules (due to coalescence of oil droplets at high temperature and aggregation of microcapsules) and incomplete encapsulation of the core component, which is often a volatileoil. The wall materials which can be used in sucha process can be various polysaccharides (starch, gum arabic) or proteins (gelatin, albumin, casein) [7,8]. It is obvious that while choosing the wall material, two important parameters should be taken into consideration: theability of the polymer to emulsify the oil phase and the behaviorof the polymer at high temperature. The latterhas a uniquemeaning regarding theuse of proteins, which can be denaturated specific at temperatures andhence form unsuitablewall material. C.HeatDenaturation Heat denaturation of proteins can also be used for encapsulation, as described recentlyin a patentby Janda et al. [g]. Briefly, the method is based on thefollowing steps:

1. Dispersionor emulsification of thecorematerialinasoluble protein slurry 2. Heating the slurry to form a protein melt followed by denaturation of the protein to render it insoluble in water in such a way that the insolublelayer coats the core droplets or particles The denaturation process can be achieved by heating only, bypH adjustment, orby using proteolyticenzymes which cause coagulation of the protein. It seems that this method could be suitable with various proteins, such as casein, albumin, corn and soy protein, wheat and rice gluten, and gelatin. 111.

SURFACE-ACTIVITY OF PROTEINS

All three methods described above have a common first step during the process of microencapsulation: adsorption of the proteinmolecule onto the

26

and

Magdassi

Vinetsky

oil-water interface. This adsorption may affect the emulsification of the oil phase (and hencethe droplet size distribution), the natureof the encapsulating wall (viapacking andinteractions between adsorbedmolecules), and the colloidal properties of the newly formed microcapsules (e.g., flocculation, zeta potential). Therefore, it is essential to understand the adsorption of proteins at oil-water interfaces, as well as the chemical parameters which may affect their surface activity. The proteins are macromolecules composed of a mixture of polar and nonpolar side chains and may have positive and negative electrical charges. These moleculesmay adsorb at theoil-water interface, through their hydrophobic side chains, and hencedecrease the interfacial tension of the emulsion system. The adsorption process is, in general, composed of three consecutive steps [lo]:(1) diffusion of the protein molecule fromthe bulk to the interface and attachment ontothe interface; (2) penetration of new (3) protein molecules, which have to overcome an energy barrier; and molecular rearrangement of the adsorbed molecules. It is interesting to note that the adsorption, or attachment at the interface, is achieved by small segments of protein having a unit surface area of 100-200 A2. The adsorption process obviously can lead to a drastic unfolding of the protein at theinterface, a phenomenon termedsurface denaturation. Under certain conditions, the protein present at the interface may coagulate; a result which may have implications on the formation of microcapsules by the protein-coated oil droplets. Since the first step in forming O/W microcapsules involvesan emulsification step, it is necessary to obtain an emulsion which has the required droplet size distribution and is stable, at least for the period of time required to complete the encapsulation process. Basically, it is preferred to use a protein which has some surface activity in terms of the initial adsorption and reduction of interfacial tension. Moreover, the proteins should be good emulsifiers, a property which can be obtained byusing nonrigid, charged protein that causes emulsion stabilization in both electrical and steric mechanisms. Such proteins are gelatin and casein, which are frequently used inprotein-based microencapsulation processes. If the emulsification step is achieved by using surfactants other than protein, the main property required of the coating protein is that it adsorbs onto the previously prepared emulsion droplets. Inthe case of simple or complex coacervation, it is required that the oil coacervate phase have suitable interfacial tensions, as shown by Arneodo etal. [ll],for gelatin coacervate phases and citrus oils. The surface activity of proteins is dependent on many factors [lo] such as themolecular parameters(MW, hydrophobicity, charge density and

Microencapsulation Emulsions Water of Oil-in-

27

isoelectricpoint,and rigidity) andthesolutionconditions(pH,ionic strength, temperature). Therefore,when forming protein-based microcapsules,oneshouldconsiderthe overall effectonboththeadsorption/ emulsification step and the formation of a wall around the oil droplets. IV. POSSIBLEFILM-FORMINGPROTEINS

As stated earlier, the of use certain proteins depends on the method used to prepare the microcapsules. From various publications, it appears thatthe number of proteins which can beused for microencapsulationis huge. In principle, any protein capable of forming a film is suitable for microencapsulationprovided the film can also be formed on the surface of the oil droplets. In a recent publication, Gennadioset al. [l21 review various edible coatings and films based on proteins. As shown, proteins such as zein, wheat gluten, soy protein, peanut protein, keratin, collagen, gelatin, milk proteins, andcasein can be used as film formers. Microcapsules were formed by emulsification and heat denaturation of egg albumin [13,14] around oil droplets; wheat gluten was used to encapsulate fat-soluble, edibleflavors by a spray-drying technique [15]; zein was used by dispersing the oil in alkaline protein solution,in which its pH was lowered to cause phase separation[16]; and casein was usedto encapsulate polyunsaturated fatsby spray drying [17-191. Qpical examples for theuse of various proteins to encapsulate oil droplets are given in Table 1. Of particular interestis the gelatin molecule, used either by itself or incombination with other componentsto form microcapsules [20]. Gelatin is a very abundant protein thatis produced from collagen of various origins. Gelatin is usually manufactured by two processes, “acid” and “basic,” which yield gelatin type A (positively charged below pH S) and type B (negatively charged above pH 5). The molecular weight of the proteins varies from 100,000 to 20,000, a property that is reflected in the bloom number, whichisindicative ofgel strength. Gelatin has unique surface activity and is a good emulsifier for various oil phases. Therefore, the various types of gelatin are often considered ideal candidates for the encapsulation process. For example, gelatinmay form capsules by the simple coacervation process, obtainedby addition of sodium sulfate [32], or by complex coacervation with gum arabic [7], sodium alginate [S], or even by using gelatin type Awith gelatin type B. We have recently reported [33] on the formation of gelatin-based microcapsules by formation of an insoluble complex of sodium dodecyl sulfate (SDS) with a positively charged gelatin. It was found that gelatin has the ability to bind anionic surfactants,such as SDS, and various soap molecules via electrostatic and hydrophobic interactions. At a certain mo-

28

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Microencapsulation of Oil-in- Water Emulsions

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Magdassi and Vinetsky

30

Soybean oil

Gelatin in water pH=6.1

T

V

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non-adsorbed gelatin

Aqueous solution of SDS, pH=6.1

.

v

d

Mix

Crosslinking agents

c

V Microcapsules

V

Removal of water (lyophilization, evaporation)

Fig. 2 Schematic presentation of formation of microcapsules of oil droplets in water by interaction of anionic surfactant with a cationically chargedgelatin.

Microencapsulation Emulsions Water Oil-in- of

31

lar ratio of surfactant to protein, the protein becomes uncharged and, therefore, is insoluble in water. Further addition of SDS leads to redissolving of the gelatin-SDS inwater, probably due tohigh charge (negative) density. It was therefore assumed that if the precipitation of the insoluble gelatin-SDS complex could occur at the surface of the oil droplets, we could obtain microcapsule walls. Since both gelatin and SDS have surface activity, this process may combine the emulsification ability of the two components, with their binding properties. The formation of microcapsules by this method is schematically presented in Fig. 2. As shown,the process is based on threesteps: 1. Emulsification of the oil by usinggelatin type A as theemulsifier. The emulsions which are formed by simple homogenizer are stable for at least several days, having a mean droplet size of several micrometers. The pH adjustment is crucial, since the SDS binding is strongly dependent on thecharges of the protein molecule. After theemulsion is formed, the freegelatin should be removed simplybyrinsing the emulsion with water during filtration or centrifugation. 2. The second step is addition of SDS solution at a specific concentration, which is predetermined according to the optimal surfactant-protein ratio which causes precipitation. Fig. 3 gives an

! l

-

$I

-1

Fig. 3 Microcapsules of soybean oil prepared by interaction of SDS with gelatin type A.

32

Magdassi andVinetsky

example of such microcapsules which are obtained without further crosslinking. 3. The last step is the addition of a crosslinking agent suchas glutaraldehyde. The hardened wallof the microcapsules allows simple separation of the microcapsules from the continuous phase, andif the continuous aqueous phase is completely removed(by lyophilization or evaporation), a dry powder of microcapsules is obtained. From the description of this process, it is clear that formation of microcapsules of oil droplets can beachieved by various protein-surfactant combinations provided that either the protein or the surfactant has some emulsifying activity,and theinteraction between the two can lead to precipitation of a suitable film. REFERENCES 1. C. A. Finch, Spec. Pub1.-R. SOC. Chem. 138(Encapsulation and Controlled

2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

12. 13. 14. 15. 16. 17. 18.

Release):35 (1993). B. H. Kaye, Kona (Hirakata, Japan) 10:65 (1992). P. Spiegl, Austrian PatentDE 244, 1890 (1975). D. Fredj and F. Dietlin, French Patent 84-14653 840925, 1986. Y. Okumura, H. Takai, M. Shiozawa, and K. Kaneko, Japanese Patent 84143268 840712,1986. H. C. Greenblatt, M. Dombroski, W. Klishevich, et al., Spec. Pub1.-R. Soc. Chem. 138(Encapsulation and Controlled Release): 148 (1993). L. Y. Sheen, S. Y. Lin, S. J. Tsai, Zhangguo Nangye Huaxue Huizhi 30(3):307 (1992), C. Arneodo, J. P. Benoit, C. Thies,S. T. P.Pharma 2(15):303 (1986). J. Janda, D. Bernacchi, S. Frieders, U.S. PatentWO-91-US7278911004, 1992. S. Magdassi and N. Garti, in Interfacial Phenomena in Biological Systems (M. Bender, ed.), Marcel Dekker, New York, 1991, pp. 289-299. C. Arneodo, A. Baszkin,J.P. Benoit, and C. Theis, in Flavor Encapsulation (S. J. Risch and G . A. Reineccius, eds.), American Chemical Society, Washington, DC, 1988, pp. 132-147. A Gennadios, T. H. McHugh, C. L. Weller, et al., in Edible Coatings and Films to Improve Food Quality (J. M. Krochta,et al., ed.), Technomic Publishing, Basel, 1994, pp. 201-277. S. Soloway, U.S. Patent 3137631, 1964. J. Kosari and G. M. Atkins, U.S. Patent 3406119,1968. P. P. Noznick and C.W. Tatter, U.S. Patent 3351531,1967. C. Brynko and J. A. Bakan, U.S. Patent 3116206,1963. T. W. Scott, L. J. Cook, K. A. Ferguson et al., Austr. J. Sci 32:291 (1970). T. W. Scott, and G. D.L. Hills, U.S. Patent 4073960,1978.

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19. G. J. Faichney, H. L.Davies,T.W.Scott et al., Austr. J. Biol. Sci 25:205 (1972). 20. L. S. Jackson and K. Zee, Lebensm. Wiss. Technol.24:289 (1992). 21. D. J. Mangold, W.W. Harlowe, andH. W. Schlamens, U.S. Patent 87-129503 871207, 1987. 22. Q.P. Corporation, Japanese Patent82-28757 820226,1983. 23. T. Kobayashi, French Patent 1568500 690523, 1969. 24. J. C. Soper, U.S. Patent 89401189,1991. 25. E. Bonatz, K.Golz, R. Nastke, German Patent90-342045 900625, 1991. 26. D. M. Whitaker, U.S. Patent 88-292495 881230,1991. 84-197776,1986. 27. S. Shimizu and H. Kato, Japanese Patent 28. R. M. Rawlings and D. Procter, Canadian Patent1086127 800923, 1980. 29. R. Shekerdzhiiski, S. Titeva, K. Mitikov et al., Formatsiya 35(4):23 (1985). 30. B. Sivik, J. Gruvmark, and M. Jakobsson, in Proc.-Scand. symp. Lipids,16th (G. Lambertsen, ed.), Lipidforum, Bergen,1991, pp. 147-152. 31. H. Jizomoto, E. Kanaoka, K. Sugita et al., Pharm. Res. 10(8):1115(1993). 32. J. Rosenblat, S. Magdassi, and N. Garti, J. Microencaps. 6515 (1989). 33. S. Magdassi andY. Vinetsky, J. Microencaps., accepted for publication.

Biodegradable Microspheres: Advancesin Production Techno Jean-Pierre Benoit Laboratoire de Phannacie Galbniqlue et BiophysiquePharmaceutique, Universitt? $Angers,and Centre dcE Microencapsulation, Angers, France

Hew6 Marchais Laboratoire de Pharmacie GaUnique et Biophysique Phannaceutique, Universitt? $Angers,Angers, France

Hew6 Rolland and Vincent Vande Velde Centre de Microencapsulation, Angers, France I. Introduction 11. Polymer Phase-SeparationMethods

111. Solvent Evaporation and Solvent Extraction Methods A. General Description of the Process B. Solvent Evaporation (Emulsification-Evaporation) C. Solvent Extraction (Emulsification-Extraction) IV. Spray-DryingMethods A. The NebulizingSystem B. Apparatus C. Applications D. AdvantagesandInconveniences V. Methods Using Fluids Under Supercritical Conditions A. Solvation of Compounds B. Precipitation of Pure Compounds C. Crystallization of Pure Compounds D. Formation of Microparticles VI. Milling Methods

36 36 43 43 45 49 50 52 53 53 55 57 58 59 60 60 62

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36 1.

INTRODUCTION

Major advancesin the therapeutic sciences and substantial developments in the field of biotechnology have taken place over the past few years which have allowed the syntheses of peptides and proteins on industrial scales. The much valued pharmacological importance of these molecules has been somewhatoffset by their poor ability to penetrate thegastrointestinal barrier. Likewise, their high sensitivities and fragilities together with their very short biological half-lives seriously limit their therapeutic applications. The delivery of proteins and peptides for therapeutic purposes thus presents a major challenge to pharmaceutical scientists. Different strategies have been proposedto overcome these problems. Various modes of administration, such as the nasal or transdermal routes, have been investigated to facilitate the resorption of some proteins and peptides. However, the parenteral route remains the dominant means of delivery of peptides and proteins. From a formulation point of view, different drug-delivery systems involving, e.g., liposomes, mixed micelles, and nanoparticles, have shown much promise, as have controlled-release implants based on the use of biodegradable polymers. The success of the latter technique has contributed to the renaissance of the microencapsulation field. A large number of microencapsulation processes, and modifications of processes, have been reportedin the literature and in various patents [l].In this chapter, emphasis is placed on the description of selected processes leading to microparticle formation froma particular series of polymers used as coating materials; namely, poly(a-hydroxy-carboxylic acids) (PHCA). This category of polymers covers polymers and copolymers of lactic and glycolic acids (PLA, PLAGA),which are the sole excipients presently approved for use by international health authorities for designing biodegradable parenteralimplants. In referring to thevarious pharmaceutical formsdiscussed herein, the authors define microparticles as polymeric entities falling in the range of 11000 pm, covering two types of forms: Microcapsules, micrometric reservoir systems Microspheres, micrometricmatrix systems This chapter deals solelywith the preparation and use of microspheres. The potentials of the various technologies addressed hereafterare also discussed.

II. POLYMERPHASE-SEPARATIONMETHODS Polymer phase-separation, in nonaqueous media, by nonsolvents or polymer addition, also referred to as coacervation, is an excellent technique for

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the entrapment of water-soluble drugs such as peptides, proteins, or vaccines to beused as microencapsulatedpreparations. This technique has beenknown sincethe 1960s [2]. Since then numerous textbooks and articles have appeared describing the principles and preparation methodswhich, if adhered to, lead systematically to theformation of microcapsules [l-61. Under certain circumstances, it has been demonstratedthat this procedure leads to the production of microspheres. Thus, the coacervation of a polymer suchas poly(d,l-lactic acid-coglycolic acid) (PLAGA) dissolved in methylene chloride with a second polymer such as silicone oil allows the formation of matricial systems[7]. If crystals of active principles are placed in suspension at thebeginning of this process, they will be capturedin these matrices after the desolvation of PHCA. The coacervation process of a PHCA, as shown in Fig. 1, can be divided into four steps [7]:

1. In step 1 (Fig. lA), the amount of phase inducer added to the appearsto form a solution is low (l-5% v/v).Siliconeoil pseudoemulsion in the organic phase. 2. In step 2 (Fig. lB), for a greater amountof silicone oil introduced in the medium, the beginning of a phase separation is noticeable. The droplets of coacervate are unstable and merge together to give larger structures which expand andfinally rupture. 3. In step 3 (Fig. IC), the quantity of polymer added is sufficient to allow a stabledispersion of PLAGA coacervate droplets. 4. In step 4 (Fig. l D ) , extensive aggregation of coacervate droplets occurs which leads to their precipitation. To prepare wellindividualized microspheres, step 3 must be attained by carefully controlling the addition of the phase inducer to avoid massive aggregation. This sequenceof events can be conveniently schematized by a phase diagram. On such a diagram, each point corresponds to a defined weight percentage of methylene chloride, PLAGA, and silicone oil. Fig. 2 shows four ternary diagramsresulting from different PLAGA batches having very close mean molecular weights (Mgpc ranging from 35,000 to 55,000) but differing in the percentage of oligomers present: 2.0, 3.1, 4.5, and 40.0% for polymers 1, 2, 3, and 4, respectively. These ternary diagrams express the influence of the proportions of the three compounds on the sequence of events describing the PLAGA desolvation. In thefollowing discussion, we have designated step 3 as the “stability window.” Significant differences in the areas of stability windows are shown in Fig. 2. PLAGA 1 leads tothe largest zone, corresponding to step 3, whereas PLAGA 3 and 4 exhibit narrower windows. Increasing amounts of

38

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Fig. 1 Sequence of events occumng ina 5% PLAGA solution following the addition of 350 CSsilicone oil. silicone oil were required to be added to PLAGA solutions to induce the formation of stable coacervate droplets according to the sequence: polymers 4, 3,2 and 1, with polymer 1requiring the greatest quantity. These results concerning the influence of the physicochemical nature of PLAGA on the stability window indicate that a relationship exists between theamount of silicone oil added to induce the formation of stable coacervate droplets and the ratio of oligomers in polymer batches involved in the phase separation. The more monodisperse the copolymer, the more methylene chloride acts as a good solvent for the coating material. Consequently, increased quantities of silicone oil are needed to desolvate the copolymer. Thus, polymer 4, which contains the highest percentage of oligomers among all of the copolymers studied, does not readily dissolve in methylene chloride. In other words, it requires low amounts of silicone oil to be precipitated. Under such conditions, the formation of well-individualized microspheres is very difficult to control (i.e., small stability window).

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SILICONE

SILICONE OIL

2c$Em

20 1

l .-c

METHYLENE CHLORIDE

MTEKLE~CHLORIDE

SILICONE

350 CS

1

/

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METHYLENE CHLORIDE

Phase diagramsfor the coacervationof different PLAGA batches. step 2; step 3 (stability window); step 4.

step

From a formulation point of view, the width of the stability window can also be modified by changing the viscosity of the silicone oil. Fig. 3 illustrates the role of the viscosity of the silicone oil on the formation of stable coacervate droplets of polymer 1. For a low-viscosity-grade silicone oil (i.e., 20 CS), no stability window could be detected. By increasing the viscosity of the phase inducer up to a value of 12,500 CS,it was possible to enlarge the stability window. Above this limit, the silicone oil could not be handled for microencapsulation purposes. The evolution of the stability window versus the viscosity of the phase inducer is common for all the polymers studied. A phase inducer of 20 CS leads either to a medium of low viscosity that is unable to stabilize coacervate droplets or toits too rapid solvation by methylene chloride, which in turn induces uncontrolled precipitation of the wall material. These two phenomena may occur simultaneously, although in practice this is verydifficult to demonstrate. Silicone oils of higher viscosities produce the reverse effect and allow one to increase the area of the stability window.

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-

METHYLENTCHL

METHYLENE CHLORIDE

M-NE

CHLORIDE

Fig. 3 Phase diagrams for the coacervation of pol mer 1. Influence of the phase inducer viscosity on the stability window. step 1; step 2; step 3 (stability window); step 4. It must be also noted that, as expected, the average molecularweight of the coating material also determines the existence, width, and displacement of the stability window within phase diagrams[8]. Fig. 4 summarizes the two main characteristics of a PLAGA batch which may influence the stability window and therefore theformulation of microspheres. Whenthe weight averagemolecular weight decreases, solvation in methylene chloride is good, and the amount of silicone oil necessary to coacervate the polymer is high. A displacementto theleft of the stability window is noted. Conversely, when the copolymer batchcontains low molecular weight compounds, i.e., oligomers, the overall hydroof more numerphobicity of the coating material decreases and the presence ous carboxyl and hydroxyl groups necessitates the use of smaller quantities of silicone oil. This induces a displacement to theright of the stability window often accompanied by a decrease in its area. The PHCA composition also exerts a strong effect on the stability window [g]. d,l-PLA appears to be less prone to desolvation induced by silicone oil addition than PLAGA copolymers of comparable molecular weights. The stability window which appears during the coacervation of d,l-PLA is therefore broadened compared with those of the copolymers.

Biodegradable Microspheres

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METHYLEECHLORIDE eflicient polymer dissolution

WHEN

"-

bad polymer dissolution

" c -

kw DECREASESWHEN

HYDROPHOBICITY DECREASES

( /low

m : STABILITY

molecular weight compounds)

WINDOW

-

Fig. 4 Influence of polymer Mw and hydrophobicityon displacement of stability window. As a practical consequence,the isolation of well-individualizedmicrospheres is facilitated when d,l-PLA is used. The authors attribute this phenomenon to a nonrandom distribution of glycolic acid in the copolymer chains, since the solubility parameters of d,l-PLA and PLA25GA50 are quite similar: 16.4 and 16.8 (J m-3)112, respectively. Glycolic acidunits tend to form microcrystalline domains which are more sensitive to desolvation and precipitation. Numerous microparticular systems resulting from thecoacervation of a PHCA previously dissolved inhalogenated solvents have been described in the literature. A summary of the main operating conditions is given in Table 1. One must bear in mind that the investigations cited in Table 1 speak of "microcapsules," whereas in reality,matrix-type systems are more likely to be obtained. DCcapeptyl LP, 3.75 mg, a speciality polymer-drug preparation commercialized in Europe byIpsenA3iotech (France) and by Ferring (Germany), is prepared following this fabrication process. It consists ofmicrospheres of PLAGA 25 : 50 (25% l-lactic unit, 25% d-lactic unit, and 50% glycolic unit) containing an analogue of luteinizing hormone-releasing hormone (LHRH), triptoreline, used inthe treatmentof prostate cancer and endometriosis [19]. Formulation parameters have been adjusted to allow a constant release of triptoreline in vivoover 1month. This kinetic profile was extremely delicate to obtain. It is known, for instance, that a compromise

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Benoit et al.

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must be made between theintensity of the initial peptide loss and the time interval over which the sustained release occurs [20]. Again, the residual oligomer ratioin a PLAGA batch is one of the variables which can drastically change the release profile of the peptide. Lanreotide, which is a somatostatine analogue, has been approved and is in use in France for the treatment of acromegaly [21]. This drug is delivered from the same type of microsphere prepared according to the aforementioned technology. In conclusion, compared withspray-drying or solvent-evaporation methods, the polymer phase-separationtechnique may protect active principles from beingaltered by exposure to heat or from their partitioning out into dispersing phases. Residual solvent concentrations in resulting microspheres, however, can be very high, especially when heptane is chosen as the hardening agent [15,22]. 111.

SOLVENTEVAPORATIONANDSOLVENTEXTRACTION METHODS

The solvent evaporation technique was fully developed at the end of the 1970s [23]. Since then numerousstudies have been carried out on thebasis of this method. A. General Description of the Process

This technique is based on the evaporation of the internal phase of an emulsion by agitation. Initially, the polymeric supporting material is dissolved in a volatile organic solvent. The active medicinal principle to be encapsulated is then dispersed or dissolved inthe organic solution to form a suspension, an emulsion, or a solution. In the following step, the organic phase is emulsified under agitation in a dispersing phase consisting of a nonsolvent of the polymer, which is immiscible with the organic solvent, which contains an appropriate tensioactive additive. Once the emulsionis stabilized, agitation is maintained and the solvent evaporates after diffusing through the continuous phase. The result is the creation of solid microspheres. On the completion of the solvent evaporation process, the microspheres held in suspension in the continuous phaseare recovered by filtration or centrifugation and are washed and dried (Fig. 5) [24]. Although the concept of the solvent evaporation technique is relatively simple, the physicochemical phenomena governing this process are very complex. This system is characterized by the existence of several interfaces through which mass transfer occurs during particle formation (Fig. 6) [25]. The formation of solid microspheres is brought about by the

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Y

Organic solution o f polymer with active principle dissolvcd o r dispcrsed with tensioactive agent

Dispersing medium

pq 0

0

/

Formation of emulsion under mechanical stirring

4)

Evaporation of solvent

Formation of solid microspheres

Fig. 5 The principle of the preparation of microspheres following the solvent evaporation technique.

evaporation of the volatile solvent L1 at interface L2/G. During the course of solvent evaporation, a partitioning is produced across the interface L1/ L2 from the dispersed phase to thecontinuous phase leading to theformation of solid microspheres. The partitioning across the interface L l L 2 is, however, not limited to the organic solvent; the active principle may also partition to some extent at this interface. The rapidity and importance of

Biodegradable Microspheres

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Fig. 6 Schematic representationof the different interfaces in the solvent evaporation process: (1) Simple emulsion; (2) multiple emulsion.

Fig. 7 Differentmodes of incorporation of active principles in an organic solution of polymers. (a) Active principle dissolved. (b) Active principle dispersed. (c)

Active principle emulsified. these different transfer processes have a direct effect on microparticle formation. Thus, by studying the various governing parameters, the establishment of optimal conditions for theformulation of individual polymer/active principle couples can be achieved. Several systems may be envisaged based on the natureof the external phase (aqueous or nonaqueous), the incorporation mode of the active principle in the organic solution of the polymer (dissolved, dispersed, or emulsified) and the elimination procedure of the organic solvent (evaporation or extraction). (Fig. 7). The classification proposed by Aftabrouchad and Doelker [26] is in part adopted in the following discussion. '

B. SolventEvaporation(Emulsification-Evaporation)

1. Oil-in-WaterEmulsion Techniques calling on water as nonsolvent to thepolymer are in general preferred.In fact, theseprocesses are extremely economical and negate

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Benoit et al.

the recycling of the external phase, which inturn results in the formationof well-individualized microparticles [27-291. In this system, the polymer is dissolved in an organic solvent such as methylene chloride or chloroform. The active principle is dissolvedor dispersed in the same medium, and then the entire mixture is emulsified inan aqueoussolution containing an appropriate surfactant. Beck et al. were thefirst to propose this procedure for the encapsulation of progesterone in PLA microparticles [23]. This technique, particularly suitable for theencapsulation of lipophilic active principles, has since been widely used for the encapsulation of different classes of therapeutic agents such as steroidal hormones [30-351, cytostatics [36-401, antiinflammatories [41-461, and neuroleptics [47,48]. Nevertheless, the microencapsulation of hydrophilic active principles by this process can pose problems. In actual fact, a partitioning phenomenon operates betweenthe dispersed and the dispersing phases which contributes to a substantial lowering of microencapsulation yields. The physicochemical characteristics of the active principle, such as its partition coefficient, its degree of ionization, or its surfactant character will play animportant role in its localization in one or otherof thetwo phases of the starting emulsion. Strategies permitting a reduction in the loss of water-soluble active principles have been proposed to increase microencapsulation yields. The solvation of medicinal substances in organic solutions of polymers can be achieved by the addition of hydrophilic cosolvents [49,50]. The advantage of solubilizing the active principle is related to the flexibility of producing particles of extremely small sizes(i.e., bordering onthe micrometer scale) whose internal structures are more homogeneousirrespective of the initial drug particle size. The water solubility of the active principle can be reduced by chemically modifying it prior its incorporation in the organic phase. This modification is simple to perform for water-soluble salts, as they are commonly lipophilic in their nonionized forms. Many medicaments have been encapsulated in this manner [51,52]. The chemical synthesis of lipophilic prodrugs fromwater-soluble species such as5-fluoro-2’-deoxyuridine[53] or cephradine [54] has likewise permitted higher microencapsulationyields. In this instance, however, structural modifications of the medicine may give rise to toxicological problems. One may also modifythe dispersing phase of the emulsion to reduce leakage of the active principle from the oily droplets. Several investigators have thereforesuggested saturating the continuous phase with the medicine [37,38,55-571, adjusting the pHof this same phase,or even addingelectrolytes [40]. It must be noted however, that thevalidity and effectiveness of these strategies vary from case to case. For example,chemical transformations only concern a limited number of active substances, and the saturation of external phases is onlyof value for substances with lowwater solubilities

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or inexpensive compounds. All the same, if the adjustment of the pH can prove advantageousfor ionizable medicines, this procedure generally accelerates thedegradation of polymers suchas poly(a-hydroxycarboxylic acid) by the hydrolysis of the ester bonds[55,58].

2. MultipleEmulsions:Water-in-Oil-in-Water Another approach has been proposed for the efficient encapsulation of water-soluble active principles by the solvent evaporation technique in aqueous continuousphases. In these systems, active principles to beencapsulated are incorporated in an aqueous solution, which is poured into a casting organic solution of the polymer to form an emulsion of the type water-in-oil (W/O). This primary emulsionisitselfemulsifiedin an external aqueous phase leading to a multiple emulsion of the type water-in-oilin-water (WlOnV). The organic phase acts as a barrier between the two aqueous compartmentspreventing the diffusion of the medicine towardthe external aqueous phase(Fig. 8 ) [29]. This technique was first proposed by Ogawa et al. [59-611 for the encapsulation of a peptide analogue of LHRH. This process proves much more effective when the watersolubility of the medicine is high(>900 mgl mL) [60] and partitioning between the organic phase is disfavorable. The same investigators have demonstrated thecrucial role of the viscosity of the primary emulsion (W/O) in the prevention of the diffusion of the active principle toward theexternal aqueous phase[59]. To this effect, they incorporated an agent (gelatine) capable of holding the active principle in the aqueous internalphase, whereas increasing this phase's viscosity. The effectiveness of such a formulation may nevertheless be compromisedby a clash of compatibility between theviscosity-enhancing agent (e.g., gelatine, pectine, agarose) and the active principle which limits its field of application. This process is particularly suitable for the encapsulation of active principles inweak doses, which are strongly water soluble (e.g., hormones, trophic factors) [62-651, and antigens [66-731. Using this technique, the Takeda Laboratories (Japan) havemarketedbiodegradable microspheres of PLA37.5GA.25 (37.5% 1-lactic unit, 37.5% d-lactic unit, and 25% glycolic unit) containing an analogue of LHRH, leuprolide acetate. In France, this speciality is marketed under the name of Enantone LPand is prescribed in thetreatment of prostate cancer along with DCcapeptylLP. In an identical manner, the release of the peptide is effected over a period of 1month [74]. 3. NonaqueousEmulsions This technique, also known under the name of oil-in-oil emulsion, is replaced by can berepresented by Fig. 5 if the continuous aqueous phase an oily phase. Naturally the dispersed phase (also of an oily nature) needs to betotally immiscible withthe continuousphase. The dispersing medium

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Benoit

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Formation o f an crnuisic (water-in-oil type)

Aqueous solution

o f active principle .

..

L

Organic solution of polymer

Formation of cmulsion undcr mcchanical stirring (watcr-in-oil-in-water type)

3

Evaporation o f solvent

0

-

Formation of solid microspheres

Fig. 8 Preparation principle of microspheres according to the multiple emulsion technique (water/oiYwater). can by constituted by a mineral or vegetable oil or a nonvolatile organic solvent [57,75-771. Although this process essentially permits one to avoid the loss of water-soluble active principles, it has only been used for the encapsulation of a very limited number of active principles, including cytostatics [49,78-801, anti-inflammatories [81,82], antimalarials [83], and

Biodegradable

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49

serum albumin [84,85]. A very interesting innovation was proposed by Iwata and McGinity [86]: multiphase microspheres of PHCA containing water-soluble drugs and proteins have been prepared using the multiple emulsion-solvent evaporation technique. A multiple emulsion of the W/O/ 0/0type was utilizedto first provide a protective barrier between the drug and the polymer-solvent phase and secondly to prevent the highly watersoluble drugs from partitioning out of the microspheres. To illustrate this, a drug was dissolved in an internal aqueous phase containing gelatin and Tween 80. This solution was then emulsifiedin soybean oil containing appropriate tensioactive agents. This primary emulsion was in turn dispersed in acetonitrile in which a PHCA was previously dissolved and a W/ 0/0emulsion was obtained. This emulsion was finallyadded to a hardening solution consisting of Span 80 and light mineral oil. It must be stressed that the protective oil phase and the external dispersing phase need to be immiscible withthe organic solution containing the polymer. In certain processes, solvent evaporation may be replaced by sublimation by the implementation of lyophilization after the emulsification step. Thus, DeLuca et al. dissolved an active principle as well as poly(glyco1ic acid) in hexafluoroacetone sesquihydrate and emulsified the mixture in carbon tetrachloride [87].The resulting medium was immersed in dry ice/ methanolandcooled to a temperature which froze the drug-polymersolvent phase and not the continuous phase. The suspension was then subjected to freeze drying, allowing the removal of the continuous phase solvent and solvent entrapped in embryonic microspheres. In addition to giving elevated microencapsulated yields for watersoluble components, this procedure canhelp prevent the eventual hydrolysis of the medicine or polymer. However, by comparison with aqueous emulsions, this technique exhibits a number of disadvantages related to the use of the nonaqueous solvents: these are often expensive and need to be recycled. Trace residues are oftendifficult to eliminate, which pose technological problems. C. SolventExtraction (Emulsification-Extraction)

In the emulsification-evaporation method, the organic solvent of the dispersed phaseof the emulsion iseliminated in two stages: 1. Diffusion of the solvent in the dispersingphase (solvent extraction) 2. Elimination of the solvent at the dispersing phase-air interface (solvent evaporation).

In theory, if one uses a continuous phase whichwill immediately extract the solvent(s) of the dispersed phase, the evaporation stage is

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no longer necessary in the formation of microspheres. In practice, this can be achieved by using large volumes of dispersing phase with respect to the dispersed phase [88,89] or by choosing a dispersed phase consisting of cosolvents, ofwhich at least one has a great affinity forthe dispersing phase. One may even formulate a dispersing phase with two solvents in which one acts as a solvent extractor of the dispersed phase [68,90].

A hybrid technology may also be envisaged where emulsificationevaporation is initialized then interrupted before the volatile solvent is totally eliminated. At this point, the native microparticles are transferred into a large volume of the continuous phase(consisting mostoften of highpurity water) where remainingsolvent is eliminated by extraction. This method was advocated by Benita et al. [31] for the preventionof the development of active principle crystals at the microparticle surface during the solvent evaporation stage or in the continuous aqueous phase, leading to decreased efficiencies inthe microencapsulation of medicines. In fact, ithas been suggested that thetransfer of embryonic microparticles in a large volume of water leads to rapid extraction of organic solvent which is found at the periphery of dispersed globules which still contain some solvent. This leads to the formation of a polymeric barrier at the organic solvent-continuous phase interface, which slows downthe diffusion of the active principle from theorganic phase towardthe aqueous external phase and prevents crystal formation of the active principle [32,91]. An interesting modification of this technology was proposed by Leelarasamee et al. [92]. If the solvent extraction method is retained, the formation of dispersed globules is assured by the injection of a polymeric support solution containing the active principle in a solvent-extracting medium. The entire mixture is placed in a tubular system with reservoirs designed to permit the total extraction of the solvent from the polymer. This system replaces the mechanical stirring blade which assures emulsification in the preceding step. The concept behind this system is to enable the eventual production of microspheres ona continual basis.

IV. SPRAY-DRYINGMETHODS Nebulization, or spray drying, is widely used in the chemical, pharmaceutical, and foodindustries [93]. The principle of spray drying bynebulization rests on theatomization of a solution (containing the product to be dried) by compressed air or nitrogen through a desiccating chamber and drying across a current of warm air (Fig. 9).

Biodegradable Microspheres

L

I 8

I

Fig. 9 Laboratory spray dryer: (1) Drying air inlet + filtration; (2) heating; (3) desiccation chamber; (4) cyclone; ( 5 ) collector for drying powder-microspheres; (6) filtration + air outlet.(A) solution, suspension, emulsionto spray. (B) compressed or nitrogen air. (C) Spray nozzle (e.g., pneumatic, ultrasonic) Four separate phases may be distinguished: 1. Nebulization of the solution in theform of an aerosol 2. Contact of the nebulized solution with the warm air 3. Drying of the aerosol 4. Separation of thedriedproductandtheaircharged solvent

with the

This technique can be used to protect sensitive substances against oxidation (e.g., fish oils, essential oils, vitamins, colorants) [94]. When volatile substancesare encapsulated, the coating materialsmost often employed are maltodextrine and arabic gum.

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A more recent application of this mode of drying has been adapted for the creation of matricial systems of the microspherical type starting from complex liquid mixtures comprising an active principle dissolved (or dispersed) with a polymer in an organic solvent. This type of process permits the isolation of microspheresor microcapsules depending on theinitial formulation, albeit in the form of a solution, a suspension, or anemulsion (Fig. 10).

A. TheNebulizingSystem W Oprincipal nebulizing systems exist[95]: 1. lbrbine method: nebulization by a rotating disk or rotary atomization 2. Atomizer method: nebulization by compressed air or pneumatic nozzle atomization

In the atomizer method,the internal diameter of the atomizer is often between 0.5 and 1.0 pm. Compressed air (or nitrogen) is then mixed directly in the central core along with the solution; nebulization thus being provoked simultaneously by the pressure applied to the interior of the liquid vein and by its ejection velocity from the nozzle. Compressed air subsequently arrives at the exit of the atomizer along with the liquid, inducing the shearing of the liquid vein. This system is convenient for the isolation of extremely fine particles ( 4 0 pm) from nonviscous solutions. In the turbine method, the pressure of the compressedair turns a disc with calibrated orifices at high speeds (25,000-35,000 rpm), provoking the

EMULSION SUSPENSION SOLUTION Liquid drug Polymer Solvent

Solid drug Polymer Solvent

Liquid drug Polymer Solvent

MICROSPHERE MICROCAPSULE

Fig. 10 '&pes of microparticlesobtained by spray drying accordingto the nature

of the active principle-polymer mixture.

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shearing of the solution from the moment of its passage through these orifices. The turbine methodis thus complementary tothe first technique, being adaptedfor very viscous solutions, and for the formation of particle sizes in the range of > l 0 pm. The turbine equipped with its nebulizers must havea desiccation chamber of >l m in diameter. B. Apparatus

A vast array of spray-drying equipment exists ranging from laboratory nebulizers (Minispray Dryer 190; Biichi,Switzerland) to pilot scale apparatus (model Minor Mobile; Niro Atomizer, Denmark) the first to productionscale equipment (Minor Productions; Niro Atomizer) and ultimately to drying towers of several meters in height. This process is thus industrializable, with users having at their disposal a range of accessories in function with the quantities of products to befabricated. Changes in particle sizes can thus beeffected in a progressive manner. Various factors, such as thermal exchanges and losses, variable yields, and the geometries of atomizers or turbines make it difficult to transform the laboratory-scale equipment to theproduction scale. Most reportsin the literature are based on results obtained from laboratory-scale equipment (Minispray Dryer190; Biichi). Being of reduced sizes, these models have limited evaporation capacities. Flow rates are typically of the orderof 15-20 mL min". The entrance temperature of the air (40-200°C) allows the user the choice of working with aqueous or organic solvents of low boilingpoints. Only a few industrial companies operate using equipment of greater dimensions [96,97], but notably none employs the atomization tower for the drying of powders. C. Applications

The formation of biodegradable microspheresby this technology, permitting the liberation of medicines over periods of months or longer from one unique injection, is a relatively recent development [96-1001. Polymers and copolymers of lactic and glycolic acids are used in the formation of some of these systems. To date, only bromocriptine (Parlodel LAR; Sandoz, Switzerland) is commercialized in the form of injectable microspheres, which originates from the spray drying method. This formulation is composed of PLAGA branched to D-glucose [96]. Other patents (busereline [loll, mifepristone [102]) make use of this process for the encapsulation of hydrophilic or lipophilic molecules, allowing their liberation over a period of 1 month after one injection only. The principal applicationsoriginating from this technology are detailed in Table 2.

54

I

0

m m

SE:

I

41

U

..

S

Y

0

3

..

E: c

8

Benoit et al.

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D. AdvantagesandInconveniences

The interest in this technique lies, among other things, on the following factors: It can be likened to a continuous process, whereas, in the pharmaceutical industry, the preparationof batches is encountered. Certain models are able to discharge their material contentsin a dry, continual manner (during which thermal treatment processes for the drying of powders must be steadily maintained over the whole process and closely monitored). The residence times of products to be dried at elevated temperatures (>lOO"C) from aqueous solvents are very short. In the case of organic solvents, evaporation temperaturescan be even lower. For example, aworking temperature of 30-40°C is sufficient to evaporate methylene chloride. These parameters are important when considering the drying of thermally sensitive products. The majority of authors evaporate polymer-methylene chloride solutions at temperatures ranging between 50 and 70°C [90,98,105,108]. Particle size distributions areusually monodispersed with a Gaussian distribution more or less depending on the pulverizing head employed (e.g., pneumatic atomizer, disc) and the chosen nebulization parameters such as, for example, the pressure of compressed air, the internal diameter of the atomizer, theviscosity, and flow rate of the solution tobe nebulized. In the literature,particle sizes are commonly described as being less than 10 pm, since the geometries of atomizers and the viscosities of nebulized solutions are almost uniform. In published articles for certain polymers, for example, dl-PLA (209,000 MW) or PLAGA (22,000 M W ) [105], the obtention of microspheres is impossible when the Kquid vein cannot be sheared into fine droplets. In these cases, the final result is the formation of filaments. This phenomenonalso arises when the concentration of the polymer is increased in the organic phase, hence increasing the viscosity of the solution [98]. It also appears that theproduction of these filaments is more readily brought about employing polymers composed of linear chains [98]. The formation of these filaments at this point is also strongly influenced by the performance of the apparatus. The influence of the pressure of the nebulizing air, the geometry of the atomizer (the diameter of its orifice), and the flow rate of the nebulizing solution are important for filament formation. Nebulization of fine particles permits, under certain conditions, to keep active principles at the heart of polymeric matrices in amorphous forms.

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On the other hand, Bodmeier and Chen[98]have observeda change in the crystalline form of progesterone nebulized on its own or in the presence of d,l-PLA. These state changes imposekinetic modifications of dissolutions, which may be exploited in pharmacokinetics. Spray drying thus permits the incorporation of lipophilic molecules in these types of copolymersin concentrations between 30 [l051 and 50% [98], withouttheappearance of crystals at particle surfaces. On theotherhand, for concentrations greaterthan 50%, or for hydrophilic molecules such as theophylline [98], the burst effect is more marked in the in vitro release kinetics. This can be explained for the former case by surface crystallizations and in the latter by weak interactions with hydrophobic polymers, which are not sufficient to prevent crystallizations. Residual levels of organic solvents are less than those from the emulsification and evaporation techniques. Levels of 0.1-0.2% can be found quoted in the literature [97]. Certain investigators have managed to arrive at concentrations of 0.05% and below [106]. For a pharmaceutical application, the concentration of organic solvent, by specified unit mass, must be inferior to thelevel set by the country of commercialization. For example, the standard set by the European Pharmacopaefor methylene chloride is less than 5OOppm (0.05%). Inferior residual solvent levels can be obtained by lyophilization of microspheres, and residual humidity levels are typically very low (
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In our experience, no kinetic variations could be observed from variations of these above parameters. The sole influence on release characteristics arose frommodifications of starting formulations: Variations in molecular weights of polymers or thetypes of polymers (L-PLA, d,l-PLA, and PLAGA) Variations of the viscosities of mixtures Variations of active principle/polymer ratios It would be of great value to have some knowledge of these variables (sizes, structural arrangements) for the transition from laboratory to industrial scales. These changes in scales entail modifications on the geometries of atomizers and their evaporation capacities and the flow rates of nebulizer solutions. From actual knowledge, the development of spray drying for the formation of microspheres based onPLA and PLAGA is interesting forthe fabrication of sizable quantities of microspheres (
METHODS USING FLUIDS UNDER SUPERCRITICAL CONDITIONS

The technique of placing a fluid at a temperature and pressure exceeding its critical point and using it asan extractant is now welldocumented [109]. Increasing applications of this technique are being discovered, with particularly promising ones being found in the field of material transformations (changing physical properties of material). Thus, in the pharmaceutical sciences, where modifications of the physical properties of drugs are necessary, several benefits can also be achieved by taking advantageof supercritical conditions. As described earlier in this chapter, drug-loaded microspheres are often formedby dissolving at least one of their two major components. In practice, one of three procedures can be followed:

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The drug can bedissolved and combinedwith the coating substance. Both the drugand coating substances are dissolved. Only the coating substances are dissolved. Thus, we have examined successively how fluids under supercritical conditions are able todissolve the drug or the container material itself. In a second step, we have studied how the drug (dissolved or not) can be embedded in the microsphere matrix.

A. Solvation of Compounds An important step in all the techniques presented hereafter is the solubilization of the active principles. Compared with the classic use of solvents, supercritical fluids offer a larger scope of choice in terms of solvents’ “performances” (Table 3). Also, the possibility of altering their abilities to solvate solute molecules existsby varying pressure and temperatureconditions. These two parameters are key features in supercritical fluid techniques. In the vicinity of the critical point especially, minute changes in temperature or pressure generate substantial density variations. Solvating properties can be related to thedensities of supercritical phases. Thus, the ability of supercritical fluids to penetrate a substance increases with their densities.

Table 3 Critical Pressure for Selected Fluids with Critical Temperature Below150°C Fluids

T (“c)

P (bars)

Nitrogen Carbon monoxide Argon Oxygen Methane Krypton Ethylene Xenon Carbon dioxide Ethane Hexafluoro-sulfur Propylene Propane Ammonia Isobutane

-147 -140 -123

34 34 48 50 46 55 50 58 73 48 37 45 42 1 1 1 36

-119 -83 -64 9 16 31 32 45 92 96 132 134

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The literatureprovides basic data concerning the solvating properties of an array of pure solvents under their supercritical conditions [109-1121. In most cases, however, the precise conditions described in the literature are not encountered, as multicomponent mixtures are more commonplace in the laboratory. Added to this, for several good reasons (e.g., like the sensibility of products to high temperatures or forreasons of toxicity), the scientist chooses the “wrong solvent.’’ An example of this situation is encountered in the use of carbon dioxide, a good choice for food applications but chemically difficultfor the solvation of polar substances. Under supercritical conditions, however, carbon dioxide exhibits more or less the same solubility as heptane. Thechemist, on the other hand, may avoidthis poorsolubility problem by adding cosolvents to the mixture (usually polar substances). Needless to say, this chemical mixture no longer acts according to the literaturewhen exposed to supercritical conditions. Its behavioris nevertheless independent on each pure component constituting the mixture. The operatorshould bear in mind that supercritical conditions can often be lost in complex mixtures and that two-phase systems may develop instead. Thus, performing practical tests using small-scalelaboratory installationsis the only reliable way to observe these phenomena and to obtain the data needed forlarge-scale project development. Another method to obtain relevant information, but somewhat exhaustive, is to search for suitable data for each particular project in the large amountof literature dealing with extraction processes exploiting supercritical conditions [113,114]. Since carbon dioxide is often employed, the solubility data provided by IUPAC is extremely useful [115]. In general, low molecular weight hydrocarbons and lipophilic substancesare good candidates tobe dissolved with supercritical carbon dioxide. Now that the desired starting material is in solution, several treatments can be applied, bearing in mind that the final step will be to recover the dissolved compound possessing the now new and interesting properties so that it can be called the product. B. Precipitation of Pure Compounds

How can the dissolved compound be recovered? Supposing that a substance A is placed in a classic solvent, S1, and that a solution of A is obtained. At the molecular level, this implies that molecules of A are far away from one another and are surrounded only by solvent molecules S1. Supposing we have a gas, G1, under supercritical conditions which exhibits a lower solubility for compound A than does S1, but which is miscible and soluble in S1. Then when the solution of A in S1 is placed in the presenceof G1, the supercritical conditions will probably disappear and a two-phase

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system will be generated. The lighter phase, predominantly composed of G1, contains a small proportion of S1. The heavier phase, comprising mainly by A and S1, also contains some amount of G1. The presence of G1 in S1 decreases the solvating properties of the mixture, so A precipitates out of the liquid medium in its amorphous state. By precipitation, we mean the formation of solid entities which “can be seen by the human eye,” having granulometric sizes in the micrometric domain. Recovering A, after supercritical processing, gives the product AGlS1, which does notnecessarily exhibit the same crystallinity, morphology, porosity, surface properties, or solubility properties as the product AS1 generated by a classic chemical methods [116-1211. The principle is helpful in explaining the processes discussed in Section V.D. C. Crystallization of Pure Compounds A variation of the aforementioned precipitation method is when conditions are adjusted to induce crystallization rather than precipitation. A solution of compound B in a concentration close to its saturation in a supercritical fluid G2 (considered as the solvent of B) can be transported to an areaof the system where the temperature is lower. In doing this, the system becomes saturated in B and crystallization occurs. Highly pure monocrystals can be obtained by this process. Later on, placed back in normal atmospheric pressure and temperature, it becomes very difficultto detect traces of the solvent G2 in the sample [122]. It should be pointed out that the retrograde solubility phenomenon (decreasing solubility with increasing temperature) can be observed [123,124]. Here also, before considering the formation of microparticles, the scientist should remember that the crystallographic arrangements of compounds may be modified by the nature of the crystallizing solvent. Thus, a compound embedded in a microsphere may very well exhibit a crystallographic lattice which differs from the starting bulk material. This is not necessarily due to the close proximity of the host microsphere matrix molecules, but it can be due to the nature of the solvent used during the process.

D. Formation of Microparticles If an excipient is added to a solution of the active principle, the basic precipitation operation as described above can be applied. Moreover, if the starting solution is fashioned before precipitation occurs, a final product can be obtained in one step. For example, if an organic solution of an active principle is sprayed through a nozzle into a column containing supercritical fluid, the organic solvent is extracted by the fluid from each finely divided droplet of the solution [125,126]. Under supercritical condi-

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tions, the solute precipitates immediately to form a microparticle powder. An interesting example is the one where biodegradable polyesters are dissolved (PLA, PLAGA). Depending on the degree of crystallinity and on the molecular weights of polymers employed, different types of microparticle formations have been observed[127]. The case where the drug N-butylscopolammonium bromideis added to poly-L-lactide can be cited [128]. Another interesting example is the formation of biodegradable microparticles of enzymes such as catalase and lysozyme and proteins such as insulin from dimethylsulfoxide (DMSO)-water or dimethylformamide (DMF)-water solutions [129,130]. The use of peptidelike materials raises the question of the solubility of DMSO, DMF, and waterin the supercritical fluid. Can solute precipitation be related to polarity changes occurring in the mixtureso that it can be described as a "gas antisolvent precipitation," or is the organic solvent extracted first by the supercritical fluid, so that theextraction is playingthe major role? Related to this question is the interesting study reported by Lokensgard [131]. Following this protocol, methylene chloride can be extracted from PLAGAmicrocapsules downto undetectable levels by supercritical carbon dioxide, whereas residual heptane is only poorly removed. Residual heptane can nevertheless be removed by the use of propane instead of carbon dioxide. In spite of these advantages, the destruction of microspheres has been reported in some instances while operating under supercritical conditions. In actual fact, the extraction properties of supercritical fluids can be tailored to extract residual solvents in pharmaceutical compounds. The purification of glycyrrhizin is an example [132]. A 20,000-ppm sample in ethanol treatedwith supercritical carbon dioxide at 40°C and under350 kg cm-' can be reducedto 455 ppm. In somecases, this operation leads to the formation of porous spongelike matters [133]. Atomization techniques (rapid expansions from supercritical solutions) can be applied to a solution under supercritical conditions. In this process, material is dissolved inthe supercritical fluid and expanded into a lower pressure environment. This results in the nucleation of the solute.By nucleation, we mean that solute molecules gather together to form nanometric entities. Following this event, heat energy needs to be given to the system, since the supercritical phase cools as it decompresses. Akin to standard atomization techniques, where the solvent undergoes a phase change from a liquid to a gas, energy needs to be supplied. This event occurs, however, at much lower temperatureranges during a typical supercritical process; e.g., k200"C in an atomization tower for water compared with 5 5 ° C for carbon dioxide [134]. A review of this particular field has been published [121,135]. Addition of a polymeric substance in the pre-

-

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expansion solution can beof particular interest in order to obtainmatrices for controlled drug-delivery devices [136-1411. VI.

MILLING METHODS

International health authorities and scientific bodies have recently focused their attention toward theinvestigation of toxic residual solvents in active ingredients and pharmaceutical formulations, and they have imposedmaximum limits on particular solvent [142]. Confronted with these new directives, microparticle manufacturers are trying to concentrate their efforts on solvent-free technologies. One of them is based on the milling of PHCA particles. This technique is preceded by the fusion of the polymer powder and can be completedby a spheronization step. Thegrinding technique by itselfwasfirst introduced byBoswell and Scribner in 1973 [lo]. They proposed melting a PHCA and thoroughly dispersing a non-heat labile drug in the melt. The melt was then congealed in a mass and ground into small particles 1-200 pm in size. Similarly, they advocated spray congealing as an alternative technique for the processing of microparticles. In actual fact, milling of a glassy solid in order to produce microparticulate implants can be performed: After cooling of molten PHCAin which appropriate amountsof drug are dispersed [143,1441 After compression of PHCA particles [1451 After extrusion of PHCA [146,147] Cooling, compression, or extrusion followed by milling culminates in the formation of irregular external particle surfaces. Hence, microparticles formed thereafter arenot truly spheroidal and display broad size distributions. This was the reason why Wichert and Rohdewald proposed complementary steps enabling the spheronization of particles [144]. Particles resulting from the milling of the cold melt (a mixture of d,l-PLA and the neurotropic agentvinpocetine) were added to a buffered 0.05% Ween 80 solution heated to 180°C. After emulsification, the microparticles were isolated either by pouring them onto ice water or by spray drying. Although the microparticles were not as smooth as those prepared by the solvent evaporation process, they looked rather spherical. Ruiz has used a similar concept to formulate LHRH- and somatostatin analogue-loadedmicroparticles [146]. The microparticles, called “microballs” by Ruiz, were obtained according to the following steps. In the first stage, the cryogenic grinding of the extruded polymer, containing a homogeneously dispersed peptide, led to the formation of heterogeneous and porous particles. In a second stage, after sieving, the microparticles

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were suspended in a gel under energetic stirring. The gel was heated and the microparticles fused, becoming spherical. When spheronization was achieved, the suspension was rapidly cooled and thegel was dissociated by the addition of an appropriatewashing agent. It should be noted that several investigators describe a reduction in the molecular weight of PHCA during heattreatment [144]. This point must be taken into account when designing biodegradable microparticles. ACKNOWLEDGMENTS

The authorswish to thankDr. Michael Lynch for his assistance in preparing this chapter. REFERENCES 1. P. Deasy, Microencapsulation and Related Drug Processes, Marcel Dekker, New York, 1984. 2. J. A. Bakan, Microencapsulation via coacervation-phase separation, National W, June 1966. Industrial Research Conference, Land O’Lakes, 3. J. E. Vandergaer, Microencapsulation, Processes and Applications, Plenum Press, New York, 1974. 1976. 4. J. R. Nixon, Microencapsulation, Marcel Dekker, New York, 5. A. Kondo, Microcapsule Processing and Technology, Marcel Dekker, New York, 1979. 6. M. Donbrow, Microcapsules and Nanoparticles in Medicine and Pharmacy, CRC Press, Boca Raton,FL, 1992. 7. J. M. Ruiz, B. Tissier, and J. P. Benoit, Microencapsulation of peptides: A

study ofthe phase separationof poly (D,Llactic acid-co-glycolic acid) copolymers 50/50 by silicone oil, Int.J. Pharm. 49:69-77 (1989). J. P. Benoit,Influenceofaveragemolecular 8. J.M.Ruiz,J.P.Busnel,and weights of poly (D,L-lactic acid-m-glycolic acid) copolymers 50/50 on phase separation and in vitro drug release from microspheres, Pharm. Res. 7(9): 9.

928-934 (1990).

C. Thomasin, B. Gander, and H. P. MerMe, Coacervation of biodegradable polyesters for microencapsulation: A physicochemical approach, Proceedings of 20th Intern. Symp. Control. Rel. Bioact. Mater., Washington, DC, 1993, pp. 358-359. U.S. Patent 10. G. A. Boswell and R. M. Scribner, Polylactide-drug mixtures,

3773,919, 1973. 1 1 . J. S. Kent, L. M. Sanders, D. H. Lewis, and T. R. Tice, Microencapsulation of water soluble polypeptides, Eur. Patent0052 510,1986. 12. J. S. Kent, L. M. Sanders, D. H. Lewis, and T. R. Tice, Microencapsulation of water soluble polypeptides, U.S. Patent 207,864, 1980. ,

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13. J. S. Kent, L. M. Sanders, D. H. Lewis, andT. R. Tice, Microencapsulation of water soluble active polypeptides,U.S. Patent 4,675,189, 1987. 14. J. Komen and J. W. Groenendaal, Process for microencapsulation, Eur. Patent 0377 477,1990. 15. P. Orsolini, R. Y . Mauvernay,andR.Deghenghi,Prockdkpourlamicroencapsulation par skparation de phases de substances mkdicamenteuses hydrosolubles, Swiss Patent 665 558 A5, 1988. 16. B. Gander, C. Thomasin, H. P. Merkle, Y. Men, and G. Corradia, Pulsed tetanus toxoid release from PLGA microspheres and its relevance for imm genicityinmice,Proceedingsof20th Intern. Symp. Control. Rel. Bioact. Mater., Washington DC, 1993, pp. 65-66. 17. C. Thomasin, B. Gander, and P. H.Merkle, Physicochemical characterization of the coacervation process of poly (l-lactic acid) and poly (d,l-lactic acid) by silicon oil, Proceedings of 18th Intern. Symp. Control. Rel. Bioact. Mater., Amsterdam, 1991, pp. 646-647. 18. D. H. Lewis, T. 0. Dappert, W. E. Meyers, G. Pritchett, and W. J. Suling, Sustained release of antibiotics from biodegradable microcapsules, Proceedings of 7th Intern. Symp. Control. Rel. Bioact. Mater., Fort Lauderdale, FL, 1980, pp. 129-131. 19. H. Parmar, S. L. Lightman, L. Allen,R. H. Phillips, L. Edwards, and A.V. Schally, Randomised controlled study of orchidectomy vs long-acting DTrp6LHRHmicrocapsulesinadvancedprostaticcarcinoma,Lancet I1 (8466): 1201-1205 (1985). 20. J. M. Ruiz andJ. P. Benoit, in vivo peptide release from poly (DL-lactic acidco-glycolicacid)copolymer 50/50 microspheres, J. Contr.Rel.16:177-186 (1991). 21. I. Heron, F. Thomas, M. Dero, J. R. Poutrain, S. Henane, F, Catus, andJ.M. Kuhn, Traitement de l’acromtgalie par une forme liberation prolongke du lanrkotide, un nouvel analogue de la somatostatine, Presse Med. 22 (11):526531 (1993). 22. B. W. Muller, J. Bleich, and B. Wagenaar, Microparticle production without organic solvent, Proceedings of 9th International Symposium on Microencapsulation, Ankara,1993, pp. 29-40. 23. L. R. Beck, D. R. Cowsar, D. H. Lewis, R. J. Cosgrove, C. T. Riddle, S. L. Lowry, andT. Epperly, A new long-acting injectable microcapsule system for the administration of progesterone, Fertil. Steril. 31(5):545-551 (1979). 24. P. J. Watts,M.C.Davies,and C.’ D. Melia,Microencapsulationusing emulsificatiodsolvent evaporation: an overview of techniques and applications, Crit. Rev. Ther. DrugCamer Syst. 7(3):235-259 (1990). 25. C. Thies, Formation of degradable drug-loaded microparticles by in-liquid drying processes, in Microcapsules and Nanoparticles in Medicine and Pharmacy (M. Donbrow ed.), CRC Press, Boca Raton,FL, 1992, pp. 47-71. 26. C. Aftabrouchad and E. Doelker, Mkthodes de prkparation des microparticulesbiodkgradableschargkesenprincipesactifshydrosolubles,S.T.P. Pharma Sci. 2(5):365-380 (1992).

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Advances in the Technology of ControlledRelease PesticideFormulations Arie Markus The Institutes for Applied Research, Ben-Gurwn University of the Negev, Beer-Sheva, Israel

Introduction I.

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11. Interfacial Polymerization

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111. PhaseSeparation

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IV.

QualityControl A. Storage Stability B. ReleaseRate of Pesticide C. Toxicity to Fish D. Acute Oral Toxicity to Mice E. Efficacy

82 82 82 83 85 86

Case Study-Antifly

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V.

1.

INTRODUCTION

The science of pesticide formulation covers a very broad field, since it deals with the development, production, and storageof the formulations [l-41, as well as withthe interactionof pesticides withthe environment,including plants, insects, animals, soil, air, and water [5]. Pesticide formulations can be classified into thefollowing types:

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Markus Aqueous solutions Emulsifiable concentrates Dispersion concentrates (aqueous and nonaqueousflowables) Wettable powders Dry flowables (water-dispersible granules) Controlled-release formulations Others (e.g., dusts, aerosols)

The choice of formulation is influenced by the following factors: the physical properties of the pesticide (melting point, solubility, volatility); the chemical properties of the pesticide (hydrolytic stability, thermal stability, irradiation stability); the mode of application of the formulation (soil vs. foliar); the crop to be treated and agricultural the practices; the biological properties of the pesticide (crop selectivity, transport, LD,, for mammals and nonmammalianspecies); and economicconsiderations. This chaptercovers controlled-release techniques. Pesticides are conventionally applied to crops by periodic broadcasting or spraying. Very high, and possiblytoxic, concentrations are applied initially, and these often decrease rapidly in the field to concentrations that fall below the minimum effective level. As a result, repeated applications are needed to maintain control[6]. The formulation of a pesticide must thus be designed to meet the multifaceted demands of efficacy, suitability to mode of application, and minimization of damage to the environment. Controlled-release formulations meet these demands in that they enable smaller quantities of pesticide to be used more effectively over a given time interval and in that their design enables them to resist the severe environmental processes that act to eliminate conventionally applied pesticides; i.e., leaching, evaporation, and photolytic, hydrolytic, and microbial degradation [6]. In most instances, the rate of removal of a conventionally formulated pesticide follows first-order kinetics [6-91. The time, t,, that an effective level of pesticide is maintained after a single application is given by:

where M, is the minimum effective level, Mm is the amount of agent applied initially, and k, is the rate constant [6]. It follows from Eq. (1) that an increase in the effective duration of action of a conventionally applied pesticide would require than an exponentially greater quantity of the pesticide to beapplied. If, however, the pesticide could be maintained at the minimum effective level by a continuous supply from a controlled-release system, then the optimum performance of the insecticide would be realized, and this duration of action, to, would be given by:

Controlled-Release Pesticide Formulations

75

- Me)(KdMe)

(2) where kd is the rate constant for pesticide delivery from the controlledrelease device [6]. The relationship between the level of application and the durationof action of a pesticide is shown Fig.1 for a conventional formulation and for a controlled-release formulation [6]. The area between the two curves represents the amount of pesticide that is wasted. It is apparent that fora short duration of effectiveness, e.g., 1 week or less, the efficiency of the conventional method is adequate. As theduration of effectiveness increases, the efficiency of the conventional system decreases exponentially (as Fig. 1indicates). In addition to the advantages described above, controlled-release pesticides have other important advantages over conventional formulations [6]: to

=

(Mm

Reduction of mammalian toxicity for highly toxic substances Extension of duration of activity at an equallevel of active agent Reduction of evaporative losses and of flammability of liquids

L

1,000

loot

CONVENTIONAL PESTICIDE

l

Fig. 1 Relationship between the level of application and the duration of action for conventional and controlled-release formulations. The graphs are plotted using equations 1 and 2 and assuming that the half-life of the pesticide is 15 days and the minimum effective level is 1 g/acre. (Reprinted from Ref. 6.)

76

Markus Reduction of phytotoxicity Protection against environmental degradation Reduction of leaching into the earth andtransportation into streams Reduction of contamination of the environment Increased convenience of use by conversion of liquid materials into solids and flowable powders Separation of reactive components Control of the release of active agents Decreased costs because less active material is needed Convenience of handling.

Although the advantages of controlled release are impressive, the disadvantages of the technology must not be forgotten. Each application has to be examined individually and the positive and negative aspects weighed carefully [ 6 ] .Some of the following aspects of controlled-release formulations may be deleterious and thus require careful appraisal: (1) costs of the materials and processing of the controlled-release preparation, which may be substantially higher than standard formulations; (2) fate of the polymer matrix (see below) and its effect on the environment; (3) fate of polymer additives, such as plasticizers, stabilizers, antioxidants, and fillers; (4) the environmental impact of the degradation products of the polymer matrix produced in response to heat, hydrolysis, oxidation, solar radiation, or biological agents; (5) cost, time, and probability of successin securing government registration of the product,if this is required [6]. Various techniques have been proposed for the production ofmicrocapsules [lo-131; one source suggests that hundreds of methods are to be found in the scientific and patent literature. In general, these methods for microencapsulation can beclassified as: Separation from an aqueoussolution Formation by polymer-polymer incompatibility Interfacial polymerization Polymerization in situ Drying froma liquid state Solvent evaporation from anemulsion Gelation in the liquid state by cooling Desolvation Physical techniques, such as spray drying or fluidized bed reactions, are not usually suitable for encapsulation of pesticides. Similarly, addition polymerization is not suitable for pesticides, since many pesticidescontain phosphorus, nitrogen, or sulfur atoms, which are known to be radical scavengers, or since the compounds may not be stable in acid or alkali media. Interfacial

Formulations Controlled-Release Pesticide

77

polymerization is the method of choice for highly toxic insecticides, since the active ingredient is completely enveloped by the polymer and in most of cases release takes place via diffusion. This technique facilitates a dramatic decrease in toxicity. Both phase separation and interfacial polymerization techniques may be used for nontoxic pesticides. The only technique suitable forbiological pesticides is phase separation: With any other technique, the active ingredient, which in this case is a biological microorganism, would not be able to cross the envelope surrounding it. II. INTERFACIALPOLYMERIZATION

The technique for the microencapsulation of pesticides by interfacial polymerization comprises two stages. First, a liquid, molten, or dissolved pesticide containing a dissolved monomer is agitated at a high speed in an aqueous solution. The pesticide in which the monomer is dissolved constitutes the organic phase. Interfacial polymerization at the dropletsurface is then completed by the addition of a second water-soluble monomer to the continuous aqueous phase. The resulting aqueous slurry may be used as such or the pesticide-containing capsules may be recovered as a dry powder. When the pesticide is water miscible, it is also possible to carry out interfacial polymerization by making a dispersion of the liquid or molten pesticide in an organic phase, such as high-boiling petroleum ether. The following factors influence the performance of the capsules:

1. Composition of the polymeric capsule wall: The monomers can be chosen so as toproduce a variety of interfacial polycondensation products as wall materials; e.g., polyamides, polyesters, polyureas, polyurethanes, or polycarbonates. Within each of these categories, a range of polymers may be produced, depending on the starting monomers. It is also possible to vary the wall composition by forming copolymers in a simple process that is based on a mixture of oil-soluble or water-soluble monomers. 2. Degree of crosslinking: For cases in which a higher degree of wall integrity is essential, multifunctional monomers are used. 3. Capsule wall thickness: The thickness of the capsule wallis a function of the concentration of the monomers. In theproduction of the capsule, the wall continues to thicken until all of the available monomer is consumed. 4. Capsule size: The size of capsules is determined by the degree of agitation and by the type of emulsifying agent used in the continuous phase. Capsule size can be varied from an average diameter of a few micrometers to a millimeter. If there is a requirement for

78

Markus a formulation that must be sprayed, the diameter of the capsule must beless than 100 pm. 5 . Physical form of product: The final product may take the form of an aqueous slurry or of microcapsules that can be filtered off, washed, and dried to a free-flowing powder. Aqueous slurries have been found to be useful in the formulation of pesticides, since they need only to bediluted with water in order to be ready for field application as a sprayable product. In addition, they are cheaper, being more economical to manufacture. 6. Additives: A range of additives, such as ultraviolet light absorbers, antioxidants, and synergists, may be dissolved inthe oil phase (pesticide) being encapsulated. It should be remembered that these additives must be soluble in the oil and must not react with the monomers.

To date, a wide range of insecticides, herbicides, and fungicides have been encapsulated within variety of polymeric shells. At present, polyamide shells, especially polyurea andpolyurethane, are particularly popular. The company that introduced encapsulated pesticidaI formulations to the market is Pennwalt Corporation. Their first product was Penncap M(encapsulated methylparathion), and this was followed by KNOX OUT 2FM (encapsulated diazinon), Penncaptrin (encapsulated permethrin), and Pennphos (encapsulated chlorpyriphos) and later by the herbicide trifluralin [17]. Polyurea was used for the envelope, and the method of encapsulation is described in patents [14-171. The Pennwalt envelope was synthesized by the reaction of polyisocyanate with amines andacid chlorides. Another company active in the market is Stauffer Chemical Company, whose interest lies mainly in encapsulated herbicides. The Stauffer process for the production of the polyurea envelope differs from that of Pennwalt in that the isocyanate also serves as a source of amine. In the reaction process, some of the isocyanate is converted intothe amine, which then reacts with the remaining isocyanate [B-211. A number of other companiesare also involved inthe production of encapsulated biocides and herbicides. Monsanto Chemical Company markets encapsulated Alachlor underthe trade name of Lasso. The envelope for the Alachlor formulation is also based on polyurea andpolyurethane, and the method andchemicals-the isocyanates, amines, and theemulsifiersused byMonsanto aredescribed in their patents [22-261. Kedem Chemicals Ltd. (Israel) producesNO-ROACH (encapsulated diazinon) [27] and Antifly (encapsulated pyrethroids), and Dow manufactures Empire (encapsulated chlorpyriphos). In all the products mentioned above,the toxicity of

Controlled-Release Pesticide Formulations

79

the formulation is at least one tenth that of the emulsifiable concentrate (EC), and the efficacy lasts longer without a reductionof insecticidal activity. Several other companies [28-321 have used polyurethane or polyurea shells as envelopes for pesticides; among the biggest of these, we can mention Hoechst Bayer AG [33], and Ciba-Geigy [34]. The process for the production of encapsulated pesticides-in this case, diazinon and pyrethroids (Figs. 2-4)”is described schematically below. So far, we have discussed the interfacial polymerization technique in which polyamides are used as the envelop for the pesticides. However, experiments have been performed with envelopes made from a variety of polymers, including urea-formaldehyde and melamine-formaldehyde polymers, polyesters, epoxy, and polysilane. The formulations based on these polymers as envelopes have, so far, not been successfully commercialized.

Fig. 2 Encapsulated diazinon.

a

F Fig. 3 Encapsulated diazinon.

cI

m

Fig. 4 Encapsulatedpyrethroids.

Controlled-Release Pesticide Formulations

81

..

Production of encapsulated D e s e

water-soluble pesticide +

withemulsifier

: Emulsification

Aqueous solution of amines

Polymerization

111.

PHASE SEPARATION

The second technique used in pesticide encapsulation is phase separation, and the capsule obtained in this case is generally of the matrix type. This technique is suitable only for nontoxic and biological pesticides, since encapsulation into matrix-type capsules does not considerably reduce thetoxicity .of the product.An advantage of this mode of encapsulation is that, should the need arise, is it possibleto grind the formulation, since in most cases, the active ingredient is absorbed onto andwithin the polymer matrix. In the phase-separation technique, the active ingredient is suspended in a solution of the wall material. The wall polymer is then induced to separate out as a viscous liquid phase by coacervation (i.e., by adding a nonsolvent), by lowering the temperature, by adding a second polymer, or by a combination of these methods. Coacervation, which maybe simple or complex or of the “salting-out” type, is manifested as turbidity, droplet formation, or theseparation of liquid layers as described below. Simple coacervation occurs when a water-miscible nonsolvent (e.g., ethanol) is added to an aqueouspolymer solution, causing the formation of a separate polymer-rich phase, A typical example of a simple coacervation system is water-gelatin dissolved in water: this system is, however, difficult to control, so it is little used in practice. Complex coacervation takes place by the mutual neutralization of two oppositely charged colloids in aqueous solution. One of the most

Markus

82

widely used methods of microencapsulation by this process is the use of a solution of positively charged gelatin (pH <S), which forms a complex coacervate with negatively charged gum arabic. Other polymer systems may be employed and other electrolytes may be used with gelatin. Complex coacervation is closely related to the precipitation of colloidal material from solution: It immediately precedes precipitation. The process was originally developed in the 1950s for the coating used in the manufacture of carbonless copying paper, with gelatin and gum arabic as the two colloids. In salt coacervation, the polymer is separated out from the aqueous solution by “salting out,” typically by adding an electrolyte to an aqueous polymer solution. The method may be used to encapsulate water-insoluble oils or dispersed solid particles, but it is difficultto control the microcapsule size and the agglomeration of particles. The system may be stabilized by altering the pH ortemperature. The polymers thatcan be used in the phase-separation technique are summarized in Table 1[6]. Biological pesticides, such as Bacillus thuringiensis isruelensis (Bti) [36] and Trichodermaharzianum [37-391, have beenencapsulated in matrix-type capsules in which envelopes were produced from two types of polymers-natural polymers, such as alginates, or synthetic polymers, such as polyethylene. The methodsof polymerization are shown below.

W. QUALITY CONTROL The following parameters must be determinedfor any biocide formulation: the amount of active ingredient, size distribution, shelf life, biological release rate, toxicity, and efficacy. Since the first two types of tests are well documented, they will not be described here. The other tests will be described in detail below.

A.

StorageStability

A sample of the microencapsulatedpesticide formulation is maintained in a closed vessel in an oven at 54°C for 2 weeks or at 40°C for 6 weeks. The amount of the active ingredient in the productis analyzed before and after heating, and the formulation is considered to be stable if the amount of active ingredient does not change.

B. Release Rate of Pesticlde The apparatus used for this test is illustrated in Fig.5. One gram of encapsulated pesticide is placed on a plate, which is then sealed into a cylindrical glass vessel.The vessel isprovided with a digital thermometer andinlet and

Controlled-Release Pesticide Formulations

83

Table 1 Polymers that Can Be Used in the Phase-Separation Technique

Natural Polymers Carboxymethylcellulose Cellulose acetatephthalate Ethylcellulose Propylhydroxycellulose Gelatin Gum arabic Starch Bark Methylcellulose Galacturonicacid salt Polybutadiene Polyisoprene Neoprene Polysiloxane Styrene-butadiene rubber Silicone rubber

Zein Cellulose nitrate Shellac Alginate Waxes-paraffin Proteins Kraft lignin Natural rubber Methocel Synthetic Elastomers Hydrin rubber Chloroprene Butyl rubber Ethylene-propylene-diene terpolymer

Synthetic Polymers Polyhydroxyethyl methacrylate Polyvinyl alcohol Polyacrylate Polyethylene Polyacrylonitrile Polypropylene Chlorinated polyethylene Polystyrene Polyvinylpyrrolidone Polvether Ethylenevinylacetate copolymer Poly@-xylylene) Polyvinylidene chloride Polymethylmethacrylate Polyvinyl acetate Polyvinyl chloride

outlet tubes. The inlet tube is equipped with a flowmeter to control the volume of air introduced into thevessel. The air stream carries the released pesticide out of the vessel into a beaker containing ethanol. The temperature of the system is maintained at a predetermined value by an external heater. The amount of pesticide in the ethanol is determined, and the release rate calculated as a function of time.

C. Toxicity to Fish The fish (10per tank) are kept in 16-L aquaria under thefollowing optimal conditions: water quality as close as possible to pH 7 (neutral); water temperature 23-25°C; and good strong light for least 12 h a day (morelight makes them grow too fast). The test is performed in a room free of insecticidal contamination as follows. Adult fish of either sex are supplied

Markus

84 Production of cncausnlated

of part of the

Drying of the

of capsules

Filtration and washing

Controlled-Release Pesticide Formulations

85

thermocouple

I

Fig. 5 Apparatus for the determination of release rate of a pesticide from an encapsulated formulation. with adequate standardized food (Europet Basic food) before and after the experiment. Food is withheld for 2 days before the experiment. A solution of the pesticide formulation is obtained by placing the formulation in water in a high-shear mixer for 5 min. After addition of the pesticide to the fish tanks, mortality is checked after 3, 6 , 24, 48, 72, and 96 h. The times for 50% and 95% mortality (LTso and LT,,) are then determined from the results. D. Acute Oral Toxicity to Mice

It is preferable to use adult male animals (2.0-2.5 months) weighing 25-30 g. Standardized mouse food is given throughout theexperiment. A solution of the formulation is obtained by placing it in water in a Vortex mixer for 5 min. The quantity of the solution depends on theweight of mouse-l mL of solution is givenper 20 g body weight.The pesticide solution is placed in a 2-mL syringe and introduced into the stomach of mouse via the mouth. and 168 h, andLD, is Mortality is checked at 0,5,24,48,72,96,120,144, determined.

86

Markus

E. Efficacy

In this method, which measures the susceptibility of a population of cockroaches (BlutfeZZugermunicu) to a given insecticide, the cockroaches are exposed to standard insecticide residues in a Petri dish, and mortality is determined. The cockroaches should be obtained, as far as possible, from the same area, andkept in a suitable container until required. It is preferable to use adult males for the test, butif it is not possible to obtainenough males, females may be used. The test should becarried out in a room free of insecticidal contamination. The cockroaches are given adequate standardized food before the experiment. The cockroaches are first anaesthetized with carbon dioxide before being put into the insecticide-containing Petri dish. The Petri dish is then placed in an experimental vessel, which is maintained at25-30°C and relative humidity >25%. Solutions of the insecticide formulation are obtained by using a high-shear mixerfor 5 min. From the results, the times for 50% and 95% knockdown (LT,, and LTg5)can be

Fig. 6 Cockroach.

Formulations Controlled-Release Pesticide

87

determined. (A cockroach is considered knocked down if it fails to move on being returned to its normal posture.) Owing to thefact that thecapsules become attached to the cockroach (or ant) leg, they are introduced into the nest, where they then exert their action on all the insects in the nest.Figs. 6 and 7 show the capsule attached to theleg of a cockroach. V. CASE STUDY-ANTIFLY

Antifly (encapsulatedpyrethroids) is manufactured anddistributed by Kedem Chemicals Ltd. under license from Ben-Gurion University of the Negev, Beer-Sheva, Israel. TheLD,, of Antifly for mice, determined by the test described above, is 12,500 mg/kg and the LD50 for golden orfe fish 2200 pg/l. The biological efficacyof this encapsulated formulation versus that of the EC against cockroaches, flour beetles, house flies, and fleas, and ticks is shown in Fig. 8, and Tables 2,3, and 4, respectively.

Fig. 7 Closeup of cockroach leg showing diazinon capsule stuck to the leg.

Markus

88

110 '20

T

.

Emusified concentration

50 40

30 20 10 0

Blattellagermanica exposedto Fig. 8 % mortalityasafunctionoftimeof pyrethroids. a, emulsified concentration;U, Antifly. Table 2 Biological Efficacy After24 h of Antifly Against the Flour Beetle (Tribolium castaneum) % mortality at a concentration (ppm) of

Pyrethroid formulation Antifly EC

0

250

0

75

it

50

500

750

80 68

95

74

EC, emulsifiable concentrate. 0, control (without pesticide-water only) in the case the of capsules and solvent in the caseof EC product. Table 3 Biological Efficacy After24 h of Antifly Against the Housefly (Musca dornestica) Pyrethroid formulation Antifly EC

% mortality at a concentration (ppm) of 0

25

75 250

125

0

100

100

100 60

60

60 80

100 68

500

750

100

1 0 0 95

EC, emulsifiable concentrate 0, control (without pesticide-water only) in the case the of capsules and solvent in the case of EC product.

Controlled-Release Pesticide Formulations

89

Table 4 Biological Efficacy of Antifly Against Fleas and Ticks8

Parasite Dogs ~

type

of

level

Mongrel Ridgeback Mongrel Mongrel M Mongrel F M Pointer German M shepherd German M shepherd F Belgium shepherd Second spraying Spitz M F Spitz F Spitz F Terrier F Terrier German M shepherd F German shepherd F F F

~

1

sex First spraying Mongrel M

Presence of parasites after days

~

Fleas

High

l1

II

,l

l1

l1

n

l1

,l

N

,l

s

I,

s

l1

+ ticks

7

14

21

28

+ o o

0

0

+ o o + o o

0 0 0 0 0 0 0

60

0 0

0 0

0 0

0

0

0

0 0 0 0 0 0 0

I,

+ o o

0

0

II

n

+ o o

0

0

Fleas

2 2 1-2 2 2 2

0 0 0 0 0 0

0 0 0 0 0 0

0 0 0 0 0 0

0 0 0 0 0 0

0 0 0 0 0 0

0 0 0 1 0 0

2

0

0

0

0

0

0

n

,, I,

II ,I

c

+ o o + o o

T h e experiment was performed in an animal shelter in Tel-Aviv, Israel under the supervision of the Chief Veterinarian. The spraying was done directly on the dogs.No untoward side effects were observed. From this table we see that spraying the dogs with Antifly gave excellent results: 28 for After the first spraying, the dogs remained without fleas and ticks days; before the spraying, they were heavily infested with ticks and fleas. After the second spraying, they wereoffree fleas and ticks for 60 days. It is well known that the commercial product keeps the dogs free of ticks and fleas for a very short time (i.e., a few days).

90

Markus

REFERENCES 1. Geissbuhler, H. Advances in Pesticide Science, Part 3. Pergamon Press, 1979,

pp. 717-826. 2. J. Miyamoto and P. C. Keierney, Pesticide Chemistry-Human Welfare and the Environment,Vol. 4, Pergamon Press, 1983, pp. 241-400. 3. W. Van Valkenburg, Pesticide Formulations. Marcel Dekker, New York, 1973. 4. K. C. Seymour, Pesticide Formulation and Application Systems,2nd Conference. ASTM, Philadelphia, 1983. 5. H. B. Scher, Advances in Pesticide Formulation Technology, A.C.S., Washington, DC, 1984. 6. A. F. Kydonieus,ControledReleaseTechnologies:Methods,Theoryand Applications, Vols. 1and 2, CRC Press, Boca Raton,F L , 1980. 7. G . 0.Fanger, General Background and history of controlled release. Pesticide Symposium (N. F. Cardarelli, ed.), Universityof Akron, Ohio, 1974. 8. R. W. Baker andH. K. Lonsdale, Principles of controlled release, in Proceedings of Controlled Release Pesticide Symposium (F. W. Hams, ed.), Wright State University, Dayton, Ohio, 1975. 9. D. H. Lewis and D. R. Cowsar, Principles of Controlled Release Pesticides. A.C.S. Symposium Series 53, A.C.S., Washington, DC, 1977. 10. R. Arshdy, Preparation of nano and microspheres by polycondensation, J. Microencaps. 6:l-12 (1989). 11. R. Arshdy, Preparationof nano and microspheres by interfacial polycondensation, J. Microencaps. 6:13-18 (1989). 12. R. Arshdy, Preparationof albumin microspheres and microcapsules, J. Controll. Rel. 14:111-131 (1991). 13. C. A. Finch, in Encapsulation and Controlled Release.(D. R. Korsa andR. A. Stephenson, eds.), Royal Society of Chemistry, 1993. 14. J. E. Vandegaer and N.J. Wayne, U.S. Patent 3,577,515. 15. J. Shimkin, U.S. Patent 4,497,793,1985. 16. P. A. Cantilli, German Patent 2,722,973, 1977. 17. R. C. Koestler, U.S. Patent 4,360,376, 1982. 18. K. B. Scher, German Patent 2,312,059,1973. 19. H. B. Scher, German Patent 2,648,562. 20. H. B. Scher, German Patent 2,706,329,1977. 21. H. B. Scher, Belgian Patent 867646,1978. 22. G. B. Beestman, Eur. Patent 17409,1980. 23. P. J. Allert, German Patent 2726539,1977. 24. G. B. Beestman, Eur. Patent 148169,1985. 25. Z . B. Becher andR. W. Magin, U.S. Patent 4,563,212,1986. 26. G. B. Beestmanb, U.S. Patent 4,640709,1987. 27. A. Markus andZ.Pelah, U.S. Patent 4,851,271,1990. 28. M. N. Hitsu and K. Kabushiki, U.K. Patent 1,091077, 1967. 29. H. P. Crodts andJ. E. Karloske, U.S. Patent 4,517,326, 1983. 30. K. Riecke,U.S.Patent4,622,267,1986. 31. G. Garber, B. Chatenet, and P. Pellenard, U.S. Patent 4,309,213,1982.

Controlled-Release Pesticide Formulations

91

32.C. H. Nemeth, U.S.Patent 4,107,292,1978. 33.Bayer,Eur.Patent50,264,1981. 34. Ciba-Geigy AG, Eur. Patent 214936,1986. 35. A. Markus, Z.Pelah, and N. Aharonson, Israeli Patent 84219, 84910, 1992. 36.J.Margalit,A.Markus,and Z. Pelah.Effectofencapsulation on the presistence of Bacillus thuringiensis var. israelensis serotype H-14, Appl. Microbiol. Biotechnol. 19:1982, (1984). 37. J. R. Connick, W. R. Walker, and W. R. Goynes, Jr., Sustained release of mycoherbicides from granular formulations. 10th International Symposium on Controlled Release Bioactive Materials. San Francisco, 1983, p. 283. 38. H. L.Walker and J. R. Connick, Sodium alginate for production and formulation of mycoherbicides, WeedSci. 31:333 (1983). 39. D. R. Farvel, R. D. Marois, W. J. Lumsden, and J. Connick, Encapsulation of potential biocontrol agents in an alginate-clay matrix, Phytopathology 75:775, 1985.

and

5 The Useof Factorial Designin the Development of Nanoparticulate Dosage Forms Baruch Magenheim,* and Simon Benita School ofPharmacy, The Hebrew University of Jerusalem, Jerusalem, I s r a e l

Pascal Wehrl6 Laboratoire de Pharmacotechnie, Centredes Recherches Pharmaceutiques, Universitk Louis Pasteur, Strasbourg, France

I. Introduction

94

11. The Principle of Experimental97Design 111. Illustration of Experimental Design Application

98

IV.100Statistical Methodology 100 A. Preliminary Study B. Sequential Experimental Design Building 100 Modelsof the C.106 Validation V. Effects of the Controlled Variables on Nanoparticle Formation Properties 109 System A. PLA-Acetone-Dichloromethane B. Dichloromethane-Indomethacin-Poloxamer188 System C. Conclusion on the Application of Experimental Design to Nanoparticle Formation Process VI.

Summary

107

120 121 124

VII. Symbols

*Currentafliliation: Agis Industries Ltd., Yeruham, Israel. 93

94 1.

Magenheim et al.

INTRODUCTION

In the last two decades, the international scientific community has expended enormous efforts to create colloidal carriers with improved drug delivery that are appropriatefor systemic use 11-91, and theinitial development of nanoparticulate dosage forms should be viewed in this context. Colloidai drug carriers mainlyinvolve submicrometer emulsions, nanoparticles, liposomes, and lipid complexes. The body distribution of the active molecules depends on their physicochemical properties. Therefore, the molecules reach the target organ but also diffuse and distribute throughout the entirebody, sometimes exercising harmful and even toxic effects to healthy organs and tissues. This is the case with various anticancer drugs [l] and some antifungal drugs such as amphotericin B [lo]. Attempts have been made to modify the pharmacokinetic profile of various therapeutic classes of drugs through their incorporation in colloidal carriers [3,11-191. These references are not intendedto represent anexhaustive or up-to-date citation of the literature but rather to reflect the efforts made to evaluate the drugbiodistribution of these loaded colloidal carriers following intravenous administration. The potential applications of nanoparticles as colloidal drug carriers by different ways of administration have been extensively reviewed [20291. Evidence has been reported by various investigators [30-361 that, following oral administration, nanoparticles may cross the ‘intestinal mucosa or traverse the mesentery lymph vessels toward the lymph nodes. Nanoparticles may consistently improve the bioavailability of poorly absorbed drugs [36]. Some success has been achieved in enhancing the efficacy and reducing the toxicity of anticancer drugs loaded in nanoparticles, as reported in some reviews [21-231. These encouraging results have focused more andmore interest on nanoparticles as promising colloidal drugs carriers. Recently attempts have been made to deliver drug-loaded nanoparticles by means of novel routes of administration (e.g., transdermal, lymphatic) [37-401. Furthermore, new strategies are now emerging in nanoparticle research to allow the delivery of antisense oligonucleotides and DNA fragments, with incursion in the expanding field of gene therapy [41,42]. The potential applications of nanoparticles in AIDS treatment are also currently under investigation [43-451. Nanoparticles have been well defined by Couvreur [22] as a solid submicrometer drug carrier of a polymeric nature in the nanometer size. The nanoparticles are divided into two main groups: nanospheresand nanocapsules. Nanospheres are polymer matrices in which the drug is dissolved or dispersed, whereas nanocapsules consist of a polymer wall entrapping an oily core in which the drug is dissolved [46,47]. Since the term

Factorial Design of Nanoparticulate Dosage Forms

95

nunoparticles refers mainly to nanospheres in the literature, this terminology is used throughout this chapter. A large number of processes for the preparation of nanoparticulate drug carriers have beendescribed in the literature and have been reviewed by some authorsin the past [48], as well as in this book. The physicochemical characterization of nanoparticles has been reviewed as well [49]. The readers are referred to the above-cited references which present the current status of nanosphere and nanocapsule manufacturing processes and characterization. Nanoparticles are usually intended for parenteral administration. The polymers for parenteral use have to fulfill a series of strict requirements. They must be nontoxic, nonimmunogenic, biodegradable into nontoxic metabolites, and free of impurities. It is also desirable that they be easy to process and possible to prepareeconomically in large quantities [50]. Polyalkylcyanoacrylates (PACA)and polyesters arethe polymers most extensively usedto form thewall or matrix of biodegradable nanocapsules or nanospheres, respectively. The research group lead by Couvreur exhaustively investigated the PACA nanoparticles, and furtherinformation should be soughtin their numerous publications [22,51]. Polyesters are biodegradable polymers with long-standing safe use and acceptance as surgical sutures. They are easily prepared as homo- and copolymers in a wide range of molecular weights. These biodegradable polymers undergo simple hydrolysis forming naturally occurring nontoxic metabolites. Their degradation pathway is byhomogeneous, bulk degradation [50]. The family of the poly(a-hydroxy acids), mainlypolyglycolic and polylactic acids, and copolymers wasextensively studied in the field of drug-delivery systems development [50]. The preparation of polylactic acid (PLA) nanocapsules was first reported in the literature by Fessi et al. [52] using an interfacial deposition process following solvent displacement. The PLA nanocapsules were thoroughly investigated and exhibited marked advantages over previously studied colloidal carriers [53]. Despite promising experimental results, it was reported thatthese nanocapsules were not able to retain the drugin their oily internal core more thana few minutes under perfect sink conditions [54]. Therefore, PLAn'anoparticles in whichthe drugwas entrapped in the polymeric matrix in the absenceof any oily core material were prepared on the basis of the method reported by Fessi et al. [52] with some modifications [55581. Other investigators have also recently adopted a similar method for investigating the incorporation of drugs in PLA nanospheres [59,60]. The authors of this chapter prepared nanoparticles able to retain drugs over longer periods of time following their immersion in the release

96

Magenheim et al.

medium under perfect sink conditions which usually prevail in physiological conditions. We also reported theinfluence of the manufacturing process variables and formulation parameters on nanoparticle characteristics and drug release profile [55-581. The materials and methods utilized were reported in detail in the various cited publications. Therefore, only a schematic representation of the nanoparticle manufacturing process is shown in Fig. 1. The most dominant variables in the search for optimal nanoparticle formulation conditions were the concentrations of PLA, acetone, dichloromethane, indomethacin, and Poloxamer 188. Between these different parameters responsible for nanoparticle formation some interactions occurred. A faster and more accurate identification of these interactions was carried out by a factorial experimental design allowing modelization

Fig. 1

Schematic representation of the nanoparticlemanufacturing (From ref. 58.)

process.

Factorial Design

of Nanoparticulate Dosage Forms

97

using multiple regression. At the same time, an attempt was made to achieve optimization of the formulation. Experimental design is the application of statistical principles for data collection [61]. A designed experiment introduces systematic changes into the variables of a system with the objective of discovering which variables are responsible for the observed changes in the output. In a designed experiment more informative data can be obtainedfrom much fewer experiments than from traditional experimental techniques. This cost-saving approach allows researchers to collect useful data from every point of the experiment. This methodology has been introduced in the last few years and is being used more frequently with high successrates in the pharmaceutical sciences [62-641. Some recent examples of the fields of application include analytical chemistry [65-701, granulation and formulation of tablets [71-741, the preparation of beads [75], microcapsules [76,77], and microspheres [78,79]. In fact, some investigators use experimental design in the formulation of nanoparticles [SO-821. Experimental design was usually utilized only for formulationoptimization purposes. By this method, few experiments allowed the selection of the optimum conditions of product manufacturing in a multivariable environment. The objective of this chapter is to illustrate the application of the experimental design technique not only for optimization but also for elucidation of the mechanistic aspects of nanoparticle formation by spontaneous emulsification, as depicted in Fig. 1. The technique enables a huge body of information to begathered and statisticallyvalidated regarding the simultaneous influence of all the various controlled variables on nanoparticle characteristics and on the interactions between these variables. The information achieved while performing a relatively reduced amount of experiments has been exploited to gain physicochemical insights of the underlying complex phenomena involved in nanoparticle formation.

II. THE PRINCIPLE OF EXPERIMENTAL DESIGN Theoretically, all the parameters that can affect the physicochemicalproperties of a final product may be included in the experimental design. However, in pharmaceutical manufacturing processes, there may be numerous important parameters due to the large number of ingredients usually required for formulation. Even the equipment used in the multiple processing steps provides additional sources of variability. If all the variables are included in the experimental design, the search process may become fastidious and lead to a prohibitive number of experiments even if a screening approach such as the Hadamard matrix is initially applied. Therefore, ex-

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perimental design can be an important tool and asset if it is exploited by an investigator who has previous knowledge on the behavior of the system. The investigators must first identify the potentially dominant parameters that should be subjected to the screening process in order to limit to a reasonable extent the number of experiments needed. Some of the variables are initially fixed for technological, regulatory, or economical reasons. Other variables are found in preliminary experiments to be critical and therefore cannot be subjected to any variation without affecting fundamentally the desired final product features. A delicate equilibrium is established between what is and is not worthinvestigating. The researcher must not neglect important factors which may lead to misleading interpretations of the system behavior. Furthermore, factors with a priori negligible or in contrast vital effects have to beexcluded inorder tosimplify the experimental design. Investigator skills, knowledge, and experience play a crucial role in selecting the appropriatevariables. It should be pointed outthat in cases where all dominant variables are not included in the experimental design, even the most sophisticated models will usually not be useful. In these cases, the researcher should consider the inclusion of additional variables previously omitted. The sequential building nature of the experimental designs enables the goal to be achieved with an addition of only a minimal number of experiments. 111.

ILLUSTRATION OF EXPERIMENTAL DESIGN APPLICATION

To illustrate the previous statements, a practical example of the application of experimental design to the study of nanoparticle formation process is given here. In preliminary investigative work, dozensof variables were examined in an attempt to select the most important variables affecting the nanoparticle properties. This obviously leads to a huge numberof experiments. However, initial results and experience gained in disperse systems allowed us to fix the nature and concentration values of most of the preeminent variables. Biocompatibility, biodegradation, and regulatory acceptability motivated the selection of polyesters as nanoparticle-forming polymers. Among these polymers, the fast release of the drug from nanoparticles prepared with different copolymers of polylactic and polyglycolic acids resulted in the choice of d,l-PLA. Technological drawbacks excluded the use of other PLAstereocopolymers. Some organic solvents like chloroform were rejected owing to potential toxicity and carcinogenicity. The semipolar solvent acetonitrile was found to have a strong interaction with dichloromethane, preventing nanoparticle formation. Regulatory require-

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ments favored the selection of Poloxamer 188and human serum albumin as hydrophilic surfactants. However, very lowdrug entrapmentwas observed, with the latterapparently due toits protein-binding capacity. Phospholipids promoted the simultaneous formation of liposomes, leading to a mixed undefined nanoparticulate-liposomal system that was not desired. The temperature of the organic and aqueous phases andtheir stirring rate did not show any effect in the range studied. On the other hand, some of the variables such as pH and thesolvents’ evaporation rate were foundto beso critical that any variation would result in a drug entrapmentyield too low to be pharmaceutically acceptable. The controlled variables initially considered worth studying were the amounts of PLA, acetone, and dichloromethane used inthe manufacturing process (PAD system). This systemallows a priori different aspects of nanoparticle formation to beclarified. It permits the evaluation of possible interactions between acetone and dichloromethane during the migration of the formerfrom the organic phase to theaqueous phase andthe degreeby which both solvents affect PLA precipitation. The consequent changes in the ratio of the PLA to thesolvents may highlight the potential impactof the organic phase viscosity on nanoparticle characteristics. Additionally, by selecting only three controlled variables, it is possible to readily obtain descriptive graphs which allow an easy visualization of the effects, thereby facilitating their evaluation. The general statistical methodology based on multiple regression using the generalized least squares theory and which allows plotting this graphs is calledthe response surface methodology [65]. Many fundamental aspects of the nanoparticle formation mechanism were elucidated from these series of experiments [58-591. The drugpartitioning between the organic and aqueous phases was shownto play an important role. Thus, a second set of controlled variables was investigated in order exhaustively to assess the partitioning contribution to drug entrapment in the nanoparticles. This set consisted of the amounts of dichloromethane, indomethacin, and Poloxamer188 used inthe manufacturing process (DIP system). They wereselected for the following reasons: (1) The solubility of indomethacin in dichloromethane and in the aqueous phase ultimately determines its partitioning behavior; (2) the solubility inthe aqueous phaseobviously is dependent on the surfactant (Poloxamer 188) concentration; and (3) variations in poloxamer concentrations may help to estimate the effect of the surfactant on incipient nanoparticle stabilization. The most appropriate way to carry out the research would be to add these threenew controlled variables (DIP) to the experimental design used to evaluate the first three variables (PAD). Indeed, dichloromethane was common to both systems. However, the results obtained with the PAD system demonstrated that a level of less than 0.5 mL of dichloromethane

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led to the formation of nanoparticles with unacceptable properties. Thus, the PAD experimentaldesign couldnot serve as a base to evaluate the DIP effects with a complementary design. Nevertheless, it did allow for the concentrations of acetone and PLA to be set preferential for use in the DIP experimental design. Before presenting a detailed explanation of the experimental design results, an introduction to the statistical methodology is presented for a better comprehension of the subject.

IV. STATISTICAL METHODOLOGY Until recently, formulation and processes were investigated and developed simply by trial and error while varyingone factor at a time. Needless to say, this approach is both time and energyconsuming as well as costly. Furthermore, in the worst case scenario, misleading conclusions can be drawn, particularly when different variables interact. Owing to the multiple requirements, the modem designer can no longer be satisfied with a method that leads to uncertain and nonvalidated results. The use of a statistical approach is necessary. The response surface methodology [64,83-861 using an experimental designisaneconomical method aimed at collecting a maximum of information with a minimum of experiments. Another important advantage of this general method is that the experimentaldesigns can be changedprogressively until an appropriately efficient model is found to describe the studied phenomenon. A.PreliminaryStudy

Before building a factorial design, the experimental field must be defined precisely. If it is too large, it may lead to nonrealistic experiments and result in microenvironments inwhichsystem behavior is different (like peaks or valleys on a topographical map) and may be not detected.If it is too small, it may be far from the optimal region. Results yielded in preliminary work allowed the experimental fields to be defined corresponding to the inner volume of the cubes described in Figs. 2 and 3 for the PAD and DIP systems, respectively. B. SequentialExperimentalDesignBuilding 1. Hadamard Matrix

In formulation or process optimization tasks, a great number of controlled variables such as amountof excipients or process parameters may have an influence on the studied characteristics (defined as response variables).

Factorial Designof Nanoparticulate Dosage Forms

101

0.50

io I

S

E

Y

5 0 D

0.00

@ (D 0 25

,W'

0

@ 15

0

PLA (mg)

Fig. 2 Description of the experimental field and the Hadamard matrix, 23,and 33 experimental designsof nanoparticle formulation for the PAD system (see details in text). (From ref. 38.)

INDOMETHACIN (%) Fig. 3 Description of the experimental field and the Hadamard matrix, 23, and Box andWilsonexperimentaldesignsofnanoparticleformulationfor the DIP system (see details in text).

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The analysis of all these factors generally leads to a prohibitive number of experiments to be performed. Ascreening experimental design is the most convenient way to reduce this number. Such an experimentaldesign allows (by multiple regression based on the generalized least squares theory) the calculation of a very simplemathematical model.The polynomial equation (Eq. 1) includes only the main effects (terms of first order) of the different controlled factors (X,) on theresponse variable (Y):

I=I

For thepresent experimental field: Y =k

+ ax, + bX, + cX,

(2)

where k is the constant term (corresponding to boin Eq. 1) and a, b, and c are the regression coefficients (corresponding to b, in Eq. 1) of the dichloroequation. X,, X,, and X, correspond toPLA,acetone,and methane for the PADsystem and to dichloromethane, indomethacin, and Poloxamer 188 for the DIPsystem, respectively. Hadamard [87] developed an original matrix allowing the evaluation of such a model, based on the following equation: =

l+Hn+Hn I +Hn-H,

So that, for example, matrix H, (for n = 2) can be established by

recurrence calculation:

+l +l +l +l -1 +l -1 H.,= +l +l +l -1 -1 +l -1 -1 +l As for any experimental design, this matrix gives incoded values -1, +l, the levels of the controlled factors that are to be set for the experiments (Table 1). For thefactors studied in this example, theactual values are reported in Table1and the corresponding experimentsare defined by any setof four nonconsecutive vertices of the cubesdefined in Figs.2 and 3. One can see that only four experiments are needed to estimate the “linear effects” of three different parameters. The four experiments are performed and the response measured. If A is the matrix column containing the constant and the regression coefficients of the model to be evalu-

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Table 1 Matrix of Experiments for the Hadamard Experimental Design Real values, PAD Real values, DIP system system L (mg) (mL) (mu (%l (%)

no.

"iirzzTG k

x 1

x 2

x 3

1 2 3 4

+l +l +l +l

+l

+l

+l

-1 +l -1

+l

-1 -1 +l

Experiment

-1 -1

75 75 25 25

15 5 15 5

0.50 0.00 0.00 0.50

0.75 0.50 0.75 0.50

8.75 8.75 6.25 6.25

2.00 0.50 0.50 2.00

ated (Eq.5 ) , if X is the Hadamardmatrix (H4) (Eq. 4) and if Y is the matrix column containing the four measured responses (Eq. 6 ) ,

the theory of the least squares allows usto write:

Y=A*X so that

A = X"

Y

The inversion operation (-l) is generally a complex calculation for which a pocket calculator or a computer is needed. But here, because the Hadamard matrix is of size n.n, symmetrical, orthogonal, and contains only +l and -1 values, the multiple regression calculation can be done very easily:

(where nis the number of experiments and tis the transposing operation). Finally,

The model is validated by performing an additional central experiment. If the experimental result value for this point is different from the

et

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expected value, the model is rejected. Other models whichtake intoconsideration theinteractions between thefactors can beapplied by adding onlya few experiments tothe previous design.

2. Experimental Design of z3Type The extremevertices of the experimental fields shown in Figs. 2 and3 constitute a typical 2kexperimental design, withk the numberof controlled variables, 3 in this example. The generalized least squares theory is applied again in order to evaluate a model including the interaction Xi Xj:

3. Models for the Analysis of Quadratic Effects If the model including the first-order terms of main effects aswell as the second-order term of interaction does not fit the data, the response surface is probably not plane. Thus, this potentially important curvature can be described by a more complex model taking into account the quadratic effects (X,') of each of the controlled variables as reflected by Eq. (12): Y =C b ob +, . C X b, +u * X , . Z X bj + u-x

(12)

a. Box and Wilsoncompositedesign. The Box and Wilson method [84,85] is the most economical way of evaluating the model described by Eq. (12). This design is defined by the addition of six experiments at an axial distance of & CY from the center of the cube in directions passing through the center of each of the six faces, to the previous design (points outside the cube; see Fig. 3). The CY value is selected according to orthogonality and rotatability criteria and depends also on the number of central point replicates and controlled variables. The equations applied for CY selection are beyond the scope of this presentation and can be found in textbooks dealing with statistics and experimental design. Orthogonality implies that theestimates of the regression coefficients of the model are uncorrelated. This property is verified for the Hadamard and 23 experimental designs. For a composite design, only quasiorthogonality can be achieved. The rotatability property leads to a variance function which is constant on circles centered at the origin. The level of this function is dependent on thedistance to the origin and not on thedirection from this origin. Eq. (12) can be translated for the DIPsystem as:

Y

=k

+ a.DCM + b.IND + c.POL

+ d.DCM.IND + e.DCM.POL + f.IND.POL +g DCM' + h.INDZ + i.POL'

(13)

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of Nanoparticulate Dosage Forms

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where k is the constant term and a-i are the regression coefficients. DCM representsthedichloromethanevolume,andINDand POL the indomethacin and Poloxamer 188 concentrations in the formulation, respectively. It should be noted that the term (DCM.IND.POL) for the triple interaction between these three controlled variables is not included in Eq. (13). This is dueto constraints associatedwith the Box and Wilson quasiorthogonality condition which excludes any third-order term in the derived mathematical model.

b. Experimentaldesign of 33 type. The efficiencyof the experimental designs is defined as the ratio between the number of terms in the mathematical equation describing the model and the number of experiments needed to perform the appropriate experimental design. For three controlled variables, the efficiency increases from 34.5% for a 33experimental design up to 58.8% for the corresponding composite design using three center replicates in both cases. However, the Box and Wilson composite design described above cannot be applied to the PAD system, because it leads to negative values for some of the controlled variables (DCM CO). Therefore, a complete 33 type design must be applied by adding 18 experiments to the23 design, 1 at themiddle of each of the 12 edges and 1at the center of each of the six facesof the cube defined in Fig. 2.Eq. (12) can be translated for thepresent system as:

+ a.PLA + b.ACE + c.DCM + d.PLA.ACE + e.PLA.DCM + f.ACE.DCM + g.PLA.ACE.DCM + h.PLA2 + i.ACE2 + j.DCM2

Y =k

(14)

where k is the constant term and a-j are theregression coefficients. DCM represents the dichloromethane volume,PLA the weight of polymer, and ACE the acetonevolume in the formulation. In contrast to the composite design, in the 33-type design, there are much more experiments than the number of terms of the derived matheso matical equation.Therefore, orthogonality canneverbereached that the third-order term (PLA.ACE.DCM) can be included as shown in Eq. (14).

c. Reducingapproach-precise analysis. Having the 33experimental design utilized inthe study of the PAD system three controlled variables and one response variable at a time, a four-dimension space is defined, which cannot be graphically represented. Hence, further analysis keeping one of the threeresponse variables constant at onelevel each timewas performed (32models). This allowed the calculation and validation of models for each described in Fig.2 and the drawing of surface face of the experimental cube response and contour plots which permited an easy visualization of the

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different effects. For example, acetone and PLA were precisely analyzed for the different constant amounts of DCM. An identical procedure was used where PLA and acetone were kept constant at different amount levels as for DCM. Applying the same concept to the Box and Wilson design utilized in the study of the DIP system results in the definition of 22 models. This occurs because, differently than in the 33design, in the present case, the experimental points added to the 23experimental design are located outside the cube defined in Fig. 3, at an a distance from its center.However, these 2* models do not allow the detection of quadratic effects. Therefore, to allow the drawing of surface response and contour plots, the equations obtained with Box and Wilson design-derived models were recalculated while keeping the value of one of the variables constant each time. Being that the models derived from Box and Wilson experimental design validated, the recalculated equations are validated as well. Only illustrative graphs obtained at fixedlevels of indomethacin of 7.5% w/v are presented.

C. Validation of the Models The above models used to evaluate the different response variables were validated by means of the following statistical tests:

1. Analysis of variance (ANOVA) of which the principle is given in Table 2. The experimental F-ratio was compared with the theoretical value:For(p-l,n -p), where a! is the chosen risk set in this study at 0.10.

Table 2 Principle of the Analysis of Variance

ss

Source

DF

Residual Total

n-l i= 1

MS

F ratio

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Forms

107

2. Multiple correlation coefficient (Rz). It gives the part of the variation explained by the model. It is calculated by using Eq. (15).

3. The lack of fit test which indicates if the lack of fit between the experimental values and the values calculated according to model equations can be explained by the experimental error. This test was performed exclusively on the Box and Wilson and the 33 experimental designs, because they were the only designs which needed to be subjected to this test according to the acceptability criteria described in Table 3. To perform this test, three additional nanoparticle batches with a formulation determined by the central point of the experimental designs were manufactured. The principle of this test is brieflydescribed in Table4. The experimental F-ratio obtained is compared with the theoretical value: Fa(n-p-c+l, c-l), where a is the chosen risk set in this study at 0.01. Further details should be sought in statistics textbooks. The Student t-test, was further applied for each term of the polynomial equations to evaluate theirsignificance. V.

EFFECTS OF THE CONTROLLED VARIABLES ON NANOPARTICLE FORMATION AND PROPERTIES

The controlled variables studied in this particular example were the amounIts of PLA, acetone, dichloromethane, indomethacin, and Poloxamer 188 used in the manufacturing process. As explained above, two sets of controlled variables were studied separately. The first set was composed of PLA, acetone, and dichloroTable 3 Acceptability Criteria Analysis of variance

Lack of fit

Correlation coefficient

R*2 0.9

9

P P

5

0.10

0.7 zs Rz <0.9

R’ <0.7 P >0.10

Model validation

2

0.01

P <0.01

9

YES YES

NO NO NO

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Table 4 Principle of the Lack of Fit Test Source

ss

DF

SSresidual-SSpurecmr

Lack of fit

iEB

MS

sslack of fit

n-p-c+l

n-p-c+l

F ratio

1

I-

MSlackoffit

MSpureemr

methane (PAD; see Fig. 2), and the secondset included dichloromethane, indomethacin, and Poloxamer188 (DIP; see Fig. 3). To better understand the experimental design approach, the results are presented and discussed separately for each set of data with a comprehensive evaluation in the conclusion section. The response variables evaluated included drug entrapment yield (yield), percentage of free drug (FD), mean particle size (MPS), standard deviation (SD), and coefficient of variation (CV). Since SD andCV provide similarly useful information regarding the polydispersity of the system, only thedataon CVis presented. Drugentrapment yieldis defined as the ratio between theincorporated and the total initial amount of drug. A drug entrapment yield of 100% corresponds to a drug content in nanoparticles of 5.88%. Particle size distribution was measured by photon correlation spectroscopy. In this example,the objective was to analyze the effects of the controlled variables ontheparametersthat characterize the nanoparticle colloidal system and to elucidate the nanoparticle formation mechanism.

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The importance of the drug entrapment yield for nanoparticlecharacterization is obvious and needs no further comment. From the percentage of free drug, itcan be deduced if low drug entrapmentyield is due to poor formation of nanoparticles or toineffective drug incorporation. The particle size distribution of the nanoparticles is one of the most important physical properties of a colloidal dispersion. Indeed, it is explicitly mentioned in the definition of nanoparticle. Sizing of particles in the suboptical range is associated with considerable difficulties, because the particle size value measured by the various sizing methodologies may be different. Thus, one single sizing procedure should be used throughout the entire set of experiments to avoid any misleading interpretation associated with the lack of consistency of the data obtained. Therefore, photon correlation spectroscopy (PCS), which is a convenient, fast, and well-established sizing technique in the submicrometer range,was carefully applied in this example while looking at thepolydispersity value of the measurements. Moreover, to rule out the possibility that small amounts of large particles might contaminate thecolloidal dispersion without being detected by PCS, all the batches were further examined by optical microscopy. From the overall results obtained during the preliminary investigation of nanoparticle formation process, the following underlying mechanism was proposed [58]. According to this mechanism, the mixing of the organic and the aqueous phases results in the diffusion of acetone from the organic to the aqueous phase. Part of the PLA which migrates together with the acetone on contact with the aqueousphase precipitates at theinterface owing to its insolubility in water. The remaining PLA isdissolvedin the dichloromethane. Following evaporation of organic solvents, the previously dissolved PLA precipitates thus forming the matrix of the nanoparticle in which the drug is entrapped. Such an hypothesis has also been proposed by other investigators [60,61]. A.

PLA-Acetone-Dichloromethane System

l. GeneralAnalysis In this example, the intent in the first part of the research was to find a generalmodel able to describe the complete system while varying all the parameters at thesame time. The datadid not fit the Hadamard matrix model (Table 5 ) , suggesting that quadratic effects and/or interactions between the controlled variables, undetectable by this model, may occur.

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Table 5 Statistical Validation ParametersValues Obtained for the PAD System

Yield

FD MPS CV

33

23

Hadamard

LOF -

P

R2

P

R2

P

RZ

P

0.454 0.623 0.316 0.303

0.867 0.737 0.937 0.943

0.499 0.755 0.136 0.209

0.933 0.712 0.958 0.935

0.005 0.017 0.000 0.584

0.732 0.672 0.799 0.350

0.030 0.409 0.267 0.157

The 23designwas then applied by adding only four experimental points to the Hadamard matrix. This design permits the postulation of an improved modelwhich has the possibility of detecting interactions between the controlled variables. The data did not fit this model as well (see n b l e 5 ) , reflecting the possible occurrence of quadratic effects unidentified by this model. In thelight of the potential presence of quadratic effects and interactions, a 33design, which allowsfor the postulation of a second-order model able to detect these phenomena, was applied. All the results obtained with the previous design form part of the new model emphasizingthe ability of saving experiments by the factorial design approach. The datafor the drug entrapment yield and the MPS fit this model with high significance (see Table 5). Owing to the numerous experimentalpoints forming the design, R2values were not very high, but the lack of fit test ascribes that to the experimental error,thus validating the model. It was observed that acetone andPLA have a significant influence on drug entrapment yieldwhich increases as the amounts of these factors increase (Table 6). The amount of acetone may affect its migration velocity during the initial formation process of the nanodroplets and subsequently the kinetics of PLA precipitation. PLA precipitation rate is concentration dependent. When morepolymer is present in the medium, the free volume available for it is reduced. Therefore, the possible conformationsthatthepolymermoleculescan adopt decrease, whereas the precipitation due to solubility constraints occur more readily. The deposition of the initial thin wall of PLA around the emulsified organic phase nanodroplets prevents further diffusion of the drug toward the aqueous medium. Hence, a faster PLA precipitation driven by higher amounts of acetone or PLA results in higher drug entrapment yield. The density of the polymer matrix formed increased with higher PLA concentrations resulting in an even higher yield.

111

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Table 6 Regression Coefficients and StatisticalValidation of the

Polynomial Individual Terms Obtained with the33 Model for thePAD System Drug entrapPercentage drugfreeyield ment Term

k PLA ACE DCM PLA*ACE PLA*DCM ACE*DCM PLA~

ACE^ DCM~ PLA*ACE*DCM

of

Mean particle size

RC

P

RC

P

RC

P

-16.27 1.87 9.49 103.69 -0.04 -2.61 -1.60 -0.01 -0.29 -131.37 0.18

0.681 0.089 0.084 0.312 0.399 0.133 0.848 0.211 0.201 0.156 0.267

7.77 -0.13 -0.53 -3.33 0.00 0.05 -0.26 0.00 0.01 16.03 -0.01

0.028 0.142 0.233 0.690 0.519 0.741 0.709 0.145 0.415 0.043 0.696

664.37 -6.59 -79.02 321.20 .0.19 6.22 -36.23 0.05 3.20 382.76 -0.44

0.008 0.273 0.015 0.574 0.505 0.513 0.447 0.342 0.020 0.452 0.615

The model describing the influence of the controlled variables on the percentage of free drug was validated by ANOVA (see Table 5), but the R2 value (0.672) was below the value set in the acceptability criteria (0.700; see Table 3). However, since the experimental R2 was so close to the preset value and the lack of fit test reached statistically acceptable levels, the effectof the controlled variables emerging from this model can be analyzed as a trend even though the model is not strictly validated. The quadraticinfluence of the amountof dichloromethane on the percentage of free drug (seeTable 6) is due toa more favorable drug partitioning into the organic phase when higher amounts of dichloromethane are present. The slight trend of effect of PLA (linear and quadratic; see Table 6) is related to the density of the polymer matrix formed, which affects the further partitioning of the drug between the nanoparticles and the aqueous phase. Acetone exhibits the greatest influence on the mean particle size, which decreases as the amountsof acetone increase (seeTable 6). A large concentration gradient of acetone acceleratesits migration to the aqueous phase during the initial formationprocess of the nanodroplets, and hence PLA precipitation occurs earlier, leading to smaller nanoparticles. It was not possible to describe the simultaneous effect of the controlled variables on the standarddeviation or thecoefficient of variation (see Table 5). This is apparently dueto thelack of accuracy of these specific measurements, as reflected by the low R' values.

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The factors that affect the drug entrapmentyield (acetone and PLA) are different from those affecting the percentage of freedrug (dichloromethane). This difference may be attributed to thedifferent mechanisms governing eachaspect of the overall process. In the formercase, the precipitation of the PLA promotes the formation of the embryonic nanoparticles. The extentof this precipitation, which is affected by the amounts of acetone and PLA, determines the amount of drug that will be entrapped. In the latter case, the drugpartition between the dichloromethane and the aqueous phase determines the extentof free drug in the aqueous phase.

2. ReducingApproach-PreciseAnalysis a. PLA and acetone variation at constant DCM levels. Drug entrapment yield dependsontheamount of acetoneat a level of 0.5 mL of dichloromethane and on PLA in the absence of dichloromethane (Table 8 and Fig.4a and b, respectively). It appearsthat the mechanism of nanoparticle formation is different in both cases. In thefirst case, a typical emulsion of dichloromethane in the aqueous phase is formed. In the second case, probably an organic acetonic phase rich in PLA was emulsifiedin the aqueous phase despite the good aqueous solubility of acetone. Therefore, in the first case, the rapid migration of the acetone is vital to the development of the first nanoemulsion, whereas in the second case, the rapid precipitation of the PLAis the key factor. No fitting to the drug entrapment yield and the percentage of free drug was observed for 0.25 mL of dichloromethane (Table 7). It appears that atthis intermediate dichloromethane level, various indefinite complex systems are obtained which are difficult tobe analyzedby one single model, and thus nomodel is validated for these response variables. The data of the percentage of free drug did not fit any model (see Table 7). Such behavior may be attributed tothe fixation of the 'dichloromethane level, which was the main factor affecting the free drug response variable. The models describing the effects of PLA and acetone on the mean particle size were validated for all the dichloromethanelevels (see Table 7). For thelevels of 0.00 and 0.25 mL, no termsof the mathematical equations were statistically significant (Table8). Therefore, nodefinitive conclusions . could be derived in these cases. The significant linear and quadraticeffects of acetone and the interaction between this solvent and PLA observed in the case of 0.50 mL of dichloromethane (Table 8) are due to the reasons explained above for the33model. It should be noted that the mean particle size of the nanoparticles obtained using 0.50 mL of dichloromethane were higher than those ob-

Factorial Design of Nanoparticulate Dosage Forms

_.-

113

- l\

9

6

(W

Fig. 4 Drug entrapmentyield as afunction of PLA and acetone amountin batches prepared using(a) 0.50 mL and (b) without dichloromethane.

Magenheim et al.

114

Table 7 Statistical Validation ParametersValues Obtained for the 3’ Models Corresponding to the Various ConstantLevels of Each Controlled Variable of the PAD System ~

~~

~~

Fixed variable

Level

Test

Yield

FD

MPS

CV

Dichloromethane

0.00 mL

P R‘ P R’ P R’ P R’ P

0.062 0.927 0.552 0.614 0.014 0.974 0.212 0.824 0.165 0.854 0.992 0.110 0.361 0.733 0.132 0.876 0.226 0.815

0.121 0.884 0.801 0.424 0.107 0.894 0.304 0.767 0.112 0.890 0.059 0.930 0.234 0.811 0.158 0.859 0.513 0.639

0.031 0.955 0.081 0.912 0.003 0.990 0.001 0.996 0.042 0.945 0.100 0.899 0.111 0.891 0.145 0.867 0.109 0.941

0.380 0.722 0.003 0.992 0.479 0.661 0.542 0.621 0.891 0.327 0.701 0.508 0.404 0.707 0.350 0.740 0.285 0.780

0.25 mL 0.50 mL

Acetone

5 mL

10 mL

R2 15 mL

P R2

PLA

25 mg 50 mg

75 mg

P R’ P R’ P R’

tained without dichloromethane (Fig. 5c and a, respectively). Mean particle size depends strongly on the amount of acetone at a level of 0.5 mL of dichloromethane and is almost independent from it in the absence of dichloromethane(see Table S). This difference is attributed to different mechanisms of nanoparticle formationas described above. The profiles of the graphs depicting the influence of acetone andPLA on the mean particle size of the nanoparticles obtained using 0.25 mL of dichloromethane are almost identical to those obtained without dichloromethane (Fig. 5b and a,respectively). This is mainlydue to thesolubility of dichloromethane in water at a low concentration.

b. DCM and acetone variation at constant P L A levels and P L A and DCM variation at constant acetone levels. PLAandacetone play primordial roles in the formation process of nanoparticles. Therefore, with a single exception, novalidated models for thedrug entrapment yield and percentage of free drug were achieved when either PLA or acetone were kept constant while the remaining parameters were varied (see Table 7). Furthermore, as previously shown, at intermediate DCM concentrationscomplex colloidal systems were obtained. Thus,in anycase where the DCM concen-

a,

U d

a,

Factorial Design of Nanoparticulate Dosage Forms

E

0

el

2

E

el m

a,

2 "d E

0

cl

4

8 "d E

cl 0

p!

2 c

E

cl 0

8

115

116

Magenheim et al.

Fig. 5 Meannanoparticlesizeasafunction

of PLA and acetone amountin batches prepared using (a) 0.00, (b) 0.25, and (c) 0.50 mL of dichloromethane. (From ref. 58.)

tration is varied, as required when PLA or acetone are kept constant, the experimental field will include a range of concentrations leading to indefinite colloidal systems which are unlikely to be fully characterized. With acetone as the major factor affecting the particle size when its value is fixed at some level, the variation of the other factors is unable to alter the particle size distribution that remained almost unchanged over a reasonable concentration variation range (Table 9 and Fig. 6a-c). At high PLA and low acetone concentrations, a dramatic increase in MPS was noted (Figure 6a) which can be attributed to thehuge increase in viscosity of the organic phase and thus preventing the formationof optimal nanoparticles. It was not possible to describe the simultaneous effect of the controlled variables on the standard deviation or the coefficient of variation (see Table 7). This was apparently due to the lack of accuracy of these specific measurements, as reflected by the low R’ values. No model could be validated in any of the levels of PLA for the particle size (see Table 7). c.

Interbatch variability. The nanoparticles prepared using 0.25 mL DCM, 25 mg PLA, and 5 mL acetone, exhibited poor interbatchparticle size measurements’ reproducibility. Some of the batches obtained were within the range of data values previously described, whereas others exhibited a

Factorial Design of Nanoparticulate Dosage Forms

117

et

118

Magenheim

Table 9 Regression Coefficients and Statistical Validation of the Polynomial Individual Terms Obtained in the Mean Particle Size Equation for the Dichloromethane and PLA Variation at Constant Acetone Levels in the PAD System 5 mL

RC

Term k DCM PLA DCM*PLA DCM* PLA~

212.57 -805.20 -2.72 4.86 2361.60 0.04

10 mL

15 mL

P

RC

P

RC

P

0.025 0.008 0.296 0.047 0.001 0.158

128.06 -95.33 -0.28 0.12 314.67 0.01

0.016 0.253 0.818 0.884 0.061 0.426

100.39 -124.00 0.19 0.44 274.67 0.01

0.069 0.278 0.909 0.705 0.163 0.685

al.

Factorial .Designof Nanoparticulate Dosage Forms

119

120al.

et

Magenheim

much larger mean particle size and polydispersed population distribution. Therefore, two independent analyses were carried out, one foreach of the nanoparticle populations. The results reported are for the reproducible type of populations. For the larger and heterogeneous populations,no general models could be validated, and therefore the results are not presented. Apparently theconcentration of the formulation components in this special combination led to complex instability problems which resulted in insufficient control of the nanoparticle manufacturing process. B. Dlchloromethane-Indomethacin-Poloxamer188 System

The data did not fit the Hadamard matrix and the 23model for any of the responses variables (Table 10). Only the Box and Wilson composite design was able todescribe the behavior of the system, accounting for the complex nature of the physicochemical phenomena involved in the formation of nanoparticles (Table 10). The decreasein the drug entrapmentyield noted while increasing the DCM concentration at low Poloxamer 188 levels (see Fig. 7a) can be attributed to therelative decrease of the PLAconcentration in the DCM phase. This resulted in a slower PLA precipitation and the formation of a less dense matrix, which allowed for a larger diffusion of the drug toward the aqueous phase. The decrease in drug entrapment yield along with the Poloxamer 188 concentration increase at low DCM levels (see Fig. 7a) is apparently due to an increase in drug partitioning. This is supported by the increase in the free drug along with the increase in Poloxamer 188 concentration (see Fig. 7b). The synergistic effect of the simultaneous increase of DCM and Poloxamer 188 on drug entrapment yield is clearly observed in Fig. 7a, accounting for the validation of the interaction term. The moderate increase in the drug entrapment yield observed while increasing the Poloxamer 188 concentration at high DCM levels is speculated to be caused by the migration of part of the excess DCM toward the Table 10 Statistical Validation ParameterValues Obtained for the DIP System 23

Hadamard

Yield

FD MPS CV

Box and Wilson

LOF -

P

R2

P

R2

P

R2

P

0.484 0.297 0.401 0.134

0.848 0.944 0.897 0.989

0.636 0.257 0.538 0.131

0.881 0.984 0.920 0.996

0.062 0.012 0.114 0.299

0.812 0.890 0.767 0.661

0.894 0.090 0.020 0.994

Factorial Design of Nanoparticulate Dosage Forms

121

aqueous phase promotedby the solubilizing power of high Poloxamer 188 concentrations. In this case, the actual PLA concentration in the DCM increases, resulting in better drug entrapment. The extentof free drug is controlled by the solubilization of the drug in aqueous phase, which increases as the Poloxamer 188 concentration increases (Table 11 and see Fig. 7b). The mean particle size increases at higher DCM concentrations because of the increase in the incipient nanoparticle volume (Tablell and see Fig. 7c). Poloxamer 188 does notaffect the particle size, indicating that low concentrations are enoughto stabilize the system (Table11and see Fig. 7c). C. Conclusion on the Application of Experimental Design to Nanoparticle Formation Process

The overall results presented in this example are in agreement with the mechanism of nanoparticle formation process that has been previously suggested (see Fig. 8). It was clearly observed throughthe factorial design approach that nanoparticle formation process is very fast, and hence the manufacturing conditions immediately after the aqueous andorganic phase mixing determine thephysicochemical properties of the nanoparticles. Only the second-order mathematical models derived from the Box and Wilson and 33experimental designs were able to describe the behavior of the system and account for the complex nature of the physicochemical phenomena involved in the formationof nanoparticles. The migration velocity of the acetone and the kinetics of PLA precipitation are the most significant parameters in nanoparticle formation. The former particularly affects the mean particle size, whereas both have an important influence on the drug entrapment yield. The mean particle size is also affected by the amount of dichloromethane, since it determines the volume of the incipient nanoparticles. The extentof free drugis controlled by a phase partition phenomena which depends fundamentally on the concentrations of dichloromethane and Poloxamer188 in the organic and aqueousphases, respectively. The rapidity of the nanoparticle formation process is also supported by other independent observations demonstrating that no difference was observed in particle diameter before and after evaporation of acetone. This may reflectthe fast and completeprecipitation of the polymerin the course of nanoparticle manufacturing process. The kinetics of PLA precipitation are highly dependent onits ratio to dichloromethane. The lack of significance of the interactions between acetone and dichloromethane suggests that both solvents behave independently andthat acetone is not retained by the dichloromethanein its migration toward the aqueous phase.

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Table 11 Regression Coefficients and Statistical Validationof the Polynomial Box and Wilson Composite Design for the Individual Terms Obtained with the DIP System Drug entrapment Percentage yield size particle Mean drug free

of

Term

RC

P

RC

P

RC

P

k DCM

56.71 180.59 0.53 -10.70 -24.33 46.12 -1.57 -65.29 -3.92 0.97

0.558 0.300 0.978 0.581 0.049 0.031 0.389 0.578 0.247 0.417

4.62 67.64 -4.49 -14.41 -2.38 6.65 0.76 40.57 0.31 3.26

0.887 0.258 0.491 0.056 0.517 0.290 0.230 0.322 0.447 0.018

353.38 -1421.61 52.32 60.00 -8.80 -57.33 -0.67 1331.75 -2.88 -9.89

0.312 0.041 0.440 0.387 0.814 0.372 0.915 0.012 0.490 0.396

IND

POL DCM*IND DCM*POL IND*POL DCM~ IND2

POL2

-

e

Fig. 7 (a) Drug entrapment yield, (b) percentage of free drug, and (c) mean particle sue as a function of poloxamer188 and dichloromethane concentration in batches prepared using indomethacin7.5% wlv.

Factorial Design of Nanopatticulate Dosage Forms

'.

123

124

Magenheim et al. Dichloromethane Acetone PLA Drug

-b

Dichloromethane PLA Acetone Drug

IC

Poloxamer Water

n Drug

Dichloromethane

Drug

Dichloromethane

Poloxamer POIOXBIIWH

Water

Polo Poloramer

Acetone Drug Acetone Poloramer

Water

Poloramer

Fig. 8 Schematic descriptionof the proposed nanoparticle manufacturingmechanism. (From ref. 58.) The nanoparticle systems obtained at the various levels of dichloromethane and their mechanism of formation appear to be different. At a level of 0.5 mL of dichloromethane, a typical emulsion of dichloromethane in the aqueous phase is initially formed, whereas in the absence of dichloromethane, an acetonic phase rich in PLA, insoluble in the aqueous phase, constitutes the internal dispersed phase of the emulsion. In the former case, the rapid migration of the acetone is critical for the development of the first nanoemulsion, whereas in the latter case, the rapid precipitation of the PLAfrom the acetonic phase is the key factor. VI. SUMMARY The use of factorial design in investigating the nanoparticle formation process clearly demonstrates the advantage of this efficient statistical technique in characterizing complex physical systems containing various controlled variables with only a minimal number of experients needing to be performed. Time andfastidious work are saved by this valuable tool which uses the sequential approach to identify the potentially dominant parameters of the studied system. It should be emphasized that the experimental design not only offers the potential of reducing the number of experiments needed to optimize a manufacturing process but may also contribute to a better physicochemical understanding of the phenomena involved in the

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construction of a specific formulation aimed at resolving a pharmaceutical formulative problem. It is well know that in the pharmaceutical manufacturing processes many important parameters exist owing to the large number of excipients usually required for stable and viable formulations. Thus, investigators cannot andshould not bypass, avoid, or neglect the optimization process of the formulations. Failure to optimize the process of the formulations can lead to problems with safety, regulatory bodies, therapeutic use, and economics. Modern pharmaceutical formulation design requires the use of efficient, comprehensive, informative, and validated techniques that can be exploited to gain physicochemical insights into the underlying complex phenomena generally involved in the formulation of any pharmaceutical dosage form. The factorial design presented in this chapter and applied to optimize the nanoparticle manufacturing process as well as elucidate the mechanism of nanoparticle formation appears to meet the various and complex requirements needed for a potent technique to achieve a global and comprehensive characterization of the drugformulation and manufacturing process.

VII. SYMBOLS PAD DIP PLA ACE DCM IND POL RC

PLA-acetone-dichloromethane dichloromethane-indomethacinAPoloxamer188 amount of polylactic acid

amount of acetone amount of dichloromethane percentage of indomethacin percentage of Poloxamer 188 regression coefficient ss sum of squares DF degrees of freedom MS mean of squares n total numberof experiments P number of terms of the model (constant included) Yi measured response Yi estimated response Y mean response B group of the experimental points, central points excluded C group of the central points LOF lack of fit Yield drug entrapmentyield

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FD percentage of free drug MPS CV

mean particle size coefficient of variation

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isosorbide-5-mononitratemicrocapsules, J. optimization of the formulation of Microencaps. 10:309 (1993). J. M. Pernot, H. Brun, B. Pouyet, M. Sergent, and R. Phan-Tan-Luu, Influence of experimental parameters on the microencapsulation of a photopolymerizable phase, J. Microencaps. 10:323 (1993). X.M. Zeng, G.P. Martin, and C. Marriott, Albumin microspheres as a means of drug delivery to the lung: analysis of the effects of process variables on particlesizesusingfactorialdesignmethodology,Int.J.Pharm.,107:205 (1994). X. M. Zeng, G . P. Martin, and C. Marriott, Tetrandrine deliveryto the lung: The optimisation of albumin microsphere preparation by central composite design, Int. J. Pharm., 109:135 (1994). M. J. Alonso, A. Sanchez, D. Torres, B. Seijo, and J. J. Vila, Joint effects of monomer and stabilizer concentrationson physico-chemical characteristics of poly(butyl2-cyanoacrylate)nanoparticles, J. Microencaps.,7517 (1990). R. Carpignano, M. R. Gasco, and S. Morel, Optimization of doxorubicine incorporation and of the yield of polybutylcyanacrylate nanoparticles, Pharm. Acta Helv., 66:28 (1991). M. C. Julienne, M. J. Alonso, J. L. Gomez Amoza, and J.P. Benoit, Preparation of poly (D,L-Lactide/glycolide) nanoparticles of controlled particle size distribution: Application of experimental designs, Drug Dev. Ind. Pharm., 18:1063 (1992). G. E. BoxandN.R. Draper, in Empirical Model Building and Response Surface, ( G . E. Box and N. R. Draper, eds.),Wiley, New York, 1987, p. 205. G. E. Box, W. G . Hunter, and Hunter. J. S., Response surface methods, in Statistics for Experimenters (G. E. Box, W. G. Hunter, and Hunter. J. S., eds.), Wiley, New York, 1978, p. 510. 0.L. Davies, The determinationof optimum conditions, in The Design and (0. L. Davies,eds.),Longman, New AnalysisofIndustrialExperiments York, 1978, p. 495. WehrlC (1990) Aspects des analyses multifactorielles et des plans d’experiences appliquCs h l’optimisationet h lavalidationdeformes et deprocedes galeniques. Ph.D. thesis No. 149, University of Paris XI. G. Sad0 and M. C. Sado, Notions de plan optimal et critkres d’optimalitk, in Les plans d’experiences, AFNOR, Paris, 1991, p. 10.

Microsphere Morphology Jean-Pierre Benoit Laboratoire de Pharmacie Galdnique et Biophysique Pharmaceutique, Universitd &Angers,and Centre de Microencapsulation, Angers, France

Curt Thies Washington University, St. Louis, Missouri

I. Introduction 11. Progesterone-Loaded Poly(d,l-Lactide) Microspheres

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INTRODUCTION

Many drug-loaded microspheres are prepared by processes that involve intimate mixing of a polymeric carrier with a drug. This creates a situation where a variety of physicochemical interactions occur, and these interactions can influence therapeutic performance of the final device. Consequently, the morphology of drug-loaded polymer microspheres warrants careful study. The purpose of this chapter is to illustrate how the morphology of biodegradable microspheres prepared by removal of avolatile 133

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solvent from oil-in-water or water-in-oil-in-water emulsions canbe characterized. In such processes [1,2], a polymer carrier is dissolved ina volatile solvent like methylene chloride (CH,Cl,). A drug is then either dissolved or suspended in the solution and the resulting mixture is emulsified in an aqueousphasethat contains an emulsifier. The solvent is allowed to evaporate, thereby giving drug-loaded microspheres. Although the procedure is conceptually simple, it is not in practice. A number of facton affect the natureof the microspheres obtained. Theseinclude solubility of the drug in the casting solvent and aqueous phase, drug-polymer ratio, nature of the polymer used as the carrier, and aqueous phase emulsifying agent. Crystalline drugs that are freely soluble in the casting solvent may exist in the final microsphere as a molecular dispersion, or they may form well-defined crystal domains in the polymer matrix. What state a drug assumes in a microsphere is defined by the drug-polymer interaction and percent loading of drug in the final microsphere. Residual casting solvent is another factor, since it could prevent drug crystallization if present in the final microsphere. When a crystalline drug is soluble in the casting solvent, its ability to crystallize in a microsphere is repressed at low drug loadings because of the nature of the solvent evaporation process. The drug-polymer-solvent solution used to form the microspheres is initiallya dilute solution of relatively low viscosity. When it is emulsified in an aqueous phase, the volatile solvent begins to partition into the aqueous phase from which it is removed by evaporation. As this process progresses, the viscosity of the organic phase increases steadily. For polymers with a glass transition temperature (Tg) above the temperature at which evaporation occurs, the viscosity of the polymer-containing organic phase increases to infinity as the volatile solvent concentration decreases toward zero. If the drug-polymer-solvent phase is sufficiently viscous when the saturation solubility of drug in the solvent is reached, and if the rate at which the remaining solvent evaporates is sufficientlyhigh, drug crystallization in the viscous polymer matrix is effectively prevented. In this manner, drug thathas little mutual miscibility with the polymer can be molecularly dispersed in the polymer. The dispersion is a metastable state formed, because the drug does not have time to crystallize before the polymer phasesets up as an organic glass. If the drug and the polymer are mutually miscible, a true molecular solution of drug in the polymer is formed, like a solution of a plasticizer in a polymer. Cases of partial drug-polymer miscibility fall between these extremes. With drugs that are weakly soluble in the volatile solvent, formation of well-defined crystalline domains occurs readily. The above list of possibilities illustrates why a drug incorporated in a

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polymeric microspheremay exist there in a wide variety of physical states. It is appropriate toillustrate some of these possibilities with specificexamples of drug-loadedmicrospheres fabricated from lactide (PLA)and lactide-glycolide (PLAGA) polymers. II. PROGESTERONE-LOADEDPOLY(d,l-LACTIDE) MICROSPHERES

Fig. 1contains SEM photomicrographs thatshow howthe surface structure of 100-300 pm PLAmicrospheres varies with progesterone loading [3]. At 23 w t . percent loading, the microsphere surface is perfectly smooth. At 35 wt. percent loading, a few crystals are embedded in the surface. At 68 wt. percent loading, the microspheres are coated by a continuous layer of progesterone crystals. The formation of free drugcrystals on thesurface of microspheres or in the aqueous phase as solvent evaporation progresses is a problem often encountered in the solvent evaporation process when the drug used is soluble in the casting solvent. Free crystal formation can be repressed if the aqueous phase emulsifier is removed from the system before solvent evaporation is complete [3,4]. Solvent evaporation is then taken tocompletion in emulsifier-free water. Fig. 2, obtained by differential thermal analysis (DTA), illustrates the thermal behaviorof a PLA microsphere sample loaded with 23wt. percent progesterone [3]. From these scans, a number of conclusions can be made. First, there is no endothermic peakat 120-132°C characteristic of the a or p polymorphs of progesterone. The progesterone is molecularly dispersed in the microsphere sample and does not form macroscopic progesterone crystal domains that can be detected by DTA. The x-ray powder diagramof this sample is a diffuse halo, thereby confirming that well-defined progesterone crystal domains are not present. Since no crystalline progesterone is present, a key question is whether or not the progesterone and PLA form a molecular solution much like a plasticizer and polymer. This question can be answered by prolonged annealing experiments carried out well above the Tgof PLA but well below the melting point (Tm) of progesterone. The logic for this experiment is as follows. Drug molecules trapped as a molecular dispersion in a polymer matrix at temperatures below the polymer’s Tg are locked in a polymer glass and cannot diffuse in a finite time to nucleate and/or grow finite domains of crystalline drug. When the polymer is heated above its Tg, chain mobility is greatly increased, thereby greatly increasing diffusivity of the drug in the polymer matrix and enabling it to crystallize if it wishes. Crystallization will occur if the polymer and drug are mutually immiscible. If they are mutually miscible, there is no crystallization. It is assumed that

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Fig. 1 Scanning electron micrographs of poly(d,l-lactide) microspheres containing (A) 23%, (B) 35%, (C) 68% progesterone.

mutual miscibility of the drug and polymer does not change measurably with temperature over the temperature range of interest. It also is assumed that the drugis not degraded measurablyduring the annealing process. In order tominimize the possibility of degradation, all annealing experiments are carried out in vacuo. Fig. 3 contains DTA scans for an annealed progesterone-loadedPLA microsphere sample [3]. Annealing was done for 22 h at 110°C. The annealed samples were quench cooled in liquid nitrogen and their thermal behavior was evaluated after a known aging cycle.All of the DTA scans in Fig. 3 have a well-defined endotherm at 130°C due to the melting of CYprogesterone crystal domains. This is evidence that PLA and progesterone are immiscible with each other at a progesterone loading of 23%. The progesterone is initially present as a molecular dispersion. This metastable dispersion existsin thePLA glass atroomtemperature,becausethe

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progesterone molecules did not have time to crystallize in the polymer matrix as CH,Cl, evaporation progressed to completion andthe PLA set up to a glassy state. When given the opportunity, it crystallized. Most of the crystal domainsformed in PLA by annealing at 110°C were the apolymorph (130°C Tm). Occasionally, a melting event at 120°C was observed. In this case, P-progesterone domains formed. A DSC scan of the freshly annealed sample used to generate Fig. 3 showed that an estimated 80% of the progesterone carried by the PLAmatrix had crystallized during the 110°C annealing process. The melting endotherm had a maximum at 130°C but was broad and skewed toward 120°C. After 3 and 7 days of storage at 22"C, the endotherm became much sharper and was still centered at130°C. This change in the endotherm onaging caused the apparent progesterone crystallinity calculated from the DSC scan to fall to 55% and suggests that at least some p-progesterone crystal domains were initially present in the PLA matrix. They either were transformed onstorage into a-progesterone crystal domains or simply dispersed in the PLA matrix. Fig.3shows that the nature of the PLA Tg event for the 110°C annealed microsphere sample also changes onstorage at 22°C. Initially, it is a rather diffuse event located at 49°C. After 3 days at 22"C, it becomes more visible and shifts upward to 51°C. After 14 days at 22"C, it is a welldefined event located at 53°C with an endothermic peak. The type of aging shown by the PLATg event in Fig. 3 is commonly observed with amorphous glasses stored close to but below their Tg [5]. Some workers attribute this to development of molecular order in the

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polymer glass, whereas others argue thatthis phenomenon is due torelaxation of stresses and strains arising from nonequilibrium cooling. Petrie [5] gives a good overview of the subject from the latter viewpoint. Regardless of theorigin of this aging behavior, it is relevant to note from Fig. 2that the unannealed progesterone-loaded PLA microsphere sample retained a comparatively diffuse Tg event at 37°C with no endothermic peak when aged 83 days at 22°C. In contrast, theannealed sample gave a Tg event thatrapidly developed an endothermic peak on storage at 22°C and fell at 49-53"C, close to theTg of drug-free PLA. The difference in aging of the PLA Tg event of drug-free and drugloaded microspheres is related to residual solvent content of the microspheres and/or degree of molecular dispersion of progesterone in the PLA microspheres before heat treatment.Although all PLA microspheres appeared to bethoroughly dry after vacuum drying at 22°C for atleast 20 h, they actually still contained between 1 and 5% CH,Cl,, the solvent from which the PLA microspheres were cast [3]. The low PLA Tg values shown in Fig. 2 could be caused solely by residual solvent trapped in the microspheres that acts as a plasticizer, thereby lowering the Tg of PLA and making it more diffuse. However, drug-free PLA microspheres develop a well-defined 57°CTg eventwith an endothermic peak within 70 days of22°C aging. This suggests that the diffuse and low PLA Tg event shown in Fig. 2 is not due solely to residual

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CH2Cl, but is caused, at least in part, by interference of molecularly dispersed progesteronewith the relaxation or ordering phenomenon responsible for developmentof a well-defined PLA Tg event by progesterone. The effect of molecularly dispersed drug on the behavior of a polymer’s Tg event merits further investigation. When progesterone is molecularly dispersed in PLA, it appears tobe vulnerable to degradation. The progesterone carriedby two PLA samples loaded with 23% progesterone and two PLA samples loaded with 31.7% progesterone experienced 24-47% degradation when stored 16months in a desiccator at 25°C [6]. All four samples contained only molecularly dispersed progesterone. Thesamples loaded with 31.7% progesterone turned yellow during the aging process. The nature of the degradation products was not determinednor was the origin of the degradation process. Residual CH,CI, could have beenacontributing factor. The key point is that progesterone molecules present as a molecular dispersion in a PLA matrix are more susceptible to degradative attack than if they are located in crystalline progesterone domains. 111.

PROGESTERONE-LOADED85/15 POLY(d,l-LACTIDEGLYCOLIDE) COPOLYMER MICROSPHERES

A series of 85/15 (w/w) PLAGA copolymer microspheres (25-50 wm in diameter) was prepared by the solvent evaporation process with CH,Cl, as the casting solvent [7]. Progesterone loading ranged from 0 to 64.6 wt. percent. As in the case of progesterone-loaded PLA microspheres, the external appearanceof the copolymer microspheres varied with drug loading; however, thevariation was morepronounced.Copolymer microspheres with 10 wt. percent progesterone loading had a smooth external surface. Increasing theloading to 28 wt. percent caused the surface of the microspheres to become serated and some free crystals embedded in the surface were visible. At 50 wt. percent loading, the microspheres were porous with progesterone crystals visible inthe surface. Table 1lists DSCthermal event temperatures for progesterone-loaded PLAGA copolymer microspheres [7]. These data illustrate a number of features ofsuch microspheres. Column 7 shows that samples with 22.8-64.6 wt. percent progesterone containcrystalline progesterone domains regardless of their thermalhistory. A sample loaded with 10.9% progesterone did not developcrystalline progesterone domains until it was annealed at80°C for 24 h. Although some freshly made samples contain a mixture of aprogesterone (130°C Tm) and P-progesterone (120°C Tm), the p form always predominates.This is interesting because progesteronesolutions evaporated to dryness from CH,C12 always yield only a-progesterone. In

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most cases, crystalline progesterone domains that form in a PLAmatrix also are the a-polymorph.Crystallization in the copolymer matrix favors formation of the p-polymorph. Samples with 22.8 and 34.4 wt. percent progesterone develop nearly all of their crystallinityspontaneously during the DSC heating cycle. Before heating,most of the progesterone is molecularly dispersed in the copolymer matrix. This crystal growth creates the exothermic peak at 80-115°C (column 6 ) . Muramatsu et al. [8], reported that a molten progesterone specimen cooled at 10"C/min was cooled so fast that it remained supercooled throughout the cooling process. Crystallization in the p form always occurred between 37 and 120°C when the supercooled specimen was reheated. Accordingly, the behavior of microsphere samples with22.8 and 34.4% progesterone parallels that of supercooled progesterone when it is reheated. In the case of the microspheres, the equivalent to a supercooled specimen was achieved at essentially constant temperatureby carrying out solvent evaporation in the copolymer matrix. As CH,Cl, evaporation progressed to completion, theviscosity of the copolymer phase increased dramatically, thereby effectively enabling the progesterone to act as a supercooled specimen and hence remain molecularly dispersed. On heating above Tg of the copolymer, the diffusivity of progesterone in the PLAGA matrix increases greatly. It is free tocrystallize and does so in essentially the p-form like a supercooled progesteronesample. Another interesting observation is the change in relative proportion of a- and P-progesteronethat occurs on aging. Theproportion of aprogesterone increases when the copolymer samples are aged at 4°C or annealed at 80°C. This is not surprising, since the a-polymorphis the more stable form.Nevertheless, it is interesting thatthis transition proceedsto a measurable extentduring 9-12 months of storage at 4"C, well belowthe Tg of the copolymer. Muramatsu et al. [ 8 ] , reported that crystals of a-and pprogesterone seemed tobe stable atroom temperature for several months or longer. As noted in Section 11, the natureof the crystalline progesterone event in an annealed progesterone-loaded (23%) PLAsample also changed on storage. In this case, the change appeared to be complete in 7 days at 22°C. These data are evidence that the p-ta transition of progesterone located in a solvent-cast polymer matrix proceeds more easily than in progesterone crystals grown in the absence of a polymer. The consistent development of crystalline progesterone domainsin a range of PLAGA copolymer microsphere samples is evidence that progesterone and the copolymer have at best a limited degree of mutual miscibility. When given an opportunity to crystallize, progesterone that initially exists as a molecular dispersion in a PLAGAmatrix crystallizes. This behavior is analogous to that observed with PLA homopolymer. For both poly-

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mers, the solvent evaporation process can be manipulated to produce microspheres that contain molecularly dispersed progesterone, provided the loading is low (e.g., below 30%), but this is a metastable state. When the samples areheated, progesterone crystallization occurs spontaneously. Why crystallization of progesterone in the lactide-glycolide copolymer matrix yields primarily the p polymorph and crystallization in a PLA matrix yields primarily the a polymorph remainsunresolved. However, it is clear that the polymermatrix affects what crystalpolymorph forms. The copolymer Tg data in column 5 of Table 1 provide further evidence that progesterone and lactide-glycolide copolymer are essentially immiscible [7]. If they were mutually miscible, a change in copolymer Tg proportional to progesterone loading would be expected. This does not occur. Several progesterone-loaded copolymer samples stored at4°C for a prolonged period have a copolymer Tgof 48-49"C, which is essentially the same as that of the copolymer before it was fabricated into microspheres. All samples annealedat 80°C for 24 h havea 47-48°C copolymer Tg. Most of the samples with low copolymer Tg values listed in Table 1were evaluated soon after manufacture.Such samples often had an endotherm at6580°C (column 4) associated with volatilization of residual CH,Cl, from the microspheres as they were heated. When heated, these samples lose 2.54.6% of their weight owingto CH,Cl, volatilization. When the samples are aged 9-12 months at4"C, the endotherm dueto CH,Cl, volatilization disappears (column 4), sincemuch of the residual CH,Cl, diffuses from the microspheres during storage. Column 3 of Table 1 shows that the residual 1%. Annealing CH,Cl, content of three samples aged at 4°C was well below for 24 h at 80°C also removes residual CH,Cl, from the polymer matrix. Thus, the low copolymer Tg values recorded in Table 1 are attributed primarily to residual CH,Cl,. Low PLAGA Tg values stored 12 months at 4°C are attributed toslow equilibration of the polymer matrix. The boiling point of CH,Cl, is 4O-4l0C, whereas the CH,Cl, evaporation endotherm for the microspheresis located between 64 and 80°C. This suggests that CH,Cl, interacts strongly withthe lactide-glycolide copolymer and is difficult to remove completely from a matrix of such a copolymer. Persistent retention ofCH,Cl,by PLA microspheres also has been observed. The question of solvent retention by microspheres formed by the solvent evaporation process and intended for injectable drug formulations warrants much closer scrutiny.A numberof papers have beenpublished on the in vivo performance of injected lactide and PLAGA microsphere samples, but the residual solvent content of the injected microspheres is never given. Although such microspheres may appear to be"dry" after anextensive drying process, they still may contain sufficient residual casting solvent (e.g., CH,Cl,) to have a meaningful effect on microsphere morphology,

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drug stability in the microsphere, and drug-release rate from the microsphere. The matter of how residual solvent affects acute and chronic toxicology of the injectable microspheres must alsobe considered. Because of the tenacity with which lactide and lactide-glycolide copolymers hold solvent, published studies of the in vivobehavior of microspheres prepared from such polymers should specify the residual solvent content carried by the microspheres andwhat the solvent@)is. Before concluding this section, it is relevant to mention that the Tg event of the PLAGA experiences an agingeffect analogous to that of PLA. The Tg event becomes more well defined on aging and develops a significant endothermic peak. The rateof development of this peak increases as the aging temperature approachesTg of the copolymer[7]. IV.

HYDROCORTISONE-LOADEDPOLY(d,l-LACTIDE) MICROSPHERES

A series of 50- to 150-pm diameter PLA microspheres loaded with 4.247.0 wt. percent hydrocortisone was prepared by continuous evaporation ofCH,Cl,[9]. Hydrocortisone has finite solubility in water and is only partially soluble in CH,Cl,. Up to hydrocortisone loadings of 32 wt. percent, partially hydrolyzed poly(viny1 alcohol) (PVA) gave stable emulsions from which spherical microspheres were prepared. At higher hydrocorwhich microspheres tisone loadings, PVA did not give stable emulsion from could be made, whereasmethylcellulose did. Accordingly, allmicrospheres loaded with 32 or more percent of hydrocortisone weremade with . methylcellulose as the aqueousphase emulsifier. A PLA microsphere sample loaded with 12.6% hydrocortisone stored at 22°C for 14 weeks had one well-defined endotherm at56°Cand a second at 208-209°C [9]. The 56°C event is characteristic of drug-free PLA and is due to theglass-transition event of PLA. This eventdid not have an endothermic peak when the microspheres were freshly made. The peak developed on storage of the microspheres at 22°C and appears to be sharper and larger than similar endothermicpeaksdeveloped by aged PLA and lactideglycolide copolymer samples loaded with progesterone. The 208-209°C endotherm that appears in the DSC scan of PLA microspheres loaded with 12.6% hydrocortisone issmall. It disappears when the microspheres are heated to 250"C, quench cooled, and reheated. The x-ray powder diagram of such microspheres taken before they were heated had d-spacings characteristic of crystalline hydrocortisone. Thus, the event is due to the melting of crystalline hydrocortisone domains located in the microspheres. The 208-209°C fusion temperature is well below the 228°C fusion temperature found for well-defined hydrocortisone crys-

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tals. This suggests that thecrystal domains in the microspheres are not well defined. Since the event is so small, much of the hydrocortisone exists in the polymer matrix as a molecular dispersion. This observation was unexpected because the initial PLA-solvent-steroid mixture used to form the microspheres was turbid and contained undissolved steroid crystals. It was thought that these would be trapped in the final microspheres to give a significant fusion event at 228°C. Direct microscopic observation throughout the solvent evaporation process revealed that this did not occur, because the steroid crystals initially present in the CH,Cl, phase migrated spontaneously into the aqueous phase on emulsification. The crystalline steroid domains in the isolated PLA microspheres were formed by crystallization of solubilized drug that occurred as solvent evaporation progressed. Since the viscosity of the PLA-steroid-solvent solution increased steadily as solvent evaporation progressed, the steroid crystal domains nucleated and grew in a progressively more viscous medium. This was assumed to hinder the development of defect-free crystal domains and the extent of steroid crystallization. Microspheres with larger steroid contents would undoubtedly have a more pronounced steroid fusion event which would fall closer to 228°C. In such cases, the crystallization would occur sooner in the process owing to thelarger amount of drug present. The study of hydrocortisone-loaded PLA microspheres providedanother illustration of how tenaciouslyPLA binds CH,Cl,. The sample loaded with 12.6% hydrocortisone had a 2.74% chlorine content after it was dried in vacuo at 22°C for 24 h. Extending the drying time to 72 h reduced the chlorine content to2.56%. Assuming these chlorine contents are dueexclusively to residual CH,Cl,, it is calculated that the microspheres contained 3.8% solvent after 22 h drying and 3.3% after 72 h drying. Increasing the drying time from 22 to 72 h had relatively little effect on residual CH,Cl, content. The hydrocortisone in PLA microspheres does not appearto degrade rapidly. Samples stored for 55 days at 25 or 37°C experienced no meaningful degradation [9]. V.

LOMUSTINE-LOADEDPOLY(d,l-LACTIDE) MICROSPHERES

Lomustine, 1-(2-chloroethyl l-3-cyclohexyl-l-nitrosourea (CCNU), is well known for its efficacy on tumorsof the central nervous system. A promising way of optimizing the action of CCNU is to incorporate it into microspheres and target the microspheresto tissue sites via selective arterial catheterization. For this reason, an effort has been madeto develop injectable CCNUloaded microspheres. Although the development process is complicated by the reactivity and comparative instability of CCNU, CCNU-loaded mi-

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crospheres have been prepared from poly(d,l-lactide) (PLA) [4,6,10] and poly(@-hydroxybutyrate)(PHB) [11,12]. The PLA microspheres were cast from CH,Cl,; the PHB microspheres were cast from chloroform. In both cases, free CCNU formed in the aqueous phase as solvent evaporation progressed to completion unless evaporation was interrupted and the aqueous phase emulsifier was removed from the system before solvent evaporation was complete. If no free crystals formed, the surface of PLA microspheres loadedwith 23%CCNU were smooth and free of defects [6]. If free crystals formed, thesurface of the microspheres was rippled and rough.The surface of 50-250 pm PHB microspheres is filled withmacroscopic pores. PHB is a relatively crystalline polymer that crystallizes during the solvent evaporation fabricationprocess, thereby generating a porous structure. Juni and Nakano [l31 show a SEM photomicrograph of50-400 pm PHB microspheres cast from CH,Cl, that illustrates their surface porosity. These large pores are notvisible in the surface of 15 pm PHB microspheres [ll]. If free CCNUcrystals formed duringsolvent evaporation, the amount of CCNU incorporated in the microspheres was reduced as one would expect. However, the efficiency of CCNU incorporation in 50-250 pm PLA or PHB microspheres was always high whether or not free crystals formed. Large amounts of CCNU were not partitioned into the aqueous phase or degraded during the fabrication process. This is significant in view of the sensitivity of CCNU to water. When efforts were made to prepare 15-pm diameter CCNU-loaded microspheres, the problem of free CCNU crystal formation increased markedly [12]. The high CH,Cl,-water interfacial area promoted development of free crystals, whereas the small size of the microspheres being formed made it difficult to minimize free crystal growth by removing the aqueousphase emulsifierbefore solvent evaporation was complete. Accordingly, smallPLA orPHB microspheres prepared to datehave CCNU loadings below 10% [12]. Fig. 4 is a DTA/TGA scan for a freshly formed 50-250 p m CCNU-loaded (23%) PLA microsphere sample [lo]. The DTA scanhas no event at 57"C, the Tg of PLA, or93"C, the crystalline melting temperature of CCNU. The broad exothermic event extending from approximately 115 to 140°C and the accompanying TGA weight loss are due to thermal degradation products. The absence of a crystalline CCNU melting point indicates that there are no crystalline CCNU domainsin this microsphere sample. The CCNU is molecularly dispersed in the PLA glass. The absence of a PLA Tg eventsuggests that CCNU plasticizes the PLA. Since the DTA scan began at 30"C, a DSC scan of a duplicate freshlyformed CCNU-loaded PLAsample was made from 0 to 70°C. It revealed a broad Tg event centeredat 25°C. The x-ray diffraction pattern of this second CCNUPLA sample was completely amorphous. The 25°C PLA Tg eventcoupled with the experimental observation that CCNU-loaded PLA microspheres

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sinter together on prolonged 22°C storage prompted the conclusion that PLA CCNU acts as a PLA plasticizer andismolecularlydissolvedin microspheres. The discovery that freshly formed progesterone-loaded PLA and lactide-glycolidecopolymer microspheres subjected to the same drying procedure as CCNU-loaded PLA microspheres retain 2-5 wt. percent CH+& led to the realization that the reduced PLA Tg reported abovecould be due in part to residual CH,Cl, and not plasticization of PLA by CCNU. The observation that CCNU tends to crystallize spontaneously in the aqueous phase as CH,Cl, evaporation proceeds is an indication that the mutual miscibility of PLA and CCNU is low. If they were highly miscible, the CCNU would not tend to form free crystals. In order togain more insight into how CCNU and PLA interact in CH,Cl,cast microspheres, the thermal behavior of a number of aged CCNU-PLA microsphere samples was evaluated. Table 2 summarizes the thermal data obtained [lo]. Question marks (?)in the table designate cases where the event could not be characterized reliably either because it was too diffuse or the thermal analysis did not extend to a sufficiently low temperature. PLA Tg values for the samples characterized fall at 21 f 4°C or 49 2 3°C. SamplesA , C, J, and E have a 21 rt 4°C Tg event which does not develop a well-defined endothermic peak.

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These samples do not have a Tm event attributable to the melting of CCNU crystal domains. Fig. 5 illustrates the nature of the PLA Tg event for sample A after 35 months of storage at -20°C followed by varying periods of storage at 20°C. There is no majorchange over the 20°C storage times examined. The small spike at 43°C for the sample stored 14 days at 20°C is believed to be an artifact. Fig. 6 also shows the 70°C CCNU fusion event characteristic of samples B and D. This peak is attributed to the melting of imperfect CCNU crystal domains. X-ray analysis confirms that crystalline CCNU is present in these microspheres. The crystal domains formed because the aqueous phase emulsifier used in the solvent evaporation process was not removed soon enough. The data presented above show that several samples with widely 4°C. In these different thermal histories have a PLA Tg event at 21 cases, the thermal history is not the key factor controlling behavior of the PLA Tg event. The absence of crystalline CCNU in the PLA microspheres when formed is the controlling factor. When the CCNU is molecularly

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Fig. 6 DSC scans of CCNU-loaded (20.4 wt. percent) PLA microspheres after 34 mo storage at -20°C followed by designated storage time at 20°C (From Ref. 10.)

dispersed, it appears to act in effectas a PLA plasticizer. When the CCNU forms crystal domains in the PLA matrix during solvent evaporation, the observed PLA Tg approaches that of CCNU-free PLA and behaves on aging like CCNU-free PLA. The contribution of residual CH,Cl, to the PLA Tg event of microspheres in which CCNU is molecularly dispersed remains to bedefined. PLA microspheresinitially loaded with 22 wt. percent CCNU storedin a desiccator in the dark show no changein CCNU contentduring 8 weeks at

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3°C [6]. Significant degradation occurred when the storage temperature was 22-37°C. After 8 weeks at 37°C approximately 46% had degraded. Free CCNU crystals stored at 22°C in a desiccator in the dark showed no detectable change in 8 weeks. Thus, CCNU molecularly dispersed in the PLA microspheres was more unstable than free CCNUcrystals. Whether this is due to residual CH,Cl, in the microspheres, CCNU-polymer interactions, oxygen, or moisture has not been determined. However, it is clear that the molecular dispersion of CCNU in the PLA microsphere is more unstable at 22 and 37°C than free CCNU crystals.

VI. CISPLATIN-LOADEDPOLY(d,l-LACTIDE) MICROSPHERES

Cis-diamminedichloroplatinum (cisplatin) is one of the most potent anticancer agents known. Accordingly,itwas incorporated in poly(d,llactide) (PLA)microspheres [14]. Spherical, unaggregated PLA microspheres were obtained by dispersing cisplatin in a CH,Cl, solution of PLA and subsequently emulsifying this dispersion in a pH 2 aqueous phase that was saturated with cisplatin and contained a mixture of 88% hydrolyzed poly(viny1 alcohol) and methylcellulose. Cisplatin loadings ranged from 4.5 to 45.5 wt. percent. At all cisplatin loadings examined, the microspheres isolated contained macroscopic pores. The higher the cisplatin loading, the more porous themicrospheres. Because of this porosity, such microspheres did not retain CH,Cl, as tenaciously as microspheres free of macroscopic surface pores. Cisplatin is insoluble in CH,Cl,, so it was present in the PLA microspheres as small, crystalline particles believed to besimilar in size to the 1- to 10-pm particles originally dispersed in CH,Cl,. X-ray microprobe analysis showed that thecross section of microspheres loadedwith 45.5wt. percent cisplatin contained a uniform distribution of cisplatin domains (Fig. 7). There was no preferential partitioning of the drug particles toward the outer edges of the microspheres or toward the interior. VII. SUMMARY OF MICROSPHEREMORPHOLOGICAL STUDIES

The microsphere examples discussed in Sections 11-VI illustrate that a diverse range of microsphere morphologiesexist. Factors that affect what morphology is obtained with a specific drug-polymer combinationinclude the natureof the drug, the nature of the polymer, the amount of drug in the microsphere, and the nature of the emulsifier in the aqueous phase. Microspheres prepared from biodegradable lactide polymers were discussed

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Fig. 7 X-ray microprobe photograph of cross sections of cisplatin-loaded (45 wt. percent) poly-d,l-lactide microspheres. (From Ref.14.)

exclusively in this chapter, because they are actively being developed for parenteral formulations. Essentially an infinite number of potential drugpolymer combinations could be used to prepare anessentially infinite number of different microspheres. Many would have morphologies analogous to those described in this chapter. However, many should differ significantly. Morphology studies of lactide microspheres containing drugs other than those discussed here indicate this is the case [2]. One way to cause significant morphological changes is to alter the mannerin which solvent is removed from the polymer-drug phase [15]. Another approach is to form microspheres from drug-polymer combinations with a high degree of mutual miscibility or mutual affinity. In such cases, drug crystallization may not occur even at relatively high loadings and polymer properties may be altered dramatically. A specific example is progesterone-loaded polystyrene microspheres [16]. Further studies such as those described in this

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chapter will provide valuable insight into thestructure of various polymerdrug combinations and suggest ways to control encapsulation process parameters so that unique microspheres are produced. REFERENCES 1. P. J. Watts, M. C. Davies, and C. D. Melia, Microencapsulation using emulsifi-

2. 3. 4.

5.

6. 7.

8. 9. 10. 11.

12.

cation solvent evaporation: an overview of techniques and applications, Crit. Rev. Ther. Drug Carrier Syst., 7(3):235-259 (1990). C. Thies, Formation of degradable drug-loaded microparticles by in-liquid drying processes, in Microcapsules and Nanoparticlesin Medicine and Pharmacy (M. Donbrow, ed.), CRC Press, Boca Raton,FL, 1992, pp. 47-71. J. P. Benoit, F. Courteille,andC.Thies,Aphysicochemicalstudyof the morphology of progesterone-loaded poly (D,L-lactide) microspheres, Int. J. Pharm., 29:95-102 (1986). J. P. Benoit, S. Benita, F. Puisieux, and C. Thies, Stability and release kinetics of drugs incorporated within microspheres, in Microspheres and Drug Therapy, Pharmaceutical, Immunological and Medical Aspects (S. S. Davis, L. Illum, J. G. McVie, and E. T. Tomlinson, eds.), Elsevier, Amsterdam, 1984, pp. 91-102. S. E. B. Petrie, Thermal behavior of annealed organic glasses,J. Polymer Sci. A2,10:1255-1272 (1972). S. Benita, J. P. Benoit, F. Puisieux, and C. Thies, Characterizationof drugloaded poly(d,l-lactide) microspheres, J. Pharm. Sci., 73:1721-1724 (1984). V. Rosilio, J. P. Benoit, M. Deyme, C. Thies, andG. Madelmont, A physicochemical study ofthe morphologyof progesterone-loaded microspheres fabricated from poly(D,L-lactide-co-glycolide),J. Biomed. Mat. Res., 25:667-682 (1991). M. Muramatsu, M. Iwahashi, and U. Takeuchi, Thermodynamic relationship J. Pharm. Sci., between alpha and beta forms of crystalline progesterone, 68~175-177 (1979). M. Cavalier, J. P. Benoit, and C. Thies, The formation and characterization of hydrocortisone-loaded poly((&)-lactide) microspheres, J. Pharm. PharmaCOL, 38:249-253 (1986). J. P. Benoit, G. Madelmont, F. Puisieux, and C. Thies, The storage behavior of CCNU-loaded poly (d,l-lactide) microspheres, Polymer Preprints 27(1):2728 (1986). M. C. Bissery,F. Puisieux, and C. Thies, A study of process parameters in the making of microspheres bythe solvent evaporation procedure, Proceedings of 3rd APGI Congress, Paris, 1983,233-239. M. C. Bissery, F. Valeriote, and C. Thies, In vitro and in vivo evaluationof 2) lactideandpoly(& CCNU-loadedmicrospherespreparedfrompoly( hydroxybutyrate), in Microspheres and Drug Therapy, Pharmaceutical, Immunological and Medical Aspects (S. S. Davis, L. Illum, J. G. McVie, and E. Tomlinson, eds.), Elsevier, Amsterdam, 1984, pp. 217-227.

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13. K.Juni and M. Nakano, Poly(hydroxyacids) in drug delivery, Crit. Rev. Ther. Drug Carrier Syst., 3:209-232 (1987). 14. G . Spenlehauer, M. Veillard, and J.P. Benoit, Formation and characterization of cisplatin-loaded poly(d,l-lactide) microspheres for chemoembolization, J. Pharm. Sci., 75:750-755 (1986). 15. T. R. Tice and D. H. Lewis, Microencapsulation Process, U.S. Patent 4389,330,1983. 16. F. Courteille, J. P. Benoit, andC. Thies, The morphology of progesteroneloaded polystyrene microspheres,J. Control. Rel., 30:17-26 (1994).

7 Drug Release from Microparticulate Systems Clive Washington The University of Nottingham, Nottingham,England

I.

Introduction

11. Measurement of Drug Release 111. Equilibrium Dialysis IV. V.

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Continuous-Flow Methods

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VI1. Mechanisms of Drug Release VIII. Diffusion in a Homogeneous Sphere IX. Diffusion Through an Interfacial Bamer

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Systems with Nonuniform Drug Distribution

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Heterogeneous ParticleMechanisms A. Dispersed and Phase-Separated Systems B. Erodable and Biodegradable Systems

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Empirical Models of Drug Release

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Summary

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INTRODUCTION

Disperse systems for drug delivery have found wide application in pharmacy; initially for external use as creams and ointments and later for oral administration and subcutaneous or intramuscular delivery. A significant number of materials have also been formulated for intravenous administration, and these represent a distinct class of products. Materials intended specifically for intravenous administration must contain an extremely small load of large particulates (5 pm is often quoted asa loose upper limit) , and this sets manufacturing and quality control criteria for these materials which are significantly more stringent than for external or oral formulations, in which the occurrence of the occasional large particle is usually immaterial. Consequently, particulates for intravenous use are normally formulated to a particle size which is well below 1 pm; often this is done simply to ensure thatmuch larger particles are absent. A typical example of such a material is the intravenous triglyceride emulsion Intralipid (Kabi Pharmacia, Sweden), which is emulsifiedto a droplet size of 200-300 nm. In addition, there is much interest in the biopharmaceutics of small particulates for drug-targeting applications. There is an extensive recent literature dealing with the preparation of microparticles for delivery of a wide range of drugs. In many cases, very small particles (those smaller than 100 nm diameter) show particularly interesting behavior; for example,such small particles can escape from the capillaries in certain fenestrated tissues and accumulate at sites of inflammation. There is also an increasing body of evidence that submicrometer particles can be taken up from the gastrointestinal tract into the systemic circulation. . Consequently, there is muchinterest in the preparation and characterization of submicron particulates containing active materials; for example, anticancer agents [l],antifungals, antibiotics [2-41, antiinflammatory agents [5,6], and peptides [7,8]. Unfortunately, the handling and characterization of such formulations is a good deal more complex than that of similar supramicron systems, a fact which, judging from the earlier literature,is only just being realized by many experimenters. Drug-release data have a number of potential applications. At the simplest level, the data can be used for quality control purposes to ensure

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the constancy of behavior of a manufactured product. It can further be used to try to understand the physicochemical structure of the delivery system and the release mechanism; however, this normally requires the fitting of the data to that computed for a model of the drug-delivery system. Finally, such data can be used in an attempt to predict the likely behavior of the system in vivo. This is a much more complex problem which involves a knowledge of the interaction of many biological components with the particle, and this area is poorly understood at present. The release of a drugfrom a carriersystem isactually a combination of a range of processes transferring drug between a range of sites. Some of these processes are illustrated in Fig. 1. The drug first has to move through the camer,usually by diffusion through a solid or pores; it has to cross the interfacial layer, at which it may be bound if it is surface active; and finally, it must diffuse into the continuous phase. If it has phase separated, the drug may have to dissolve inthe carrierparticle prior to diffusion, and the timevarying hydration of the carrier particle may be important in many systems. The experimental release rate is the result of all these processes, although one is usually considered to be rate limiting and So dominates the kinetics. Some of these processes can be driven in reverse if the concentrationof the solute in the external phase is sufficiently high; indeed, this may form a method for loading microparticles. Under these conditions, the reversible

Db- \+ D

D

Fig. 1. Some possible sites of distribution of a solute drug(D) or phase-separated drug (shaded regions) in a microparticle.

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*

k, /

kf

[Dlcontinuous

Fig. 2. Equilibrium of drug release between particle and continuous phase. equilibrium nature of drug diffusion willbe evident. The kinetics will depend on thedetailed mechanism, but as an example,consider the simplest scheme of Fig. 2, in which the drug in the particle is in first-order equilibrium with that in the aqueous phase, with no interfacial terms being significant. The rate of appearance of drug in the continuous phaseis givenby:

If the concentration in the external phase is zero, this reduces to:

.Consequently, the experimental release rate is only indicative of the release process behavior athigh dilutions; this is termed a perfect sink experiment, and it is generally desirable in order to simplify the interpretation of the experimental data.In concentrated systems, the experimentalrelease kinetics are also influenced by the reverse reassociation reactions, which are represented here by k, (which may include other compartments, such as readsorption to a surface site on the particles). Obviously this model may be complicatedby such factors as surface binding, carrier degradation, and diffusion in the carrier. More complex models involving ionization of the solute and surface binding have alsobeen analyzed[9,10], but in general, it is simpler to understand the behavior of perfect sink systems, so experiments are normally performed under these conditions. II. MEASUREMENT OF DRUG RELEASE

The measurement of drug release from submicronparticulates poses many difficulties which are not encountered for larger particulate systems. The

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main difficulty arises when an attempt is made to separate the released drug in the continuous phase from the suspended colloidal camer; this becomes increasingly difficult and slow as the particles become smaller; since the release rates increase with decreasing particle size, a size is reached at which the separation time is the same order as thedrug-release time. Under such circumstances, the time resolution of the experiment is inadequate anduseful data cannot beobtained. The size limitat which this occurs obviously depends on the camer and solute, butmost for purposes, one can expect to encounter difficulties for particles smaller than approximately 1pm. Emulsion systems may prove particularly troublesome owing to extremely rapid drug diffusion coupled with their polydisperse nature. Consequently, it ispreferable for many systemsto use analytical techniques which do not depend on separation of camer and solute. The methods available fall into four diffuse groups: (1) equilibrium dialysis, (2) sample and separate, (3) continuous flow, and (4) in situ methods. 111.

EQUILIBRIUMDIALYSIS

In equilibrium dialysis, a small volume of the concentrated drug-particle suspension is contained in a dialysis bag, which is immersed in a larger volume of continuous-phase acceptor fluid [ll-141. Preferably both compartments are stirred, and the drug then diffuses out of the particulate carrier into its local continuous phase andthen through the dialysis membrane into theacceptor phase, which is periodically sampled and assayed. This technique is highly misleading, since the particulate system is not diluted into a sink phase, and so sink conditions will not beachieved; under these circumstances, it will be difficult to interpret the release data. A simple analysis of this experiment (15) demonstrates thatthe measured rate of drug appearance in the external sink does not depend significantly on the particle release rate; instead it depends largely on thepartition coefficient of drug between its carrier and local acceptor phase. If we consider the model shown in Fig. 3, it can be shown that the concentration (A,)of drug in the external acceptor phase is given by:

where Kp is the partition coefficient and k, the dialysis membrane diffusion rate. This is independent of the particle release rate, k,, whichwe are seeking to measure. This partition-rate correlation is easily discernible in some published data; for example, the data of Sasaki et al. [l21for the release of a series of mitomycin C prodrugs from a triglyceride emulsion.

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DIALYSIS BAG

I

SINK VC

Fig. 3. Kinetics of nonsink membrane dialysis technique.

Levy and Benita [l61 confirmed the disadvantages of this method in studies of an emulsion containing diazepam in an Intralipid-like base; they also demonstrated a reverse dialysis technique, in which the emulsion was diluted into asink and small preequilibrated dialysis bagscontaining buffer were used to sample the continuous phase. This method has the advantage that a large sink can be used, but it was still found to be rather slow owing to the diffusion rate through the bags (about 1 hr), and the drug release rate from the emulsion could not be distinguished from the “release” of a true solution preparation using this method. The investigators concluded that emulsion carriers released their drug very rapidly, and they suggested that this could be studied if smaller dialysis microcapsules were available.

W.

SAMPLING AND SEPARATION METHODS

Examples of this method normally start by preparing a concentratedsuspension of the particulate carrierand rapidly diluting it into amuch larger bulk of acceptor phase [17-191. Samples of this diluted system are then removed at intervals, and the particles are separated so that the continuous phase may be assayed for released drug. This technique is perfectly adequate for larger microparticles, in which the separationcan be effected in seconds or minutes by filtration or centrifugation, and the release times are measured in hours. However, as the particle size decreases, the difficulty of separa-

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tion increases. Centrifugation times increase dramatically for thecomplete sedimentation of submicron particulates, and if the particles are polydisperse, extended centrifugation may be needed to sediment the smallest component. Filtration through membrane filters is an attractive alternative; however, the possibility of released drug binding to thefilters must be considered. Ultrafiltration membranes withlow binding properties are available, but these still adsorb small amounts of many drugs [20]. Even mildly hydrophobic materials may be strongly adsorbed to many filtration materials; in addition, the particles mayblock the filters, so that only limited sample volumes may be obtained. A rapid and useful variant of this technique used in our laboratories involves diluting a small amount (approximately 10 mg) of the particles in suspension into 100 mL of continuous phase ina 100-mLsyringe equipped with a 0.2-pm encapsulated membrane filter, so that samples of continuous phase may be expressed at suitable intervals; a time resolution of approximately 30 S may be obtained in this manner, but blockage of the filter normally limits the number of samples which can be obtained. A more technically sophisticated version of this method was described by Magenheim et al. [20], who used low-pressure ultrafiltration to separate samples of the released drug from a relatively large volume of sink fluid. This apparatus had a time response of a few minutes, which made itsuitable for studying microparticle formulations but too slow for emulsion release studies. The ultrafiltration step can be accomplished rapidly by centrifugal filtration [21], and simple centrifugal ultrafilters are available which make this a most convenient method. A particularly useful device in this respect is the Sartorius@ Centrisart (Epsom, UK) which uses an (initially empty) ultrafilter tube that floats on the top surface of the sample to becentrifuged inside the main centrifuge tube, and the filtrate enters this inner tube via centrifugal force. The advantage of this method is that the particles in the sample are forced downward away from the ultrafilter, thus avoiding filter clogging. An interesting final example of sampling and separation measurements was described by Widder et al. [22], who studied drug release from magnetite-albumin microspheres; these could rapidly be separated from the continuous phase using a magnet. V.

CONTINUOUS-FLOWMETHODS

The continuous-flow method most commonly uses an ultrafiltration cell to recover samples of clean continuous phase; this is fed from a pressurized reservoir and flows continuously through a detector (usually ultraviolet [UV]) for a direct continuous measurement of drug release [23,24a]. This technique is conceptually simple but presents a number of sources of error,

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with the most significant being variations (usually a decrease) in the flow rate through the ultrafiltration membrane as the experiment proceeds, thus distorting the release profile. The response time of such an experiment is approximately the time taken to replace the continuous phase in the ultrafiltration cell; consequently, a thin flat cell (i.e., one having a large ultrafiltration membrane area but a small total volume) is preferable. Qpical flow rates of 1 mL min" in a 10-mL cell suggesta 10-min time resolution; if the drug release rate is comparable to this, the time resolution can be improvedby deconvoluting the cell response function from the observed release function as we have described previously [25]. We suggest (but have not yet had time to implement) a significant improvement to this experiment. If the ultrafiltrate analysis is performed using a multichannel diode array spectrometer with multicomponent analysis software, it should bepossible to identify multiple components simultaneously in the filtrate. Consequently, the drug-carrier system in the cell could be spiked with an inert nonabsorbing marker dyein the continuous phase and theconcentration of both dye andreleased drug measuredin the ultrafiltrate. This would allow variations in flow rate to bemeasured and, more importantly, provide a simultaneous instrument response function for deconvolution of the cell response time. Scholsky and Warner (26) have described an ultrafiltration technique in which the diluted colloid was circulated around one side of an Amicon (Mass.) microfiber ultrafiltration cartridge, and the release buffer was circulated over the opposite side. The appearance of the drug on the acceptor side was monitored by continuous UV spectrometry. The time resolution of this instrument was not assessed but appears to be of the order of hours rather than minutes, presumably owing to the membrane diffusion rate.

VI. IN SITU METHODS In situ methods depend onthe use of analytical techniques which are only sensitive to the drug in solution and not to that in the solid particles or bound to theirsurface, so that theassay of released drug may be performed without any separation step, The difficulties discussed for many of the alternative methods would suggestthat in situ techniques would be particularly valuable; however, they have not found widespread use, largely on account of the difficulty of identifying suitable assay methods formost drug substances of interest. The most obvious method is to use optical absorption, since many molecules will have different spectroscopic properties depending on whetherthey are boundto theparticulate or freein solution. This methodwas used by Illumet al. [27] to study the release of rose bengal

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from cyanoacrylate microspheres. Scattering from the microspheres is a significant problem, and themethod can only normally be performedif the drug is strongly colored; the carrier particles often absorb or scatter light too strongly for measurements to be made in the UV region of the spectrum. It,is possible that these difficulties could be minimized by the use of multichannel spectrometers with multicomponent analysis, but this has apparently not been explored. A novel in situ sampling method was described by Minagawa et al. [28], who attempted to measure the release of isocarbacyclin methyl ester from an emulsion formulation using silanized glass beads to adsorb the free hydrophobic drug. This technique resulted in a time resolution of approximately 15-30 S and an apparentrelease time of a few minutes from a typical triglyceride emulsion. As discussed below,techniques of this type are really too slow to study dynamic distribution processes in submicron emulsions, but they could prove to bevery useful for solid microparticle studies. Electrochemical techniques would appear to be valuable, but again have been little used. Polarography and ion-selective detection would have a response time of seconds, but very few drugs are amenable to detection using these methods. However, an attractive alternative is to use pH as an indicator of drug release. Many drugs are weak acids or bases, and their release can be monitored by measuring the neutralization of a suitable medium as the drug diffuses into it.We have used this method to study the release of model organic acids and the basic drug chlorpromazine from triglyceride emulsions [29]. This experiment has a time resolution of approximately 1S, and this is sufficiently rapid to measure drug release from emulsions, which occurs over a period of tens to hundreds of seconds. The results demonstrate thatthe release of drug is limited by interfacial transport, andso is sensitive to thenature of the emulsifier as well as the drug. The rapidity of drug releasein these emulsion systems casts doubt on many earlier measurements in this area.

VII. MECHANISMS OF DRUG RELEASE It is rarely possible to make any interpretation of drug release dataunless the experimenter has some preconceived model concerning the structure and composition of the system under study. Unfortunately, many workers give the impression that their systems consist wholly of monosized spherical particles uniformly loaded with drug; this is rarely the case, and it is useful to discuss some of the ways in which real formulations may depart from this ideal. First, the particles can depart from a monosized distribution of spheres. They usually have a distribution of sizes, with the occasional large

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particle having a significant fraction of the total drugmass. Without going into details of release mechanisms, this will imply a range of release rates, with the smaller particles showing rapid release and the larger ones a slower component. In theory it is straightforward to integrate the release rate expression over the particle size distribution if this is known. The particles may also be nonspherical; many papers show electron micrographs of “spherical” particles which show minor or gross distortions; these can be inducedby freeze-drying, so that the dried formulation shows very different release profiles to thesuspension formulation. Also, the structure of the particle may not be uniform; it could display core-shell behavior, with the drug being nonuniformly distributed across the particle section, or multiple phases of drug and carrier. It is evident that if the particles are porous or have a fractal interior structure, an even greater rangeof uncertainty exists concerning the drug distribution and release behavior. Finally, most polymer particles have varying degrees of crystallinity, which can be related to thedrug release rate from them. Second, we must consider all the possible locations for the drugload in the formulation. Since smallparticles have relatively rapid release rates, the drug will probably be in thermodynamic equilibrium between the carrier andits continuous phase. To ascertain the position of this equilibrium, it is usually helpful to isolate the continuous phase by ultracentrifugation and assay for unentrapped drug. The drug in the continuous phasemay be present in true solution or more likely in micellar solution if a significant amount of excess surfactant, from the particle preparation, is present. Removal of this surfactant can be troublesome; if it is dialyzed out, the drug it carried can be precipitated in the sample, so that two particle populations, one of carrier and one of pure drug, are present. If lecithin surfactants are used, an extensive liposomal phase may be present (30). The effects of this liposomal material on release of isocarbacyclin methyl ester froma triglyceride emulsion was highlighted by Minagawaet al. (28); over a third of the drug was present in the liposomes, and this caused a surprisingly long release profile for this fraction of the drug. Finally, if the drug is surface active, it will adsorb to theoutside of the particles and may provide a burst effect on dilution. Consequently, it will be appreciated that release data can only be unequivocally modeled if great effort is taken tocheck allof these possibilities prior to measuring the release profile. Despite all these difficulties, many experimenters still try to draw conclusions about the particle structure andrelease mechanism from therelease kinetics. As we will see, there is one furtherhurdle; that is, many of the model release mechanisms from particles of different types lead to very similar release kinetics, normally exponential or exponential-sum. As a result, it is rarely possible to draw

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conclusions about the structure of the particle from the release profile unless a considerable amount of supporting evidence is available.

VIII. DIFFUSION IN A HOMOGENEOUS SPHERE This is the simplest model of a drug-releasing particle; it assumes that the drug is initially uniformly distributed in a sphere made of material with a uniform diffusion coefficient. Because of its simplicity, it can be modeled analytically; the mathematicsfor this process are well known [31] and were applied to the drug-release problem by Guy et al. [32]. The expected release-time profile for this model is

where M, is the amount of drug in the particle at time t and the dimensionless time, T, is given by:

D is the diffusion coefficient of the drug in the oil droplet and r is the droplet radius. This sum-of-exponentials series provides a decay rate that initially is steeper than a single or best-fit exponential, and at long times is slower than a single or best-fit exponential; this emphasizes thedifficulty in obtaining a sustained release effect from a monodisperse sphere formulation of this type, and indeedthe rapid initial release may be indistinguishable from the burst effect observed in many systems. In order to simplify a discussion of this model, simplified forms of this equation have been used. Expansion and truncation of the series yields:

which suggests that the initial release rate is inversely dependent on the particle size and proportional tothesquareroot of time. This latter relationship is often termed the Higuchi law; this is in fact an incorrect use of the term, since Higuchi derived this relationship for a completely different model system. Truncated series of this type show an increasing error at later times, and they should only be used for analysis of the release of the first 5-10% of the drug or unacceptable errors will result.

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Despite this unavoidable consequence of the model, many investigators emphasize that their data obey the square root law over much longer intervals; much of theliterature in this area is clouded by the blind application of this law. Addition of a further termin the series yields [33]

which provides a fit for data at“longer short (!)” release times. An approximation for drug release at long times (i.e., the release of the last few percent of the drugload) is given by:

This expression can be rearranged intothe linear form:

Thus, a graph of ln(1 - M j M o ) against time will have a limiting slope of (-$D)/? at long times, enabling the diffusion coefficientof the drug in the oil droplet to becalculated. The simple uniform diffusion model is applicable to those systemsin which the drug is uniformly dissolved in the particle, and the matrix does not degrade during the experiment or present an interfacial barrier. In emulsions, the most obvioussystem, release is controlled by the interfacial transfer characteristics, and it is only in solid systems that bulk diffusion becomes limiting. Some swelling noneroding polymersmay release by this mechanism; normally these contain pores of solute dimensions through which the drug can diffuse; if the pores are of larger dimension than the drug molecules, then a heterogeneous model (see below) would be more appropriate [34]. An interesting variant of the diffusion controlled model was discussed by Cremers et al. 1351 to describe the release of drugs from ion-exchange microspheres. These are prepared from combinations of proteins (normally albumin) and negatively charged polyelectrolytes such as heparin [36,37] or poly (L-glutamate) [38]. In such systems, the drug is retained in the particle by ionic interaction with the polyelectrolyte and is released by an ion-exchange process when ions from the release medium diffuse into the particle. Cremers et al. [35] modeled this process by assuming that the ion transport into theparticle was rapid and in equilibrium with the rapid

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solvent swelling of the particle. Under these conditions, diffusion the of the drug in the matrix is rate limiting, and so the release kinetics is similar to that of the simple diffusion model. However, there is one important difference; theactivity of free drug (i.e., not bound to the polyelectrolyte)in the microsphere is controlled by the ionic strength of the release medium. Thus, the fraction of drug extracted increases as the ionic strength of the medium is increased, although the release rate is constant and controlled by the diffusion coefficientof the drug in the microsphere. One oft-suggested method of circumventing the problem of nonlinear release and obtaining a constant release profile is to use a collection of phases spheres of varying sizeso that small spheres contribute to the initial of drug release, whereas larger spheres contribute to release at longer times. A few moments thought will reveal that this argument is flawed; a collection of spheres with varying sum-ofexponential releaseprofiles, however weighted, can only yield an overall sum-of-exponentials release profile; addition of a wider range of particle sizes will just make the initial slope steeper and the tail longer. A flat release profile cannot be decomposed into a sum of exponentials. However, we should note that this approach may be feasible if the release rateof the individual particles was not identical to this theoreticalspherical particle releaseprofile. IX. DIFFUSION THROUGHAN INTERFACIAL BARRIER The previous model assumed that the release of drug was limited by the rate at which it diffused through the carrier particle. Although this model may be valid for particles of larger sizes or those madeof solid materials, it is unlikely to be limiting for smaller liquid droplets such as those found in parenteral emulsions. In such cases, the expected diffusion coefficients would lead to drug release times of a few tens of microseconds; we have measured the release ratesfrom model emulsions to be of the orderof tens must beof much greater relevance of seconds; and so the interfacial barrier in this case. The model for releaseof drug from a spherical particle in this situation is more complex than that for the situation in which a barrier is not present. In fact Guy et al. [32] state that no complete solution exists and present approximatesolutions for shortand long times. At short times, the release is zero-order andis given by:

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where K is a reduced interfacial rate constant related to the trueinterfacial rate constant k, by:

It is difficultto verify this regimen for real systems, since the presence of a burst release component often overlaps on this timescale. At long times, where T >> 1, the release is given by a single exponential decay:

M,

-3kf MO - l - e x p ( T ) This is a useful method for obtaining k,, since it is straightforward to linearize: "

Thus, a graph of ln(1 - M,/M,) against time will have a limiting slope at long times of -(3k,)/$ enabling the interfacial transport rate constant of the drug betweenthe particle and the release medium to befound. X. COATEDPARTICLES

The coated spherical particle is a rather fundamentaltype of delivery system, in a range of macroscopic controlled-release devices, as and it has been used well as microspheresin which the outer coating or membrane is distinct in composition from the inner core. Liposomes may be classified as coated particles, and Pekarek et al. [39] described a method of making two-layer microspheres by the control of surface wetting and interfacial energetics. A full analysis of diffusive release from a coated particle is complex, but it has been fully detailed by Lu and Chen [40]. The full solution can be simplified by taking a number of specific cases; the most useful is that for release into aninfinite sink. Solutions are also givenfor the cases where the diffusion coefficient inthe core or the inner matrix are rapid. XI. SYSTEMS WITH NONUNIFORM DRUG DISTRIBUTION

It has been suggested that the burst effect observed in many systems is due to weakly bound drug at or near theparticle surface, and if the particles were briefly washed, this would deplete the drug concentration near the particle surface and lead to a more uniform release of drug. Lee [41a,41b,42] studied release from macroscopic devicesof this type, but the

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results are equally applicable to particulate systems. The results suggest that by suitable selection of concentration distribution it ispossible to achieve steps or peaks in the release rate rather thana continuously declining rate, and that,for example, a pseudo-zero-order release may be possible by using a multilayer structure with varying drug concentrations in the layers. Although it is easier to visualize these techniques being applied to relatively large particles, it maybe technologically feasible to preparecolloidal systems withthese characteristics. XII.

HETEROGENEOUSPARTICLEMECHANISMS

The simple uniform spheremodels discussedabove are probably only valid for a small number of systems, including emulsion formulations and some lightly doped nondegradablemicroparticles. The majority of microparticle systems are heterogeneous, andso drug diffusion through themis complex. We could imagine, for example, polymericmicrospheres, in whichthe crystallinity varies spatially with polymer blend; microspheres in whichthe drug and carrier are phase separated intodistinct domains; and systems where drug diffusion is so slow that degradation of the microsphere was rate limiting. For example,Magenheim et al. [20] reported variations in the release of indomethacinfrom poly-(lactide-co-glycollide) (PLGA)and poly(1actic acid) (PLA) nanoparticles which could be attributed to variations in the solvent permeability of the polymer. Many examples of heterogeneous systems have beenstudied in the literature tovarying degrees of detail. A. Dispersed and Phase-Separated Systems

A potentially common system is that in whichthe drug has phase separated from theparticle matrix into pure domains and is then released by diffusion through the polymer. Although systems of this type have been widely studied as macroscopic devices for controlled release (e.g., see ref. 43), many microscopic particles also display these characteristics. Baker and Lonsdale [33] studied a system of this type, and they found thatthe release rate was given by:

where C, is the drug solubility in the dissolution medium and A the drug loading per unit volume. The drug particles are assumed to be small compared with the particle radius. Release of drugs from heterogeneoussystems has often been treated using the expression derived by Higuchi [M] for the flux, Q, from a planar slab of polymer containing the drug:

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Q = ~ D C , ( ~ -C c,)t , In this expression, c, is the solubility of the drug in the matrix, c, is the initial drug concentration, and c, is the drug solubility in the sink phase. Unfortunately, this expression refers to an infinitely deep matrix which is not significantly depleted, and so it is rarely applicable if the majority of the particle’s drug load is released. However, the use of expressions in which release is proportional to the square root of time are so firmly ensconced in pharmaceutics that little thought is normally given to their significance or applicability. As the loading of solid drug in the matrix is increased, ultimately a point will be reached at which the drug particles are in contact with each other; this normally occurs at loadings of around 10-20%. In this case, as the drug diffuses out of the matrix, solvent-filled channels are left, which act as pathways through which the remaining drug is preferentially released. The full expression for release from a spherical matrix of this type is rather complex [44] and involves terms for the porosity and tortuosity of the matrix. These terms can only be accessed with difficulty, and it would appear that such treatments are straining the limits of the pure analytical approach to drug-release modeling. Ultimately numerical simulation may prove to bemore useful for these systems. Examples of pure multiphase systems are difficult to find, since many fall into the category of erodable or degradablesystems rather than diffusive systems. However, one importantclass of multiphase systems ismultiple emulsions, in which the drug is carried in an inner (normally aqueous) phase within an oil droplet. These systems have been extensively studied, although they have not been widely exploited owing mainly to stability problems of the inner aqueous droplets within the oil droplets [45-471. Magdassi and Garti [48, 491 suggested that the Baker and Lonsdale [33] expression should be applicable to release from multiple emulsions, and they found a reasonable agreement with experiment, as did Raynal et al. [50], who studied the release of sodium lactate from a liquid paraffinisopropyl myristate multiple emulsion system. B.ErodableandBiodegradableSystems There is an extensive literature on hydrating and eroding systems, which is largely applicable to macroscopicdevices for sustained-release applications. Eroding devices differ from those previously discussed, because drug release is by definition controlled by erosion of the particle rather than diffusion of drug within the matrix. This represents an increasingly important class of particles owing to the currentinterest in biodegradable particles for drug delivery and particularly for the delivery of macromolecules

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such aspolypeptides and DNAfractions. The most commonlyused materials for such applications are poly(1actic acid) (PLA) andpoly(glyco1ic acid) (PLG) and their copolymers (PLGA); polyanhydrides are also receiving much attention [51,52]. These polymers can be madeto degradein vivo on a timescale of hours to weeks, depending on the device size and polymer blend, and can easily be made intoparticles by solvent evaporation methods. Allemann [53a,53b] described a salting-out method which appeared to reduce the burst effect from the resulting microspheres. These polymers degrade by backbone cleavage by simple hydrolysis, also known as type 3 erosion [54]. It is fairly certain that both macroscopic devices and microsphere formulations degrade uniformly in the bulk rather thanprogressively from the surface. The erosion rates are largely controlled by the crystallinity of the polymer, with amorphous polymers showing the most rapid degradation, and polymers degrading more rapidly above the glass transition temperature [55,56]. The older literature on these systems has been reviewed by Heller [57]. More recent studies have provided a detailed description of the effects of polymer chain length and copolymer mix on release; for example, the work of Le Corre et al. [58] demonstrated that release rates of bupivacaine from pure PLA microparticles varied with polymer chain length, with the shortest polymers showing the most rapid release rate, and an increased proportion of glycolide also causing an increase in release rate. The drug release from a surface eroding particle, in contrast toa bulk eroding particle, is given by:

where k, is an erosion rate constant, r is the initial radius of the sphere, and c, the concentration of drug in the sphere [59]. There is little modeling work on the release from bulk eroding particles, although these appear to lose no mass until they have reached a critical chain length (m.w. 15,000), after which breakdown is rapid [55]. In general, however, it is inappropriate to use equations of this type to describe release, because the particles are rarely homogeneous. For example, the PLAPLGA particles produced by Le Corre [58] referred to above showed a porous cross section; the origin of the poresis uncertain, but may be due tophase separationof oily droplets of the drug (bupivacaine base) within the polymer as the dichloromethane solvent was lost. This complex structure led to a rapid initial release rate, presumably frompores close to thesurface, followed by a slower rate which the authors suggested was due to the onsetof polymer degradation. These polymers appearto behave similarly when prepared as spray-dried particles [60];such particles are normally hollow microcap-

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sules. Particle degradation appeared to be localised at specific pore sites, which presumablywere also responsible for drug release. Iwataand McGinty (61) described multiphase PLGAparticles prepared by a multiple emulsion method, which degraded internally over several weeks in the release experiment despite maintaining a uniform wall structure. It is evident thata detailed modeling of the release kinetics from such increasingly complex systems will be demanding, to say the least. However, there is much interest in these systems because it may be possible to use them to provide delayedpulsatile release of drugs (see below). The other majorcandidates for the production of biodegradable microspheres are naturally occurring proteins and polysaccharides. Albumin microspheres have been studied for many years in this regard (e.g., see ref. 62), as have a number of other proteins. Gelatin [63] and hyaluronic acid [64] are among a range of polysaccharides studied for this purpose. These systems are normally made by a water-in-oil emulsion technique, leading to microsphere sizes of a few micrometers down to hundreds of nanometers if high shear dispersion is used. There is some difficulty in the interpretation of release data from these systems, since they are normally fairly stable in sterile aqueous media, andso proteases are normally added to the release medium in order to bettersimulate their degradation in vivo. It may thus appear that surface erosion modelswould be applicable; however, the actual behavior of the particles is more complex, since the microspheres normally swell prior to being digested, and so one must assume that some enzyme is transported into the core of the particle during this initial swelling phase. This behavior was demonstrated by Magee et al. [65], who studied the size of albumin microspheresby laser diffraction as they were digested in a trypsin-containing medium; the microspheres swelled to several times their initial diameter over 20-30 mins prior to being degraded over a further30-60 min by the enzyme. Control of release rates from protein or polysaccharide particles can be achieved by varying the particle size or the degree of crosslinking. For example, Jayakrishnan et al. [66] demonstrated that the release of methotrexate from casein microspheres was influenced by the degree of crosslinking as a result of varying the amount of glutaraldehyde crosslinker used in the particle preparation; similar results were found by Rubino [67] and by Dilova andShishkova [68]for chemically and thermally denatured albumin microspheres.

XIII. REGULATEDDRUG RELEASE The most sophisticated drug-delivery systems now attempt to control or regulate the release of drug according to some local chemical signal or

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biological need in the tissue. Such systems are called regulated-release devices, and they evidently pose significant difficultiesfor in vitro characterization, since it is necessary to confirm that the release rate responds to the external stimulus. Although the study of such devices is in its infancy, a number of systems have been developed; these have largely been macroscopic rather thanparticulate systems. It seems inevitable, however, that this technology will be applied to particulates in the near future. For example, Yui et al. [69] demonstrated that crosslinked hyaluronic acid was degraded by hydroxyl radicals, and thatthis could be used to trigger degradation of the matrix at inflammation sites. Release can also be triggered by an external stimulus. The best known of these systems are the magnetic microspheres, in which the application of an external magnetic field causes both local accumulation and modifies the drug-release rate. Saslawski et al. [70] demonstrated that this technology could be used to deliver insulin from ferrite-containing alginate microspheres, with an oscillating magnetic field causing a 50-fold increase in the insulin release rate.

XIV. PULSATILE RELEASE SYSTEMS There is much interest in producing systems which release their payload in one or morepulses over a controlled time period. Such a system would be useful for the delivery of multiple-challenge antigens and peptide hormones. Pulsed systems are generally produced by engineering a time delay into the particle-degradation mechanism; for example, Kibat et al. [24b] demonstrated that liposomes coencapsulated with phospholipase would release their drug payload after a delay, corresponding to thetime taken for the phospholipase to destroy the integrity of the liposomes. Delayed release can also be achieved with some of the larger polylactide/glycolide microsphere systems [71], since these normally show a lag phase prior to release; this corresponds to thetime taken forsolvent swelling and internal polymer degradation. Other pulsed-release systems under study include polyiminocarbonates [72] and poly(ortho esters) [73]. XV. SIMULATION METHODS FOR DRUG RELEASE The gradual increase in complexity of drug-release models ultimately causes them to become mathematically intractable or incapable of analytic solutions. In such cases, it is possible to tackle the problem by simulation techniques, in which a particle is set up on a lattice, and Fick’s laws applied to calculate the diffusion of the drug as a function of space and time. Models of this type can take intoaccount processes such as the hydration of the parti-

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cle, and the consequent spatial variation in the diffusion coefficient, variable drug distributions, and drugbinding at interfaces, depending mainly on the amount of computing power available. The use of such methods is in its infancy, but a number of simple deviceshave beenstudied in this way. Armand etal. [74] used this technique to study release from Eudragit matrix formulations, in whichthe primaryrelease mechanism wasthe diffusion of solvent into the polymer, followed by dissolution of drug and its diffusion out of the device. Both experiment and data demonstrated that drug release follows a square root law for small release fractions after a lag phase. Liu et al. [75] used simulation techniques to study the release from a loaded sphere coated with a further layer of polymer, in which solvent penetration was taken intoaccount. Their experimental data was obtained from macroscopicsystems (several millimeters in diameter), and goodcorrelation with the calculations was found for the thinner polymer coatings.

XVI. EMPIRICAL MODELS OF DRUG RELEASE Many investigators have regarded the analytic modeling of drug release as unjustifiably complex and have used more empirical techniques for data analysis. These approaches vary from those which provide useful data about the system dynamics to those which one is convinced have been adopted simply to minimize the size of error bars in a graph. Probably the most useful of the former group is the diffusional exponent approach described by Peppas and colleagues [76,77],which goes some way to explainning the behavior of hydrating or eroding systems. In such systems, the diffusion coefficient is not constant, and the term anomalous d i m i o n is often used to indicate that a constant value of the diffusion coefficient does not satisfactorily fit the data. The termsFickian and non-Fickian are also used to indicate whether or not a material is diffusing with a temporally and spatially constant diffusion coefficient; these are somewhat misleading, since, at a particular instant and point in space, diffusion always occurs according to Fick's law. The diffusional exponent method proposesa power law relationship for drugrelease:

The constant n is termed the diffusional exponent, and it should equal 0.5 for diffusional (Fickian) release from a planar slab. Values greater than0.5 indicate anomalous diffusion, -and are generally indicative of a system which swells in the solvent prior to diffusional release. Analysis of the

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model and comparison to the exact solutions demonstrates thatn is equal to 0.5 for a flat slab and 0.432 for a sphere. Since simple diffusive release from spheresis often adequately fitted when n = 0.5, it is evident that this approach requiresprecise data toallow the extractionof a useful value for n. A corollary to this is that many of the experimental techniques in the literature require improvement before they can be used to discriminate between the various mechanisms of drug release. One of the morepopular empirical relations is to describe release as a biexponential process:

of the two lifetime components into Where k, and k, are the rate constants which the decay function is being decomposed. The exponentials usually consist of a rapidand a slow function, being assigned to “burstphase” and “sustained release,”respectively. The decomposition of a monotonically decreasing function into exponential componentsshould be performed with some caution,since it is an ill-posed problem. Lanczos [78]demonstrated that asum of three exponentials waswell fitted by a sum of two exponentials with distorted time constants and amplitudes, and further thatrecovery of exponential components requires data of an extremely high degree of accuracy, far beyond that observed for most drug release experiments.Attempts to obtain more than two release componentsfrom experimental datamust often be treated with scepticism. A corollary of this problem is that it is extremely difficult to extract true multiple exponentials, so that when these exist, a double exponential function provides an apparently adequate model. This has further led many experimenters into the misconception that the exponential components correspond totwo distinct physical processes (which may be true in some cases), and further that monodisperse homogeneous microspheres display a single exponential releaseprofile (which we have seen is completely false). The use of empirical functions to model release data has been taken to an extremeby Lin [79], who tried 10 separatesimple functions as fitting equations forthe release of theophylline microcapsules. Although abest-fit equation was found (lly = A/x + B ) , this approach does not significantly increase our understandingof the physical chemistry of the system. XVII. SUMMARY Our understanding of microparticle systems has developed considerably over the past 10 years. In themid 1980s, a typical drug-microparticlepubli-

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cation was characterized by a rather fuzzy electron micrograph of vaguely spherical adhering objects, with possibly a “laser sizer” size measurement that did not agree with the microscopic dimensions. Fortunately, we now have much more well-characterized systems, with details of internal morphology and accurate size distributions, and we have gained experience in the reproducible preparation of high-quality dispersed systems. Despite these achievements, the two central problems still remain. First, how do we design a microsphere system that will have specific desired properties? Second, evenif we know exactly whatis required, how do we manufacture it? The second question is still largely a black art, with pockets of knowledge centered around specific drug-particle combinations and only rather vague ground rules being known. It ispossible that the application of expert systems may assist in the correlation of the enormous amount of scattered information and experience that at present is poorly accessible. The first question is more tractable; as we have seen, models of many particle types can be created and understood. However, many details can only be explained qualitatively after the event. As a result, microparticle development is still an iterative process, with developers cycling round a “make-try-modify” loop which in many cases never reaches a usable conclusion. It would save many hours of experimentation with expensive materials (not to mention the lives of many animals) if such systems could be tested by simulation prior to production. Ultimately, a full understanding of the behaviorin vitro and in vivo of drug release from a complexsystem like a microparticle suspension is probably beyond the possibility of a rigorous analytic solution. Systems may ultimately be evaluated using numerical simulation of dissolution and diffusion in three dimensions, with full details of their heterogeneous in vivo environment. This would require a massive computing effort,but it is well withinthe capabilities of modem parallel processors. It would then be possible to study, for example, the effect of changing the polymer blendor drug partition coefficienton the pharmacokinetics of an intramuscular particle depot. Under these circumstances, it finally would become possible to design a microparticle system rationally by computational methods, by much the same philosophy that new chemical entities are developed at present. REFERENCES 1. D. J.Kerrand S. B. Kaye, Chemoembolism in cancer chemotherapy, Crit. Rev. mer. Drug Camer Syst., 819-37 (1991). 2. S. S. Davis, C. Washington, P. West, et al., Lipid emulsions as drug delivery systems, Ann. N.Y. Acad. Sci., 507:75-88 (1987).

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3. K. A. Lamb, C. Washington, and S. S. Davis, Toxicity of amphotericin B emulsion to cultured canine kidney cell monolayers, J. Pharm. Pharmacol., 43~522-524 (1991). 4. C. Washington, M. Lance, andS. S. Davis, Toxicityof amphotericin B emulsion formulations, J. Antimicrob. Chemother., 31:806-808 (1993). 5. K. Yokoyama, H.Okamoto,M.Wantanabe, et al.,Developmentofa corticosteroidincorporatedinlipidmicrospheres,DrugsExp.Clin.Res., 11~611-620 (1985). 6. Y. Shoji, Y. Mizushima, A. Yanagawa, et al., Enhancement of antiinflammatory effectsof biphenylacetic acid by its incorporation into lipid microspheres. J. Pharm. Pharmacol., 38:118-121 (1986). 7. S. S. Davis, L. Illum, andE. Tomlinson (eds.), Delivery Systems for Peptide Drugs. NATO Advanced Science Institutes series, Vol. A125, Plenum Press, New York, 1986. for delivery 8. P. Couvreur andF. Puisieux, Nanoparticles and microparticles the of polypeptides and proteins, Adv. Drug Deliv. Rev., 10:141-162 (1993). S. L. Silvestri, Theoretical considerations of drug 9. R. T. Lostritto, L. Goei and release from submicron oil in water emulsions, J. Parent. Sci. Techno]., 41: 214-224 (1987). 10. S. Silvestri, L. L. Wu, and B. Bowser, Release of polyionizable compounds J. Pharm.Sci.,81:413-418 fromsubmicrometeroilinwateremulsions, (1992). 11. M. Hashida, T. Yoshioka, S. Muranishi, and H. Sezaki, Dosage form charac1: Stability and drug release, Chem. teristics of microsphere in oil emulsions. Pharm. Bull., 28:1009-1015 (1980). 12. H. Sasaki, Y. Takakura, M. Hashida, et al., Antitumour activityof lipophilic prodrugs of mitomycin C entrapped in liposomeor o/w emulsion, J. Pharm. Dyn., 7:120-130 (1984). 13. S. Benita, D. Friedman, and M. Weinstock, Pharmacological evaluation of an injectable prolonged release emulsionof physostigmine in rabbits, J. Pharm. Pharmacol., 38:653-658 (1986). 14. S. Miyazaki, N. Hashiguchi,W. M. Hou, et al., Preparation and evaluation in vitro and in vivo of fibrinogen microspheres containing adriamycin, Chem. Pharm. Bull., 34:3384-3393 (1986). for the measurement 15. C. Washington, Evaluations of non-sink dialysis methods of drug release from colloids: effects of drug partition, Int. J. Pharmaceut., 56~71-74 (1989). 16. M. Y. Levy and S. Benita, Drug release from submicronized olw emulsion: a new in vitro kinetic evaluation model., Int.J. Pharmaceut., 66:29-37 (1990). 17. D.C.Tsai, S. A.Howard,T. F. Hogan, et al., Preparation and in vitro evaluation of polylacticacid-mitomycinCmicrocapsules, J. Microencaps., 3~181-193 (1986). 18. N. Farah, J. Bouzon, M. Rollet, et al., Dry emulsion-A sustained release form: Modelling of drug transfers in liquids, Int. J. Pharmaceut., 36:81-88 (1987).

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34. R. S. Langerand N. A.Peppas,Presentandfutureapplicationsofbiomaterials in controlled drug delivery systems, Biomaterials, 2:201 (1981). 35. H. F. M.Cremers,R.Vemjk, H. P.J.M. Noteborn, et al., Adriamycin loadingandreleasecharacteristicsofalbumin-heparinconjugatemicrospheres, J. Control. Rel., 29:143-155 (1994). 36. W. E. Hennink, J. Feijen, C. D. Ebert, and S. W. Kim, Covalently bound conjugates of albumin and heparin: synthesis, fractionation and characterization, Thromb. Res., 29:l-13 (1983). 37. H. F. M. Cremers, G. Kwon,Y. H. Bae, et al., Preparation and characterization of albumin-heparin microspheres, Biomaterials, 15:38-48 (1994). 38. W. E. Longo, H. Iwata, T. Lindheimer, and E. P. Goldberg, Preparation and drugreleaseproperties ofalbumin-polyglutamicacidadriamycinmicrospheres, Polymer Prep., 2456-57 (1983). 39. K. J. Pekarek, J. S. Jacob, and E. Mathiowitz, Double-walled polymer microspheres for controlled drug release, Nature, 367:258-260 (1994). 40. S. M. Lu andS. R. Chen, Mathematical analysis of drug release from a coated particle. J. Control. Rel., 23:105-121 (1993). 41a. P.I. Lee, Diffusion release of a solute from a polymeric matrix: Approximate analytic solutions,J. Membrane Sci., 7:255-275 (1980). 41b. P. I. Lee, Kinetics of drug release from hydrogen matrices, J. Control. Rel., 2~277-288 (1985). 42. P. I. Lee, Initial concentration distribution as a mechanism for regulating drug release from diffusion controlled and surface erosion controlled matrix systems, J. Control. Rel., 4:l-7 (1986). 43. J. R. Cardinal, Matrix systems, in Medical Applications of Controlled Release (R. S. Langer and D. L. Wise, eds.), CRC Press, Boca Raton,FL,1984, pp. 86-97. 44. T. Higuchi, Mechanism of sustained-action medication: theoretical analysis of rate of release ofsoliddrugsdispersedinsolidmatrices,J.Pharm.Sci., 52~1145-1149 (1963). 45. A. T. Florence andD. Whitehill, Some features of breakdown in water-in-oilin-water multiple emulsions, J. Coll. Interface Sci., 79:243-256 (1981). 46. A. T. Florence and D. Whitehill, The forulation and stability ofmultiple emulsions, Int. J. Pharmaceut., 11:277-308 (1982). 47. J. A. Omatosho, T. L. Whateley, T. K. Law, and A. T. Florence, The nature of the oil phase and the release of solutes from multiple W/O/W emulsions, J. Pharm. Pharmacol., 38:865-870 (1986). 48. S. Magdassi and N. Garti, Release of electrolytes in multiple emulsions: Coalescence and breakdown or diffusion through oil phase, Colloids Surfaces, 12:367-373 (1984). 49. S. Magdassi and N. Garti, A. kinetic model for release of electrolytes from W/O/W emulsions, J. Control. Rel., 3:273-277 (1986). 50. S. Raynal, J. L. Grossiord, M. Seiller, and D. Clausse, A topical W/O/W multiple emulsion containing several active substances: formulation, characterization, and studyof release, J. Control. Rel., 26:129-140 (1993).

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51. Y. TabataandR.Langer,Polyanhydridemicrospheresthatdisplaynearconstantreleaseofwater-solublemodeldrugcompounds,Pharm.Res., 10:391-399 (1993). 52. Y. Tabata, S. Gutta, and R. Langer, Controlled delivery systems for proteins using polyanhydride microspheres, Pharm. Res. 10:487-496 (1993). E. Doelker,Invitroextended 53a. E. Alleman, J. C.Leroux,R.Gurny,and release properties of drug loaded poly (DL-lactic acid) nanoparticles produced by a salting-out procedure, Pharm. Res., 10:1732-1737 (1993). 53b. E. Allemann, R. Gurny,and E. Doelker,Drug-loadednanoparticlesJ. Pharmaceut.BioPreparationmethodsanddrugtargetingissues,Eur. pharmaceut., 39:173-191 (1993). 54. c. G. Pitt, T. A.Marks,andA.Schindler,Biodegradabledrugdelivery systems based on aliphatic polyesters: Applications to contraceptives and narcotic antagonists, in Controlled Release of Bioactive Materials (R. W. Baker, ed.), Academic Press,New York, 1980, p. 36. 55. C. G. Pitt, M. M. Gratzl, G. L. Kimmel, et al., Aliphatic polyesters11. The degradationof poly (DL-lactide), poly (e-caprolactone) and their copolymers in vivo, Biomaterials, 2:215 (1981). 56. S. Izumikawa, S. Yoshioka, Y. Aso, and Y. Takeda, Preparations of poly (1lactide) microspheresof different crystalline morphology and effect of crystalline morphology on drug release rate, J. Control. Rel., 15:133-140 (1991). 57. J. Heller, Bioerodible systems, in Medical Applications of Controlled Release (R. S. Langer and D. J. Wise, eds.), CRC Press,Boca Raton, FL, 1984, pp. 1-23. 58. P. Le Corre, P. Le Guevello, V. Gajan, et al., Preparation and characterization ofbupivacaine-loadedpolylactideand polylactide-co-glycolide microspheres, Int.J. Pharmaceut., 107:41-49 (1994). 59. H. B. Hopfenberg, Controlled release from erodible slabs, cylinders, and spheres,inControlledReleasePolymericFormulations(D.R.Pauland F. W. Harris, eds.), American Chemical Society, Washington,DC, 1976, pp. 182-194. 60. B. W. Wagenaar andB. W. Muller, Piroxicam release from spray-dried biodegradable microspheres, Biomaterials, 15:49-54 (1994). 61. M. Iwata andJ. W. McGinty, Dissolution, stability, and morphological properties of conventional multiphasepoly(DL-lactic-co-glycolicacid) microspheres containing water soluble compounds, Pharm. Res., 10:1219-1227 (1993). 62. J. J. Burger, E. Tomlinson, E. M. A. Mulder,and J. G. McVie,Albumin microspheres for intra-arterial tumour targeting: I. Pharmaceutical aspects, Int. J. Pharm., 23:333-344 (1985). 62. R. Narayani and K. P. Rao, Controlled release of anticancer drug methotrexate from biodegradable gelatin microspheres, J. Microencaps., 11:69-77 (1994). 64. L. Illum,N.F, Farraj, A.N. Fisher, et al., Hyaluronic ester microspheres aas nasal delivery system for insulin,J. Control. Rel., 29:133-141 (1994). 65, G. A. Magee, N. Willmott, and G. W. Halbert, Development of a repro-

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duceableinvitromethodforassessingthebiodegradation of protein microspheres,J. Control. Rei., 25:241-248 (1993). A. Jayakrishnan, W. A. Knepp, and E. P. Goldberg, Casein microspheres: preparation and evaluation as a camer for controlled drug delivery, Int. J. Pharmaceut., 106:221-228 (1994). 0.P. Rubino, R. Kowalsky, and J. Swarbrick, Albumin microspheres as a drug delivery system: relation among turbidity ratio, degree of crosslinking, and drug release, Pharm. Res., 10:1059-1065 (1993). V. Dilova and V. Shishkova, Albumin microspheres as a drug delivery system for dexamethasone, Pharmaceutical and pharmacokinetic aspects, J. Pharm. Pharmacol., 45:987-989 (1993). N. Yui, J. Nihira, T. Okana, and Y. Sakurai, Regulated release of drug microspheres from inflammation responsive degradable matrices of crosslinked hyaluronic acid, J. Control. Rel., 25:133-143 (1993). 0.Saslawski,C.Weingarten, J. P. Benoit,and P. Couvreur,Magnetically responsive microspheres for the pulsed delivery of insulin, Life Sci., 42:15211528 (1988). et al.,BiodegradablemiJ. H.Eldridge, J. K. Staas, J.A.Muelbroek, crospheres as a vaccine delivery system, Mol. Immunol., 28:287-294 (1991). S. Pulapura,C.Li,and J.Kohn, Structure-property relationships for the design of polyiminocarbonates, Biomaterials, 11:666-678 (1990). P. Wuthrich, S. Y.Ng,K.V.Roskos,andJ.Heller,Pulsatileanddelayed release of lysozyme from ointment-like poly-(ortho esters), J. Control. Rel., 21:191-200 (1992). J. Y. Armand, F. Magnard, J. Bouzon, et al., Modelling of drug release in gastric liquid from spheric galenic forms with Eudragit matrix, Int. J. Pharm., 40~33-41 (1987). H. Liu, P. Magron, J. Bouzon, and J. M. Vergnaud, Spherical dosage form with a core and shell. Experiments and modelling, Int. J. Pharmceut., 45:217227 (1988). G. W. Sinclair and N. A. Peppas, Analysis of non-Fickian transport in polymers using simplified exponential expression, J. Membrane Sci., 17:329-331 (1984). P. L. Ritger and N. A. Peppas, A simple equation for descriptionof solute release. 11. Fickian and anomalous release from swellable devices, J. Control. Rel., 2:37-42 (1987). C. Lanczos, Applied Analysis. Prentice-Hall, Englewood Cliffs, NJ, 1957, pp. 272-280. S. Y. Lin, In vitroreleasebehaviour oftheophyllinefromPIB-induced ethylcellulose microcapsules interpreted by simple mathematical functions, J. Microencam.. 4213-216 (1987).

8 Nanoparticles: Preparation and Characterization Patrick Couvreur, Guy Couarraze, Jean-Philippe Devissaguet, and Francis Puisieux Universitk de Paris-Sud, URA CNRS 1218, Chdtenq-Malaby, France

I. Introduction-Definitions 11. Nanoparticles Obtained by Polymerization of a Monomer

A. Nanospheres B. Nanocapsules

183 184 184 191

111. Nanoparticles Obtained by Dispersion of Preformed

Macromolecules A. Nanospheres B. Nanocapsules C. Drug Loading and Release

IV. Summary

1.

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INTRODUCTION-DEFINITIONS

Nanoparticles as drug colloidal carriers have attracted more and more interest in the last decade, since these systems were found to be able to increase efficacy and/or to reduce toxicity of potent drugs [1,2]. More recently, they have even been found to be efficient vectors to deliver synthetic fragmentsof DNA into cells [3]. 183

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Nanoparticles may be defined as submicron (
NANOPARTICLES OBTAINED BY POLYMERIZATION OF A MONOMER

A. Nanospheres

Most of the methods for preparing nanospheres based on thepolymerization of monomers consist of introduciog a monomer intothe dispersed phase of an emulsion, an inverse microemulsion, or dissolved ina nonsolvent of the polymer. The polymerization reactions in these systems take place in two phases: a nucleation phase followed by a growth phase(propagation). I . Poly (methylmethacrylate) and Polyacrylamide Nanoparticles Poly(methylmethacry1ate)nanospheres have been produced by a radical dispersion polymerization in whichthe monomeris dissolved in an aque-

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ous phase [4]. The polymerization is initiated by y-irradiation from amCo source (500 Krad yrays) by heating at elevated temperature or by using a chemical intitiator (potassium peroxodisulfate) [5]. The oligomers formed precipitate to form aggregates which may be stabilized by surfactant molecules. The final polymer particles are obtained by the growthof aggregates. In this process, the molecularweight as well as the particle size increase with increasing monomer concentration, decreasing initiator concentration, and decreasing temperature [6]. Polycrylamide nanospheres were proposed by Birrenbach andSpeiser [7] in the 1970s. They were prepared by an inverse microemulsionpolymerization mechanism. Acrylamide was usedas a monomer and N’N’-methylene bis-acrylamide as a crosslinking agent. In thksystem, microemulsions consisted of monodispersed spherical swollen aqueous micelles stabilized by a layer of surfactant molecules and dispersed in an organic phase. The nucleation process is continuous, and it occurs throughout the duration of the polymerization reaction [8]. Thus, the numberof polymer particles formed increases steadly with time but their size remains constant. The polymer particles result from the fusion of several micelles by collision, which occurs during the growth state of the nucelated micelles [9]. By this method, Birrenbach and Speiser [7] succeeded in preparing polyacrylamide nanospheres in the presenceof antigens for the developmentof vaccine adjuvant. Like for poly(methylmethacry1ate) nanospheres, the polymerization was initiated by y-irradiation which is a drawback for the entrapmentof unstable drugs that are easily degraded under a needed dose of 0.5 Mrad. But the main disadvantage of those nanoparticulate systems wasthat they could not be used for intravascular administration because of the nonbiodegradability of the polymers used.

2. Polyalkylcyanoacrylate Nanospheres To open wider prospects for nanoparticles, nanospheres consisting of polyalkylcyanoacrylates (PACA) were developed by Couvreur et al. [10,11]. These polymers,which have beenused for several years as surgical glues, are bioresorbable. a. Preparationand characterization. Preparation of PACA nanospheres can be achieved by emulsion polymerization, in which droplets of water[121. Anionic polyinsoluble monomers are emulsified inan aqueous phase merization takes place in micelles after diffusion of monomer molecules through the water phase andis initiated by negatively charged compounds. Classically, the cyanoacrylic monomer is added to an aqueous solution containing both dextran 70 (1%)as a protectant of colloids and glucose (5%) for isotonicityin MHCl.The polymerization of the alkylcyanoacrylate monomers occurs spontaneously at room temperature with stir-

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ring, leading to the formation of 180 nm (220 nm) colloidal nanospheres. Acid polymerization medium is necessaryto avoid too quick a polymerization of the monomer, which can lead to unwanted agglomerates. Drugs can be combined with nanoparticles after dissolution in the polymerization medium eitherbefore the introduction of the monomer or afterits polymerization. The pH of the final suspension is then adjusted to pH 7, and nanospheres can be freeze-dried with their drug content. Various studies have shown that after the freeze-drying process, PACA nanospheres canbe easily resuspended in water [13]. Furthermore, comparative particle size measurements showed no significant modification of the carrierdimensions after freeze-drying. It has been shown that such a formulation was very reproducible regarding s u e and drug adsorption rate even after preparation at a semi-industrial level [13]. Moreover, when prepared under sterile conditions in an aseptic roomunder laminar flow,cyanoacrylic nanospheres met the usual requirements neededfor intravenous administration, such assterility and lack of bacterial endotoxins. Concerning the morphological characterization of PACA nanospheres, scanning electron microscopy (SEM) generally shows spherical particles of a diameter of about 200 nm [14]. After freeze-fracturing, it was observed that these particles consisted of a solid core, whose inner structureappeared highly porous [15]. The polymer which constitutes the nanospheres doesnot appear as a hollow envelope but rather asa ball with a dense polymeric network. The specific surface area of polybutylcyanoacrylate (PBCA) nanospheres was calculated by Kreuter [l61 to be 106.5 mVg. Other physicochemical parameters of cyanoacrylic nanospheres were measured [161. Polycyanoacrylates had a slight negative charge with a tendency for a decrease in charge with increasing ester side chain length. Furthermore, serum yielded an even more pronounced decrease in surface charge, which was caused by the adsorption of the serum contents, demonstrating a significant interaction of these components with nanospheres. The water contact angle decreased with decrasing hydrophobicity in the following order: PBCA > polyethylcyanoacrylate (PECA) > polymethylcyanoacrylate(PMCA) [16], Using a gel-permeation chromatographic method, it was shown that PACA nanospheres appearto be built by an entanglement of numerous small oligomeric subunits rather than by the rolling up of one or a few long polymerchains [17]. Chromatographic profiles suggested an uniform distribution of molecules with unexpected low molecular weights ranging between 500 and 1000. Furthermore, no monomeric residues were found. The low molecular weight of oligomeric subunits is consistent with later-discussed observations concerning excretion and metabolization of nanoparticles. The most significantadvantage of

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alkylcyanoacrylate over otheracrylic derivatives previously used for preparing nanospheres is in the polymerization step. In contrast to other acrylic derivatives, requiring an energy input for this process that can affect the stability of the adsorbed drug, alkylcyanoacrylates can be polymerizedeasily without suchcontribution.

b. Drug incorporation and adsorption. A successful nanoparticle system may be one which has a high loading capacity to reduce the quantity of carrier required for administration. Two theoretical curves can be proposed to describe the adsorption of drugs onto nanoparticles: Langmuirian-type and constant partitioning-type-isotherms. In fact, it was found that nanoparticles can entrap a drug according to a Langmuirian adsorption mechanism, because of their large specific area [18]. The drug can either be incorporated into nanospheres during the polymerization process or beadsorbed ontothe surface of preformed particles. In the former case, the drug-polymer interaction may result in the covalent linkage of the drug with the polymer. This was observed with vidarabine, an antiviral agent, whose nucleophile N in positions 3 and 7 may playa role of initiator for the anionic polymerization mechanism of the cyanoacrylic monomer [19]. To a lesser extent, a similar interaction was described with peptidic compounds such as growth hormone-releasingfactor (GRF) [20]. In fact, with GRF, it was shownthat if the drugwas added very early after the beginning of the polymerization process (5 min), 50% of the peptide was found covalently linked to thepolymer. On thecontrary, when the peptidewas added later on(i.e., 60 min after the beginning of the polymerization), it was not chemically modified, but the loading capacity was poor. Thus, it was found that thereis a narrow window of time forthe addition of GRF to thepolymerization medium which results in both the preservation of the chemical structure of the peptide and a satisfactory drug-loading capacity [20]. In contrast, surface pressure and surface potential studies of polyisobutylcyanoacrylate monolayers spreadat theair-water interface in the presence of ampicillin revealed that theassociation which forms ampicillin with the polymer was weak and could be easily disrupted [21]. Based on the surface potential and surface pressure experiments, models of polyisobutylcyanoacrylate-ampicillin arrangements in the interfacial region have been proposed for the low- and high-polymer surface densities [22]. Concerning loading capacity of cyanoacrylic nanospheres, it was found that both the nature and quantities of the monomer used influenced the adsorption capacity of the carrier. Generally, the longer the alkyl chain length, the higher is the affinityof the drug. Taken in its entirety, the capacity of adsorption is related to the hydrophobicity of the polymer and

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to thespecific area of the carrier. Moreover, the percentage of drug adsorption generally decreases with the quantity of drug dissolved inthe polymerization medium, according to theLangmuirian isotherm[23]. Special attempts have been made for the association of synthetic fragments of DNA to PACA nanospheres [3]. In fact, the association of antisens oligonucleotides with nanoparticles was possible only inthe presence of a hydrophobic cation, such as triphenylphosphonium (Fig. 1) or quaternary ammoniumsalts. The poor yield of oligonucleotide association without cations could be explained by the hydrophilic character of nucleic acid chains that areknown to be soluble in water. In the proposed method, oligonucleotide adsorption on the nanosphereswas mediated by the formation of ion pairs between the negatively charged phosphate groupsof the nucleic acid chain and the hydrophobic cations [3]. The adsorption efficiency of oligonucleotide-cation complexes on nanospheres was found to be highly dependent on several parameters: oligonucleotide chain length, nature of the cyanoacrylic polymer, hydrophobicity of the cations used as ion-pairing agents, and ionic concentration of the medium [3]. When adsorbed onto nanospheres, oligonucleotides were found to be protected from the degradationby a 3'-exonuclease in vitro [3].

0

50

100

Hydrophobic ion

150

(PM)

Fig. 1 . Adsorption of oligothymidylate to polyalkylcyanoacrylatenanoparticles. Oligothymidylate (2 mM) complexed with increasing concentrations of positively charged tetraphenylphosphonium chloride (PP)was added to PIBCA (W) or PIHCA (+) nanoparticles (10 mg/mL). Adsorption of oligothymidylate to PIHCA nanoparticles in the presence of the negatively charged tetraphenylboron sodium (0)was measured under the same experimental conditions, as a control. (From ref. 3.)

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c. Biodegradationanddrug

189

release. Another characteristic of cyanoacrylate polymers is that their rate of degradation is dependent on the length of their alkyl chain. The dominating mechanism of particle degradation was found to be a surface erosion process [24]. This process occurs through the hydrolysis by enzymes of the ester side chain of the polymer [25]. By this mechanism, the polymeric chain remains intact, butit gradually becomes more and more hydrophilic until it is water soluble. This degradation pathwayis consistent with the productionof alcohol during the bioerosion of PACA nanospheres in vitro in the presenceof esterases [25]. Indeed, the action of rat liver microsomes and tritosomes on the ester hydrolysis of polyisobutylcyanoacrylate nanospheres was clearly demonstrated [25]. A relationship between isobutanol production (i.e., nanosphere bioerosion) and esterase concentration was found. Therefore, we consider that the production of formaldehyde via chemical degradation (reverse Knoevenagel reaction) as proposed by several investigators [26] plays a marginal role in the overall degradation of PACA nanospheres under physiological conditions. Indeed, the amount of formaldehyde produced after nanospheredegradation was found to be5% of the theoretical quantity that would have been producedif the polymer had been entirely degrated by this pathway [25]. The suggestion of Vezin and Florence [26] that esterases do not affect the degradation rate is therefore contested. Since the biodegradability of polyalkylcyanoacrylate depends on the nature of the alkyl chain, it is possible to choose a monomer whose polymerized form has a biodegradability corresponding to theestablished program for drugrelease [14]. Indeed, by using a double radiolabeled technique (14Clabeled nanospheres loaded with [3H]actinomycin), it was found that drug was released from nanospheres as a direct ,consequence of the polymer’s bioerosion [25]. This was confirmed using GRF, another drug model[20]. With this peptidic compound, in the absence of esterases, no drugrelease was observed, whereas in the nanospheres suspension turbidity remained unchanged, indicating no polymerbioerosion. Drug release appeared to be dependent on the esterase content of the incubation medium. The liberation of GRF was quicker in rat plasma (content of esterases around 150 @mL) than in Ringer lactate medium containing esterases (100 pg/mL). At the same time, the turbidity (i.e., the expression of nanosphere bioerosion) decreased with the release of the peptide (Fig. 2). That the GRF release directly resulted from the polymer’s bioerosion was confirmed by the fact that themore slowly bioeroded polymerpolyisohexylcyanoacrylateyielded the slowest decrease of turbidity and the most progressive release of the peptide. Finally, the fact that the drugrelease was related to the turbidity decrease suggested that GRF was homogeneouslydispersed throughout the polymeric matrix rather than adsorbed atthe surface of the polymer.

Couvreur et a/.

790

GRF

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Time (h) Fig. 2. Release of GRF from PIHCA nanoparticles is the consequence of polymer's bioerosion (decrease of the turbidity of the suspension). (From ref. 20.)

With rose bengal, a biphasic release from PACA nanospheres was observed [18]. The release profile of this compound was described by using a simple biexponential function: Pt = Ake-a

+ Be-b'

where Pt is the concentration of compound remaining in the nanosphere at time, t; the concentrations Aand B and the two rate constantsa and b can

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be used to characterize the release of the compound. The initial release rate was faster when the marker was adsorbed only at the surface of the carrier, whereas it was reduced when the compound was incorporated into the polymeric networkof the nanospheres. By employing dextransulfate as the stabilizer in the preparation of PACA nanospheres, a slower release rate was obtained for the model drug rose bengal [27]. Thus, with this compound, the release was not only dependent on the kinetic of bioerosion of the polymer. B. Nanocapsules

After the development of nanospheres obtained by emulsion polymerization of alkylcyanoacrylates as described in the preceding section, it has been possible using the same monomer to manufacture particles of nanometric size containing an oil-filled cavity. This procedure, which was first developed by Al Khoury et al. [28], has beenapplied for insulin delivery byDamge et al. [29].The characterization of polyalkylcyanoacrylate nanocapsules has also been performed by different investigators [30-321. I . PreparationandCharacterization Nanocapsules of PACA are obtainedvia an interfacial polymerization process in emulsion. Nanocapsules are formedby mixing an organic phase with an aqueous phase. The organic phase is classically an ethanolic solution of the monomer mixed together with the oily core material and the lipophilic drug to be encapsulated and to which, in some cases, soya bean lecithin is added as an additional surfactant. Oils which have been used to prepare nanocapsules include Miglyol, benzylbenzoate, ethyl oleate, and Lipiodol. The encapsulation efficiency of a lipophilic drug generally depends on its partition coefficient between the oil and the aqueous phase,so the oil must be chosenaccordingly. The aqueous phase is a solution of a nonionic surfactant, usually Synperonic PE-F68 at 0.5% at a pH between 4 and 10. To form nanocapsules, the organic phase is allowedto runslowly-for example, through a wide-bore syringe needle or a micropipette tip-into the aqueousphase, which is magnetically stirred. The mixture immediately becomes opalescent. After stirring for 15-30 mins, the ethanolis removed by evaporation under reduced pressure. If required, the nanocapsules canbe furtherconcentrated by evaporation; this allows them to be rediluted in a physiological buffer for injection. Nanocapsules formedin this way have a mean diameter between200 and 300 nm witha narrow polydispersity. The speed of the magnetic stirring has noinfluence on theparticle size, which depends solely on the nature and

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the volumeof the oil and on thevolume of the diffusing organic phase. The proportion of oil to monomer must be correctly chosen to avoid simultaneous formation of either flakes of polymer or of a single oily emulsion. The presence of a surfactant in the aqueous phase is, in fact, not necessary for thesuccessful formation of nanocapsules, but it does prevent them from aggregating on storage to a cake which is difficult to disperse. Nanocapsules formedin this way are physically stable for several years at ambient temperature andmay be sterilized by autoclaving at 120°C for 20 min (the effect of such a procedure on any encapsulated drug must, of course, be considered.) However, as a result of their vesicular character, nanocapsules are not easily lyophilized, since they tend to collapse releasing the oily core. In fact, many factors could influence the type of colloid formed by anionic interfacial polymerization ofPACA: the nature of the aqueous phase, the pH, thecomposition of the organic phase (monomer, oil, ethanol), the ratio of monomer to the aqueous phase, and the emulsification conditions. The degree of polymerization and, therefore, the molecularweight, depend ona balance between initiation, propagation, and termination [33]. The number of growing chains depends on the concentration of initiators (OH-, CH,O-, CH,COO-, CN-). Forthe same quantity of monomer, when the numberof live chains is high, the degreeof polymerization (DP) is low. Similarly, DP is reduced when the concentration of terminating agents (H+) is high [34,35]. The concentration of the initiating and the terminating agents available for the monomer depends on thephysicochemical nature of the system in which the componentsare dispersed at themoment of the formation of the colloid. In thecase of a system composed of two immiscible phases, such as an emulsion, the presence and theconcentration of the solutes in the different phases is a function of the polarity of the solute molecules and of the dielectric constant of the medium. In contrast, at an interface between an aqueous and an organic medium, there can be large local variations in properties which can themselves cause changes in the interfacial properties in a dynamic system. For example, spontaneousemulsification depends on the interfacial tension. Thus, although the degree of polymerization depends on the propagation reaction, other characteristics of the colloid formed suchas particle size and morphology depend on interfacial phenomena inducted by the dynamic mixing of an organic phase with an aqueous phase. Thus, when the quantity of ethanol was increased, there was a tendency to form a suspension of polymeric flakes.However, with an increase in the volume of oil, an oily layer appeared onthe surface of the suspensions.

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For systemshaving an oiVethano1ratio of some percentage, the macroscopic aspect is markedly more homogeneouswith the presence of a majority of nanocapsules. Since the preparation of nanocapsules of PACA is rather similar to the preparation of PACA nanospheres, it is necessary to verify that the colloidal suspension does not consist of a simple mixture of an oil-in-water emulsion containing polymeric nanospheres of similar size. Several different experiments confirmed that the preparation was indeed composed of oil-filled nanocapsules [32]. First, turbidity measurements showed a clear difference between nanocapsules and a mixture of nanospheres and the emulsion containing the same amount of polymer and oil. Second, the vesicular nature of nanocapsules prepared in this way was confirmed by electron microscopy. Scanning electron microscopy showed a smooth exterior to thenanocapsules,whereasthethincapsulewallandtheinternal cavity could be visualized by transmission electron microscopy [28]. Fig. 3 shows a polyisobutylcyanoacrylate nanocapsule after cryofracture andobservation by transmissionelectronicmicroscopy. The membrane thickness ranges from 5 to 10 nm. 2.

Mechanism of Formation Theoretically, nanocapsules result from the interfacial polymerization of the cyanoacrylic monomersat the surface of the oil droplets dispersed in the aqueous phase. A drugcan be encapsulated in the internalcavity of the polymerized membrane during the formation of the system provided that its oil-water partition coefficient is infavor of the oily phase. Nevertheless, a simple emulsification of an organic phase, containing alkylcyanoacryrate monomers and oil, in an aqueous phase does notallow nanocapsule formation. As a matter of fact, because of the affinity of alkylcyanoacrylates for the oily phase, a bulk polymerization of the monomers in the oil droplets occurs preferentially. Polymeric matrix systems, which can be compared with nanospheres in which the oil is dispersed throughout the polymeric network,result. To obtain nanocapsules, a dynamic process must be created which brings the monomer to the oil-water interface. This transfer is performed by the diffusion of a cosolvent from the organic phase to the aqueous phase. This cosolvent must be a solvent for the monomer and forthe oil on one hand and miscible with the aqueous phase on theother hand. Usually, ethanol can be used as diffusing solvent. So, nanocapsules are prepared by emulsificationof an organic phase containing the monomer in an aqueousphase. The main point is the compositionof the organic phase in which the oil and the diffusing solvent are mixed together. In the case of nanocapsules, the natureof the emulsification process is spontaneous.

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Fig. 3. Polyisobutylcyanoacrylate nanocapsulesobserved by transmission electron microscopy after cryofracture. (From ref.32.)

In addition to their vesicular structure, the nanocapsules made from alkylcyanoacrylates differ from nanospheres by their size, the molecular mass of the polymer, and the characteristics of the drug release. Usually, nanocapsules are larger than the corresponding nanospheres and the degree of polymerization of the polymer is also significantly higher. These differences are theresult of the specific mechanismof nanocapsule formation. This mechanism is complex, becauseit results from boththe chemical characteristics of the polymerization and the physical characteristics of the polymerization medium (viscosity, interfacial tension, and cosolvent diffusion from the organic phase to theaqueous phase). These processes usually lead to theformation of mixed systems where nanocapsules coexist with other polymeric particles. Figure 4 shows photographs takenby transmission electronic microscopy of a nanocapsule preparation after separation by ultracentrifugation. Particles of lower density are true nanocapsules containing an oily cavity;

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(b)

Fig. 4. Transmission electron micrographs of a nanocapsule preparation after separation by ultracentrifugation. (a) Floating layer.(b) Pellet. particles of higher density are in fact nanospheres with a size comparable to that of nanospheres obtained by emulsion polymerization. Nevertheless, the mechanism of formation of these nanospheres is not anemulsion polymerization, because the molecular mass of the polymer of these particles is high, similar to that associated with nanocapsules (about 100,000 versus about 1000 for nanospheres) [31]. Since the nanospheres obtained during the formation of nanocapsules have the same polymeric molecular mass as the nanocapsules, it is possible to conclude that thepolymerization mechanism is the same. Froma chemical point ofview, the anionic polymerization mechanism of alkylcyanoacrylate is not very much modifiedduring the formation of nanocapsules as

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compared with nanospheres. It could be possible to consider that the nucleophilic initiation of alkylcyanoacrylate polymerization, which is normally performed by the hydroxyl ions of water, could be performed by ethoxy ions, although the dissociation constant of ethanol is very low. In fact, polymerization of alkylcyanoacrylates in the presence of ethanol is very restricted and leads to the formation of small oligomers with low molecular mass. Although the presence of ethanol seems not to modify the polymerization of the monomer from the chemical point ofview, the diffusion of ethanol is the major physical factor which imposes the conditions of polymerization and, thus, those of nanocapsules formation. Indeed, when the total volume of ethanol is kept constant, but is initially distributed differently between the organic phase and the aqueous phase, it appears thatthe molecular mass of thecreated polymer doesnotdependon the total amount of ethanol but only on that part of the ethanol diffusing from the organic phase to the aqueous phase. The very high molecular mass of the polymer making up the nanocapsules, is only obtained when the diffusing fraction of ethanol is more than half the total.The role of ethanol diffusion in the formationof nanocapsules is illustrated in Fin. - 5 . The rapid diffusion the aqueous phase leads to primary of ethanol from the organic phase to

Aqueous phase

2

Nanocapsules

/ 0

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fragmentation of thepolymeric film

Marangoni effect:

69 @ @ Nanoparticles

interfacialpolymerization of monomer

Fig. 5. Mechanism of formation of nanocapsules

by interfacial polymerization.

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convection currents, which are perpendicular to the interface, yielding an higher concentration of monomers on the peak of the surface undulations of the interface between the organic and the aqueous phases. Since alkylcyanoacrylate monomers have tensioactive properties, interfacial tension gradients appear on the interface, leading to secondary convection currents at a tangent to the interface (Marangoni effect). So primary and secondary convection currents promote spreading of the monomer at the interface where an interfacial polymerization will be able to propagate. This kind of polymerization leads to a higher molecular mass than ina single emulsion polymerization where the oligomers, being widelydispersed in the aqueousphase or in the micelles, have a greater probability of meeting cationic species (H,O+) which terminate the polymerization. In contrast, during interfacial polymerization, the oligomers are relatively protected from termination agents and the polymerization can develop owing to the monomer reservoir set up within the organic phase. The interfacial turbulences due to the secondary convection currents of the Marangoni effect initiate the fragmentation of the interfacial polymeric film and initiate the spontaneousemulsification of the system. During this emulsification, fragments of films fold back on themselves at the,interface around the oil droplets to form the nanocapsules, whereas the remainder of the film fragments lead to theformation of nanospheres of smaller size with a polymer molecular mass retaining the characteristics of interfacial polymerization (high molecular mass). The structure of the mixed systems (nanospheres and nanocapsules) formed during the preparation of nanocapsules is confirmed by the analysis of the drug-release kinetics. Fig. 6 shows the comparison of the in vitro release of tetracaine from polyisobutylcyanoacrylate nanospheres (true nanospheres) and from the two fractions (obtained after separation by ultracentrifugation) of the nanocapsule suspension (true nanocapsules and nanospheres). The similarity of the release kinetics of the two nanosphere systems confirms that they have an identical structure (matrix structure). Analysis of the release kinetics of nanocapsules, compared with those of nanospheres, shows that thetime constant of nanocapsules is superior to that of nanospheres which wouldhave the same diameter. This is coherent with the vesicular structure of the nanocapsules. 111.

NANOPARTICLES OBTAINED BY DISPERSIONOF PREFORMED MACROMOLECULES

Nanospheres A.

As presented above, nanoparticles can be obtained by inducing the polymerization of a monomer in certain conditions. However, many polymeric

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I

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nanwpsulcs obtained

0 nanosphcres obldncd

by inlerfacirl polymubtlon

* 0

2

h u e nannosphcrcs obtained

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6

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Fig. 6. Invitroreleasekinetics nanoparticles.

of tetracain from polyisobutylcyanoacrlate

materials not capable of being prepared by emulsion polymerization, such as polyurethane, epoxy, polyester, and others,including semisynthetic polymers such as cellulose derivatives, remained unavailable for aqueousdispersion. This problem has led to the development of pseudolatexes (artificial latexes) obtained by dispersion of preformed polymers. For drug delivery purposes, following an intravenous injection, the polymeric material needs to meet physicochemical and biological requirements adapted and optimized for this specific application. Among these requirements, submicron size to avoid the risk of embolization following intravenous injection and biodegradability to nontoxic metabolites are of crucial importance, thus limiting the number of available polymers. If the method of preparation implies the use of nonsafe ingredients (solvents, stabilizers), steps of purification should be added in order to assure the required qualities of the final product. Most of the synthetic and artificial latexes preparedfor industrial applications did notmeetthese requirements,and various approaches were progressively developed

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in order to obtain polymeric nanospheres fulfilling the pharmaceutical criteiia. Polymeric nanospheres prepared by emulsion polymerization may encounter somedrawbacks: With the exception of alkylcyanoacrylate, most of the monomers suitable for a micellar polymerization in an aqueous phaselead to slowly or nonbiodegradable polymers. The polymerization process is mainly limited to the vinyl addition reaction, and the molecular weight of the polymeric material cannot be fully controlled. Residues in the polymerization medium (monomer, oligomer, organic solvent, surfactant, initiator of polymerization) can be more or less toxic and may necessitate further purification of the colloidal material. During the polymerization process, activated monomer molecules may interact with the drug molecules, thus leading to theirinactivation (20). In order to avoid some of these limitations, the methods of obtaining nanospheres from various preformed macromolecular materials, whose physicochemical and biological properties can be well controlled, have been developed. Two approaches using the emulsification of a solution of the macromolecular material or using a controlled desolvation of this material have beensuccessively considered.

1. NanospheresPrepared by Emulsification The methods of preparing nanospheres byemulsification were adapted from theindustrial techniques available for obtaining the artificial latexes used in surface-coating applications (e.g., paints, adhesives, textilesizing, paper coating). Initially, artificial latexes were developed as waterbased dispersions in order to avoid economical and environmental problems encountered with the use of organic solvent-based coatings. They present the major advantage that any water-insoluble polymeric material can be used and all the industrial techniques for obtaining artificial latexes rely on oneof the threefollowing processes: 1. Solution-emulsification [36]: The polymer is dissolved ina volatile solvent immiscible with water. This organic phase is dispersed in an aqueous phase by conventional emulsification techniques and appropriate emulsifiers. The polymeric particles are formed from the oil-in-water ( O W ) droplets by removing the organic solvent following its vaporization.

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2. Phuse-inversion [37]:The polymer is first ground with a fatty acid. An alkaline aqueous solution is then added and mixed to form a water-in-polymer dispersion. Further addition of water leads to the inversion of phases and to a dispersion of the polymer in the aqueous solution. In this case, the emulsifier is the soap formed from the reaction between thefatty acid and the alkali. 3. Self-emulsification [38]:Some polar groups such as amine or quaternary ammonium, when attached to polymer molecules, may confer autoemulsioning properties and remove the need for any added emulsifier for the dispersion of the polymerin an aqueous phase. However, the presence of polar groups may also modify the water sensitivity, more specifically the pH sensitivity, of the colloid.

If pharmaceutical applications are considered, the emulsification techniques may present some limitations:

To obtain small (
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tion of some vegetal proteins such as zein or gluten, they need physical or chemical modifications to become water insoluble.

Nunospheres of synthetic polymers. Gurny et al. [41] and Krause et al. [42] have prepared poly(1actide) nanospheres according to thesolutionemulsification technique. The polyester is dissolved in an organic volatile solvent immiscible withwater, such as chloroform, and the organic solution is dispersed in an aqueous phase to form an O N emulsion. Continuous emulsification under mixing prevents coalescence of organic droplets and allows the spontaneous evaporation of the solvent at room temperature and the formation of the colloidal particles. Residual organic solvent is removed under reduced pressure. The emulsifiers can, if necessary, be removed by dialysisof the suspension or by washings following the separation of nanospheres by ultracentrifugation. Ibrahim et al. [43] and Allemann et al. [ M ] have proposed replacing water-immiscible solvents, such as chloroform (all trace of which must be eliminated for aninjectable dosage-form) by a miscible one such as acetone, which can easily be removed by evaporation. The method consists of adding under continuous stirring to an acetonesolution of polymer an electrolyteor a nonelectrolyte-saturated aqueous solution containing poly(viny1 alcohol) (PVA) as a viscosity agent and stabilizer. The saturated aqueous solution prevents acetonefrom mixing withwater by a salting-out process and a W/O emulsion is obtained. Addition of water to this emulsion allows complete diffusion of acetone in theaqueous phace inducing the formation of nanospheres. Subsequent washings by ultracentrifugation or dialysis allow the elimination of the salt or of the nonelectrolyte used to trigger the saltingout process. One of the investigator’s claim is that surfactants are notnecessary to obtain nanospheres, since a protective colloid is used. However, when injectable surfactants are available the presence of PVAremains questionable if the dispersion is intended to beinjected. For that reason, poly(1actide) and poly(1actide-co-glycolide) nanospheres were developed by solvent evaporation using serum albumin as a colloidal stabilizer [45]. The characterization of the albumin coating has been studied [46].

U.

b. Nunospheres ofnuturulpolymers. Kramer [47] hasprepared albumin nanospheres according to the emulsification technique. Albumin is first dissolved in water andthis solution is dispersed in cotton oil to form a W/O emulsion. This emulsion is then poured into oil heated to morethan 100°C, thus resulting in an heat-denaturation of the protein and, simultaneously, in the vaporization of the aqueous phase. Subsequent washings and purifications are necessary to yield an aqueous suspension of albumin nanospheres, which can finally behardened by crosslinkage by aldehydes.

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Yoshioka et al. [48] also suggested a method leading to the formation of protein nanospheres at room temperature following the direct crosslinkage of gelatin by an aldehyde added to a W/O emulsion. Schroder and Stahl [49] proposed a similar method (W/O emulsion) to prepare nanospheres from polysaccharides such as dextran and its derivatives.

2. Nanospheres Prepared by Desolvatation As previously mentioned, one drawback of the emulsification methods is the use of emulsifiers and eitherchlorinated solvents or oily phase or saturated salt solutions, which implies more or less tedious purifications to obtain the final aqueous dispersion. Another limitation is the size and the homogeneity of the droplets containing the macromolecularmaterial, and it is almost impossible to get a monodisperse population of nanospheres. Attempts have been made to omit this step of preparation and have led to the development of methods using a controlled desolvation to obtain the colloidal aqueous dispersion of the macromolecularmaterial. Desolvation is a well-known method for isolating macromolecules from liquid media. Thus,polymers can beprecipitated from theirsolutions following the addition of a third component or of a nonsolvent miscible with their solvent. The sameeffect is obtained with an aqueous solution of proteins when neutral salts or alcohol are added. However, the process generally leads to a bulk precipitate of polymeric or protein material rather than colloidal particles. a. Nanospheres of synthetic polymers. Fessi et al. [50] proposed a new, simple, and mild method yielding nanometric and monodisperse polymeric particles without the use of any preliminary emulsification. Briefly, an organic solution of the polymeris prepared and added under moderate stirring to a nonsolvent liquid phase. Both solvent and nonsolvent must havelow viscosity and high mixing capacity all in proportions. As an example, acetone and water meet these conditions. The only complementary operation following the mixing of the two phases is to remove thevolatile solvent by vaporization under reducedpressure. Further concentration of the aqueoussuspension can becarried out under the same conditions or by freeze-drying. The mean diameter of the particles obtained is about 200 nmwith a low dispersity. It was observed thatthe size of the particles was mainlyrelated to the type of polymer used and less to the experimental conditions. This method has been successfully applied to various biodegradable preformed polymers such aspoly(1actide) ,poly(glyco1ide),poly(1actide-co-glycolide), poly(&-caprolactone), poly(alkylcyanoacrylate), and other nonbiodegradable preformed polymers such as poly(acrylic),poly(vinylch1oride-coacetate), ethylcellulose, cellulose-acetophtalate, and poly(styrene). Small

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amounts of surfactants, either lipophilic or hydrophilic, can be added to the solvent or to the nonsolvent (for an injectable suspension, phospholipids and Synperonic PEF68 can used). be However, it was observed that surfactants were notnecessary to obtain the colloidal dispersion (which forms spontaneously) but only to stabilize it and to facilitate its redispersion following caking by sedimentation. It was alsonoteworthy thatthe sense and the rate of mixing of the two phases had no influence, the organic solution of the or vice versa, with polymer beingslowly or rapidly added to the nonsolvent or without the aid of additional stirring. In fact, themost critical conditions for obtaining the spontaneous formationof colloidal particles alone, avoiding any bulk precipitation of the material, were thelow concentration of the polymer in the organic phase and the 1/2 ratio of solvent to nonsolvent volumes. One of the most interesting and practical aspect of this method is its capacity to bescaled up fromlaboratory to industrial amounts. Chang et al. [51] and Bodmeieret al. [52] described a method, similar to that of Fessi et al. [50], to prepare nanospheres without any surfactant byusingacrylic copolymers bearing quaternary ammonium groups. As previously mentioned, these autoemulsioning polymers are water sensitive, more specifically pH sensitive, and they may become soluble, thus limiting both the applications and the choice of the starting material. Byusing carboxylated polymers such as Eudragit S, Fessi et al. [50] were able to obtain colloidal particles by the simple addition of an aqueous alkaline solution of the polymerto anacidic medium. However, subsequent neutralization of the suspension to physiological pH led to a clear solution.

b. Nanospheres of Natural Polymers. Marty et al. [53] have proposed a method of preparing gelatin nanospheres by controlled desolvation from an aqueoussolution. The slow addition of a desolvating agent (neutral salt, alcohol) to a gelatin solution containing a surfactant induces progressive modifications of the protein’s tertiary structure to give a more and more hydrophobic material. The initially clear solution gradually turns opalescent asthe gelatin becomes less soluble in waterandtends to form nanometric aggregates of the desolvated protein, leading to a milky suspension. If salts or alcohol are added in excess, bulk precipitation occurs and the supernatant becomeslimpid once more. Thus, Martyet al. [53] recommended the monitoring of the desolvation process by continuous nephelometric measurements. Gelatin nanospheres formedin this way need to be hardened by crosslinkage with an aldehyde, the excess of which is neutralizedwith a sulfite. Finally,purification to remove desolvating agent, surfactant, and other reactants is carried out by dialysis or by washings following ultracentrifugation. The elimination of the desolvating agent can be by vaporization if alcohol is used. This method presents some draw-

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backs, such as the monitoring of the desolvating process and the need for purification to eliminate various added materials. Stainmesse et al. [54] proposed a simpler, single-step technique to prepare protein nanospheres. Adapted from the method developed for polymeric nanospheres[50], the technique consists of pouring an aqueous solution of the protein (i.e., albumin) into anonsolvent. In particular, the nonsolvent can be ethanolor boiling water, thus leading to heat denaturation of albumin and to the formation of colloidal particles as small as 80 nm. Further strengthening of albumin nanospheres can be obtained either by heat sterilization (121°C) or by crosslinkage. It is noteworthy that no additive (surfactant, protective colloid) was necessary to obtain a stable dispersion, owing to the small and monodisperse size of albumin nanospheres, even under theconditions of heat sterilization. The sameprocess was usedto obtain an aqueousdispersion of a water-insoluble protein, such as zein, by dissolving the proteinaceous material in an organic solvent, such as ethanol. B. Nanocapsules

Nanocapsules present the unique characteristic of being a solid vesicular system. The polymeric wall renders nanocapsules much more stable than nanoemulsions andliposomes, whereas the inner core allows a large loading capacity according to the solubility of the drug moleculein this core. It might be considered that nanocapsules could be obtained by adapting some of the various and well-known techniques for microencapsulation to yield smaller particles. However, all attempts to reduce the size of microcapsules to below 1 pm, essentially based on conventional emulsification techniques, were unsuccessful. Two methods of preparing nanocapsules have been successively developed in our group:

1. The first one, which have been previously considered in this chapter, was the method of Al Khoury et al. [28], using an interfacial polymerization of alkylcyanoacrylate monomer. When prepared by interfacial polymerization of alkylcyanoacrylate monomer, nanocapsules may have similar limitation to the one previously mentioned for nanospheres; thatis, the possible chemicalinteraction between the monomer andthe drug molecules [20]. 2. The second one was the method of Fessi et al. [55], using an interfacial deposition of a preformed polymer. The advantages of using well-characterized performed polymers have been previously emphasized and is particularly interesting when perfectly safe and biodegradable polymers mustbe used, such asthe polyes-

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ters already used for surgical sutures, implants, and other biomedical applications. Fessi et al. [S51 were the first to propose a method for obtaining nanocapsules from preformed polymers, and their approach was to apply the principle of spontaneous emulsification. Spontaneous emulsification [56]can occur when a solution of a liquid L1 in a liquid L2 is carefully added at the surface of a third liquid L3, L2 being soluble simultaneously in L1 and L3, whereasL1 is insoluble in L3. Molecules of L1 are progressively transferred to L3, where they are notsoluble, from their solution in L2, while reciprocal diffusion occurs between L2 and L3. This process, called “diffusion and stranding,” allows the formation of droplets of L1 in L3 to give an ultrafine emulsion. Various explanations of this phenomenon, beyond the scope of this chapter, have been given, including localvariations of the interfacial tension between L1 andL3, due to the reciprocal diffusion between L2 and L3, or the Marangoni effect. The method of obtaining nanocapsules [55] was adapted from the method used to preparenanospheres [50].Briefly, the preformed polymer, the phase intended to become the core of the nanocapsule, and the drug molecule to be loaded in the carrier are dissolved in a volatile solvent perfectly miscible with the nonsolvent. This solution is then poured in the nonsolvent of both the polymer and the core, and the resulting mixture immediately turns milky owing to the spontaneous formation of nanocapsules. The solvent can be removed by evaporation at room temperature under reduced pressure. The size of nanocapsules is about 200 nm with a low dispersity ( 5 0 nm). The loading capacity of nanocapsules is related to the solubility of the drug in the core and the maximum loading could be obtainedwhen the core is the pure drug molecule. Smallamounts of surfactants, either lipophilic or hydrophilic, can be addedto thesolvent or to the nonsolvent. However, it was observed that surfactants were not necessary to obtain nanocapsules. The same critical conditions as previously cited for thepreparation of nanospheres by desolvation of preformed polymer apply to the preparation of nanocapsules, that is, the concentrations of both the polymer and the core in the solvent and the solvent to nonsolvent ratio. It should be mentioned here that, in contrast to nanospheres, nanocapsulesoffer the major advantageof a low ratio of polymer to othercomponents. Moreover, nanocapsules can be obtained with either an oily or an aqueous core according to a careful selection of the polymer, the solvent and the nonsolvent. The feasibility and reliability of this method have beenstudied for either laboratory or industrial batches. One of the very intriguing aspects of the method of Fessi et al. [55] is why the mixing step does not lead to the simultaneous formationof poly-

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meric nanospheres on one hand and of a core emulsion on the other. Moreover, themechanism of formation of nanocapsulesis not totally understood, but a partial explanation could be as follows: When the solution of the preformed polymer and the core is poured in the nonsolvent, core droplets are formed by spontaneous emulsification as the solvent diffuses in the nonsolvent. The polymer, insoluble in both the core and the nonsolvent, could be desolvated at theinterface of these two immiscible components. This interfacial polymer deposition following solvent displacement leads the polymer to act as a barrier and to stabilize the emulsion formedby the corematerial. It can be hypothesized that the polymer might playa role analogous to the roleof a surfactant in emulsions.

C. Drug Loading and Release The loading capacity of colloidal delivery systems and theirrelease kinetics, which make the drugmolecule available, are important criteria. Depending on both thepreparation process and the physicochemical properties of the drug molecule and thecarrier, the drug entrapment can be either by inclusion within the carrier and/or by superficial adsorbtion onto this carrier. Electrical charges on both the drugmolecule and thecarrier may influence the loading capacity.

1. DrugLoading Whatever their process of preparation, polymerization of monomers or dispersion of preformed polymer, entrapment within nonporous nanoin the macromolecular spheres requires the solubility of the drug molecule material, whereas porous nanospheres may entrap the drug molecule by adsorption either superficially or within the macromolecular network,The described following the adsorption of drugs ontonanospherescanbe Langmuir-type or the constant partitioning-type isotherms. In fact, nanospheres generally entrap drugmolecules according to a Langmuir adsorption mechanism owing to their large specific surface area. Entrapment within the core of nanocapsules implies the solubility of the drugmolecule in the oily phase. It should be mentioned that the drug to polymer ratio can be as large as 500% in nanocapsules (inner core madeof the drugitself) when this ratio is usually under 10% in nanospheres. 2. Drug Release Drug release from colloidal carriers is dependent on boththe type of carrier and the loading mechanisminvolved. As the specific area of a nanometric dispersion is very large, the release rate may be more rapid than from other larger structures (microspheres and microcapsules) and colloidal carriers are generally not intended to act as sustained-release

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delivery systems. It has to be mentioned that drugrelase data from colloidal delivery systems are highly dependent on the experimentalconditions of the study [57]. More specifically, precautions have to be taken toavoid any artefactual rate-limiting step, particularly the presence of a membrane in a diffusion study or theabsence of sink conditions. a.

Nunospheres. Releasefromnanospheres may be different according

to the drug-entrapment mechanism involved. When the drug is super!% cially adsorbed, the release mechanism can be described as a partitioning process (rapid and total release if sink conditions are met). When the drug is entrapped within the matrix, diffusion plus bioerosion will be involved with a biodegradable carrier, whereas diffusion will be the only mechanism if the carrier is not biodegradable. From this, it can be inferred that entrapment within the matrix of nanospheres may lead to sustained release, the rate of which may be related to the rate of biodegradation of the polymer.

b. Nanocapsules. Releasefromnanocapsules is related to partitioning processes within immiscible phases. The equilibrium between the carrier (loaded drug) and the dispersing aqueous medium (freedrug) is dependent both on the partition coefficient of the molecule betweenthe oily and the aqueous phases and on thevolume ratio of these two phases. This means that the amount released is directly related to thedilution of the camerand that the release is practically instantaneous when sink conditions exist. Diffusion of the drug through the polymeric wall of nanocapsules does not seem to be a rate-limiting step [58]. Drug leakage from nanocapsules can be advantageously reduced by coating the polymeric wall with an outer layer of phospholipids. IV. SUMMARY

From this chapter, it emerges that a number of methods are available today for the preparation of nanoparticles. These methods are based on a wide variety of principles, but theyare generally concerned with either the polymerization of a monomer into an homogeneous or an heterogeneous phase or with the precipitation of a preformed polymer. Some of these processes present drawbacks from technological standpoint, whereas others are probably unsuitable for intravascular administration owing to the solvents or surfactants used in the method of preparation. The methods available for the entrapment or encapsulation of hydrophilic compounds are few innumber anddelicate, since they often requirea step involving the transfer of the nanoparticles from an oily preparation medium to an aqueousmedium more suitable for pharmaceutical purposes.

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In the future, effortsshould be made tohave a better understanding concerning the mechanism of formulation of these colloidal systems at the molecular and supramolecular level. This could lead to new formulation process and could open new prospects in the area of drug delivery by means of nanoparticle systems.

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Couwreur et a/. terionicPolymerisationofCyanoacrylates,Makromol.Chem.,190:1613 (1989). E. F. Donnelly, Ionic and Zwitterionc Polymerisation of n-Alkyl 2-Cyanoacrylates, Polymer Let. Ed., 15:399 (1977). D. C. Pepper, Anionic and Zwitterionic Polymerisation of a-Cyanoacrylates, J. Polymer Sci.: Polymer Symp., 62:65 (1978). D. Aleony and H. Wittcoff,U.S. Patent 2,899,397,1959. W. Cooper, U.S. Patent 3,009,891, 1961. P. Judd, Brit. Patent 1,142,375, 1969. G. S. Banker, U.S. Patent 4,330,338, 1982. J. W. Van Der Hoff, U.S. Patent 867,031,1977. R. Gurny, N. A. Peppas, D. D. Hamngton, and G. S. Banker, Development of biodegradable and injectable latices for controlled release of potent drugs, Drug Dev. Ind. Pharm.,7:l (1981). H. J. Krause, A. Schwartz, and P. Rohdewald, Polylactic acid nanoparticles, a colloidaldrugdeliverysystemforlipophilicdrugs,Int.J.Pharm.,27:145 (1985). H. Ibrahim, C. Bindschaedler, E. Doelker, et al., Aqueous nanodispersions by a salting-out process,Int. J. Pharm., 87:239 (1992). E. Allemann, R. Gurny, and E. Doelker, Preparationof aqueous polymeric nanodispersions by a reversible salting-out process: Influence of process parameters on particle sue, Int. J. Pharm., 87:247 (1992). Rhone-PoulencRorer,Microsphtres,leurprocCd6 de prtparation et leur utilisation, Fr. Patent 2660556, 1990. T. Verrechia, P. Huve, D. Bazile, et al., Adsorption / desorption of human serum albumin at the surfaceof poly(1actic acid) nanoparticles prepared by a solvent evaporation process, J. Biomed. Mater. Res., 27:1019 (1993). P. A. Kramer, Albumin microspheres as vehicles for achieving specificity in drug delivery, J. Pharm. Sci., 63:1646 (1974). T. Yoshioka, M. Hashida, S. Muranishi, and H. Sezaki, Specific delivery of mitomycin C to the liver, spleen and lung: Nano- and microspherical camers of gelatin, Int.J. Pharm., 8:131 (1981). U.Schroeder and A. Stahl, Crystallized dextran nanospheres with entrapped antigen and their use as adjuvants, J. Immunol. Meth., 70:127 (1984). H. Fessi, J-P. Devissaguet, F. Puisieux, and C. Thies, ProcCdC de prCparation de systkmes collo'idaux dispersibles d'une substance sous forme de nanoparticules, Fr. Patent 8618446, 1986. R. K. Chang, J. C. Price, and C. Hsiao, Preparation and preliminary evaluation of Eudragit RL and RS pseudolatices for controlled drug release, Drug Dev. Ind. Pharm., 15:361(1989). R. Bodmeier, H. Chen,P. Qle, and P. Jarosz, Spontaneous formation of drug8: 161 (1991). containing acrylic nanoparticles, J. Microencaps., J. J. Marty, R, C. Oppenheim, andP.P. Speiser, Nanoparticles-a new colloidal drug delivery system, Pharm. Acta Helv., 53:17 (1978). S. Stainmesse, H. Fessi, J.-P. Devissaguet, and F. Puisieux,ProddB deprtpa-

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rationdesystbmescolloidauxdispersiblesd‘uneprotkinesousforme de nanoparticules, 1st Add. to Fr. Patent 8618446, 1988. 55. H. Fessi, J-P. Devissaguet, and F. Puisieux, Procede de preparation de systbmes colloidaux dispersibles d’une substance sous forme de nanocapsules, Fr. Patent 8618444,1986. 56. C. V. Sterling andL. E. Scriven, Interfacial turbulenc: hydrodynamic instability and Marangoni effect,A. I. Ch. E.J., 5514 (1959). 57. C. Washington, Evaluation of non-sink dialysis methods for the measurement of drug release from colloids: Effects of drug partition, Int. J. Pharm., 56:71 (1989). 58. N.Ammoury, M. Dubrasquet, H. Fessi, et al., Indomethacin-loaded poly(d,llactide) nanocapsules : protection from gastrointestinal ulcerations and antiinflammatory activity evaluation in rats, Clin. Mat., 13:121 (1993).

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Biodegradable Colloidal Drug Carrier Systems Based o n Solid Lipids Kirsten Westesen Friedrich-Schiller University Jena, Jena, Germany

Britta Siekmann Astra Arcus A B , Sodertalje, Sweden

I. Colloidal Carriers in Drug Delivery and the Use of Biodegradable Lipids A. Polymeric Carrier Systems B. Lipid-Based Carrier Systems 11. Lipid Suspensions as an Alternative Delivery System for the Administration of Poorly Water-Soluble Drugs A. Lipid Nanopellets for Persorption B. Fatty Acid Nanoparticles Prepared from Oil-in-Water Microemulsions C. Lipospheres-Lipid Particles Used for Sustained Drug Release D. Colloidal Lipid Suspensions Prepared by Precipitation in Emulsified Organic Solvents E. Colloidal Lipid Suspensions Prepared by MeltEmulsification

111. Physicochemical Characterization of Melt-Emulsified Colloidal Lipid Suspensions A. Investigations of the Physical State of Colloidal Lipid Dispersions Prepared from Crystalline Glycerides B. Investigations on the Crystallinity and the Recrystallization Process of Melt-Emulsified Glyceride Nanoparticles

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C . Mechanism of Gel Formation in PhospholipidTripalmitate Stabilized Suspensions D.Incorporation of Drugsinto Solid Lipid Nanoparticles IV. Prospects Summary Future and

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1. COLLOIDAL CARRIERSIN DRUG DELIVERY AND THE USE OF BIODEGRADABLE LIPIDS

Many drug substances are characterized by poor solubility in aqueous media and thus cause formulationproblems with regard to parenteral administration. Besides the use of cosolvents, drug complexation and solubilization in surfactant micelles, incorporation in colloidal carrier systems represents an alternative way to render poorly water-soluble drugs applicable by the parenteral and in particular the intravenous route. Futhermore, incorporation of drugs in particulate carriers provides a possibility to mais desired. In recent nipulate drug releaseif controlled or sustained release years, colloidal carrier systems have also attractedgrowing interest concerning drug delivery to site-specific targets, especially incancer chemotherapy [1,2]. The objectiveof drug targetingis an increase of drug concentration at the desired site of action with a simultaneous decreaseat nontarget sites, with the rationale being the enhancement of therapeutic efficacy with a simultaneous reductionof unwanted side effects. During the last decades, several approaches have beeninvestigated to develop submicron-sized drug-delivery systems. Based on the carrier material, the conventional vehicles used as drug carriers can generally be divided into two groups, polymeric andlipidic systems. A.PolymericCarrierSystems

Polymeric nanoparticles are (amorphous) solid colloidal particles consisting of nonbiodegradable syntheticpolymers or of biodegradable macromolecular materialsof synthetic, semisynthetic,or naturalorigin. Theyusually exhibit a longshelf life and a good stability on storage. Drug release from polymeric nanoparticles is in many cases found to be biphasic with an initial burst phase followed by a sustained release [3]. The burst effect can be related to drug adsorbed to the camer surface. The methods for the prep ration of polymeric nanoparticles such as emulsion polymerization and solvent evaporation techniques often involve the use of toxicologically harmfulexcipients and additives,forexample,organicsolvents,cancerogenic monomers, and reactive crosslinking agents, the complete re-

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moval of which from the productis hardly possible from atechnical point of view [4]. Moreover, the carrier material itself can represent a potential toxicological risk. Apart from polymer accumulation on repeated administration owing to slow biodegradation, toxic metabolites may be formed during the biotransformation of polymericcamers; for example, formaldehyde as a metaboliteof polycyanoacrylates [5]. B.Lipid-Based

Carrier Systems

In order to avoid potential toxicological problems associated with polymeric nanoparticles, a great deal of interest focuses on lipid-based carrier systems. Lipidic camers comprise, among others, liposomes, lipoproteins, andlipid oil-in-water emulsions. These vehicles are composed of physiological lipids such as phospholipids, cholesterol, cholesterolesters,and triglycerides, and owing to thebiological origin of the carrier material, the toxicological risk is much lower than thatof polymeric particles. There are,however, a number of drawbacks inherent in conventional lipid carriers which are basically related to physicochemical instabilities. Liposomes, for example, tend to fuse, therebyreleasing drug from the vesicles. In particular,small unilamellar vesicles (SUV) are in a thermodynamicallyunfavorable state owing to the high curvature of the phospholipid bilayer [6]. The problem is even more pronounced when drugs are incorporatedin the liquid crystalline phospholipid bilayer. The storage stability of liposomesis therefore limited. Owing to lipid exchange with lipoproteins, liposomes are often also unstable in the vascular system [7]. Moreover, the reproducibility of liposome manufacture in terms of vesicle sizeand properties is limited and thus complicates largescale production. These technological problems have so far impeded the widespread clinical useof liposome formulations. Lipoproteins are thephysiological carriers of water-insoluble lipids in the vascular system. Since lipoproteins exhibit a considerable circulation time in blood and are not taken up by the reticuloendothelial system (RES), they have been considered as drug-delivery systems [8,9]. Lowdensity lipoproteins (LDL) have attracted particular interest in this respect, because they areableto leave the vascular system by specific receptor-mediated uptake into cells. Tumor cells exhibit an increased uptake of LDL owing to their high cholesterol need, which forms thebasis of the LDL drug-targeting concept in cancer chemotherapy [lo]. As with liposomes, the shelf life of drug-loaded LDL is, however, only very limited on account of physicochemical instabilities. Furthermore, isolationof LDL from human blood is a time-consuming procedure, and large-scale production is limited by the access to human blood. The isolation procedure also bears the potential infectious riskwith respect to hepatitis and human

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immunodeficiency virus (HIV). A circumvention of these problems might be the preparation of artificial LDL [ll]. Production of artificial LDL containing the apolipoprotein compound which mediates receptorrecognition has, however, not yet been reported so far. Submicron-sized vegetable oil-in-water ( O N ) emulsions have been used as a calorie source in parenteral nutrition for decades [12,13]. These systems are manufactured in large scale and display an acceptable longterm stability. Lipidemulsions have therefore been extensively investigated as drug-carrier systems [14,15]. However, despite these efforts and the available production technology, there have been remarkably few drugcontaining lipid emulsions on the market until now, whichpoints to formulation problems caused by the susceptibility of the carrier toward incorporation of drug. This can be attributed to drug crystallization within the oil droplets and perturbations of the stabilizing emulsifierfilm by drug crystals or diffusing drug molecules which have a high mobility in the liquid oil phase. These perturbations may induce instabilities of either mechanical (reduction of film elasticity, film ruption) or electrochemical nature (influence on the zetapotential) causing coalescence, particle growth, and drug leakage. Drug leakage is the unwanted loss of drug from the carrier on storage. The high mobility of incorporated drug molecules also causes a fast release of drug fromthe carrier in biological fluids, so that it is hardly possible to achieve sustained release by an emulsionformulation [161.

II.

LIPID SUSPENSIONS AS AN ALTERNATIVE DELIVERY SYSTEM FOR THE ADMINISTRATIONOF POORLY WATER-SOLUBLE DRUGS As outlined above, most drawbacks associated with conventional lipid drug-carrier systems can be attributed to the liquid or liquid crystalline state of the dispersed lipid phase. It was therefore obviousto combine the superiorities of colloidal lipid carriers, such as the biodegradability and biocompatibility of the carrier material as well as the easeof manufacture, with the advantages of the solid physical state of polymeric nanoparticles with respect to size stability, drug leakage, and sustained drug release. One approach was the idea to prepare aqueous suspensions of lipid nanoparticles by using solidlipids and thus circumventing the toxicological problems associated with polymer particles. Possible advantages of the solid physical state of the dispersed lipid phase are summarized in Table 1. By definition, solid particles cannot coalesce, and they may therefore exhibit a better physical stability than liquid or liquid crystalline carriers. Since the mobility of incorporated drug molecules is drastically reduced in a solid phase, drug leakage from the camer and drugprotrusion into theemulsi-

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Table 1. Possible Advantagesof the Solid Physical State of Lipid Carrier Systems 1. Avoidance of coalescence enhanced physical stability 2. Reduced mobility of incorporated drug molecules reduction of drug leakage circumvention of instabilities due to interactions betweendiffusing drug mole-

cules and emulsifierfilm sustained drug release 3. Static interface solidlliquid facilitated surface modification

fier film would be counteracted. In addition, drug release from the carrier is not a diffusion-controlled process but is matrix-dependent provided the drug is distributed in the matrix and the matrix does not melt at body temperature. Thedegradation rate of the carrier lipids is assumed to determine drugrelease, which inthat case could be controlled to a certain extent by the choice of matrix constituents. Moreover, the presence of a static interface might facilitate a surface modification of the carrierparticles after solidification of the lipid matrix; for example,by subsequent adsorption of nonionic surfactants. The latter might be of relevance to reduce carrier uptake by the reticuloendothelial system, which is known to be related to surface properties. Surface modification is therefore one approachto drug targeting using colloidalcarriers [171. It is known to the authors that first attempts to develop lipid-based colloidal suspensions already date back to decadesagowhen the first parenteral lipid emulsions becamecommercially available. The early concepts considered to transfer the production principles of submicron-sized phospholipid stabilized O W emulsions tothepreparation of colloidal lipid suspensions by substituting the dispersed liquid oil phase by solid triglycerides. However, the attempts to formulate long-chain triglycerides into phospholipid-stabilized aqueous suspensions failed because of stability problemssuchas the formation of semisolid gels, whichwas also observed by the authors [l81 and will be discussed later (cf. Section 1II.C). Until the beginning of this decade, the development of colloidal lipid suspensions which exhibit a satisfactory long-term stability had not been reported in the pharmaceutical literature. Despite obvious stability problems, a patent application on the production of lipid nanopellets for persorption (cf. Section 1I.A) by melt-emulsification was filed by Speiser in 1985 [19]. Product stability is, however, not taken into consideration in the application.

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In contrast to submicron-sized lipid suspensions, the production of solid lipid matrices in the micrometersize range has already been reported in the late 1950s and the beginning of the 1960s [20,21]. Different preparation techniques have been described, such as milling of drug-containing lipid phase [20]; melt dispersion, for example, by stirring an emulsifier-free wax emulsion in water [21,22]; solvent methods in which the wax is dissolved in an organic solvent that is evaporated [22]; spray-drying of lipids dissolved in a volatile solvent [23]; and spray-congealing of the lipid melts [20,23]. Lipid microspheres are predominantly used as sustained-release oral solid-dosage formulations. Stability problems with respect to sedimentation or gel formation have not been described. The microparticles are generally separatedfromtheaqueous dispersion medium.These microparticulate systems are, however, not suitable as intravenous drug carriers. Moreover, the long-term stability of lipid microparticles has been rarely investigated. Most investigators focus their interest on product parameters and drug release of freshly prepared systems only.The few studies published on polymorphic behavior of spray-dried and spray-congealed lipid micropellets reveal that the micropellets are obtained in an unstable polymorphic formwhich transforms gradually into the stable modification on storage [24]. This polymorphic transition is accompanied by changes in the surface structure of the micropellets. A possible influence of particle aging on drugrelease is commonlynot discussed inthe literature, although physicochemical alterations during storage such as polymorphic transitions of the lipids, changes in particle size and shape, andincreased crystallinity might be of relevance. In the beginninngof the 1990s, a number of research groups focused their activities on the development of solid lipid-based colloidal carrier systems, reflecting the pressing importance of alternative parenteral carriers. The different approaches are briefly summarized in the following sections. It has to be mentioned that the prevailing number of these studies are still at an early stage of development, and up to now most systems under investigation are only poorly characterized from a physicochemical point of view. In particular, stability data of these lipid carriers are lacking, and some investigators seem to disregard or underestimate this important aspect.

A.

Lipid Nanopellets for Persorption

In a patent application, Speiser [l91 discloses lipid nanopellets as an oral drug-delivery system with a particle size small enough for persorption. Persorption is definedas the transport of intact particles through the intestinal cell layer into thevascular or lymphatic system under avoidanceof the

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first-pass effect in the liver. Persorbed lipid nanopellets are believed to be concentrated in adipose tissues to form a sustained-release depot of incorporated drugs [19]. This theoretical mechanism is, however, not substantiated by experimental data. Aqueous suspensions of lipid nanopellets are described to exhibit a particle size of predominantly 80-800 nm. The carriers consist of waterinsoluble solid lipids such as long-chain glycerides, fatty acids, and fatty alcohols, and surface-active agents, preferably physiological emulsifiers. Colloidal suspensions are obtained by melting the lipids and adding them to the warm aqueous phase. The lipid melt is dispersed in the aqueous phase by high-speed stirring and sonication. Drugs can be incorporated into the carrier matrix by dissolution or dispersion in the melt or by addition of a solution of the drug in a volatile organic solvent to themelt and subsequent solvent evaporation. Data regarding the storage stability of these lipid nanopellets are not specified in the patent application. Based on Speiser’s patent application [191, Kecht-Wyrsch investigated the preparation of aqueous suspensions of lipid nanopellets [25]. However, Kecht-Wyrsch succeeded only in preparing lipid suspensions with a lipid concentration as low as 0.2 to 1.0% (w/w). These suspensions exhibited a documented storage stability of 3 months, whereas higher concentrated suspensions displayed a considerable particle growth and tended to form semisolid gels [25]. This investigator reports furthermore that preparation of lipid nanopellets by high-pressure homogenization was not feasible and yielded particles in the micrometer size range only [25]. Comprehensive physicochemical studies on lipid nanopellet suspensions of the type described by Speiser [l91 are yet outstanding. However, the limited set of published data demonstratedalready some of the physicochemical difficulties and problems associated with the suspension state compared with the emulsion state. B. Fatty Acid Nanoparticles Prepared from Oil-ln-Water Microemulsions

Gasco et al. have developed a method to prepare solid lipospheres from O/W microemulsions [26-281. Microemulsions were prepared from mixtures of melted fatty acids which may contain a drug, a surfactant, and possibly a cosurfactant in warm water by stirring. Typical formulations comprise stearic acid, m e e n 20, taurodeoxycholate, and butanol [27,28]. The warm microemulsion is diluted and cooled down by dispersion in a cold aqueous phase. In a subsequent step, the liposphere dispersion is washed by diaultrafiltration. Lipophilic drugs can be incorporated in the melted stearic acid prior to formation of the microemulsion. The method

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typically yields lipid particles with an average size of around 200-300 nm depending on production variables such as mixing rate, temperature protocol, and pH of the aqueous medium[28]. In a study of critical formulation parameters for the formation of stearic acid lipospheres from diluted microemulsions using butanol as a cosurfactant, it was found that the growth kinetics typically displayed a biphasic pattern with an initial, rapid size increase up to 10h after dispersion followed by a slower equilibration phase [D]. Incorporation of decanoic acid in the lipid phase was observed to impede submicron particle formation at concentrations about 25% and to disrupt lipid phase equilibrium resulting in continuous and accelerated particle growth. Below around pH4, it was found that minimal variation of acidity could strongly modify growthkinetics. Moreover, differences in butanol amount strongly affected both initial particle growth and equilibrium stability, and at optimum butanol concentration, maximum liposphere stability was observed. The investigators hypothized that the observedeffects might be attributed to interfacial interactions in the microemulsion or to differentcrystallization pathways of stearic acid [29]. Studies on the crystallization of washed and dried stearic acid lipospheres by x-ray powder diffraction and differential scanning calorimetry indicated that polymorph C is always present in the lipospheres and that additional crystallization of polymorph B from the microemulsion systemis favored in the presence of small amounts of butanol, whereas this polymorph is not present in mixtures of stearic acid, butanol, and Tween 20, which had undergone the same thermal cycle and had the same composition as the dried lipospheres [30]. At present, it is not clear whether polymorph B has its own liposphere population separate from that of polymorph C or whether the two polymorphs cocrystallize in the same liposphere [30]. It has to beconsidered that thecrystallization studies were conducted ondried lipospheres which had been separated fromthe dispersion medium. The crystallographic state of lipospheres in aqueous dispersion was not investigated but might be different from the findings on dried lipospheres, as crystallization is generally affected by the crystallization medium. Moreover, the composition of lipospheres in dispersion is different from dried lipospheres, since butanol is evaporated from the lipospheres during the drying procedure as indicated by gas chromatographic results on dried lipospheres [30]. In a modification of the microemulsion method developed by Gasco et al. [27,28], ?keen 20 and the organic solvent were substituted by purified egg lecithin [31]. The obtained average diameters of drug-loaded lipospheres ranged between 70 and 180nm. The shape of the stearic acid particles was described to be spherical [31]. The stability on storage of

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stearic acid liposphere suspensions with respect to properties which might be relevant for drug distribution and release such as particle size, particle shape, and degree of crystallinity was not discussed by the investigators. The fact that the lipospheres were freeze-dried for storage [31] points, however, to stability problems of these lipid suspensions. Results from our own laboratory indicate that stearic acid liposphere suspensions prepared according to Gasco et al. [31] display a significant particle growth within a couple of hours or days after preparation necessitating their lyophilization for pharmaceuticalapplications [32]. Timolol-containing lipospheres that comprisedpalmitic and decanoic acids as carrier material were suggested as an ocular delivery system by Gasco et al. [33]. Timolol was mainly present as ion pairs in the lipospheres in order toincrease its lipophilicity. Differences in the incorporation capacity were attributed to thedifferent lipophilicity of the ion pairs [33]. It has not yet been investigated how far themicroemulsion method can betransferred to the use of carrier lipids other thanfatty acids. Although the patent by Gasco [26] claims that triglycerides and fatty alcohols also can be employed as lipid components, no evidence thereof is provided in the patent examples, which represent exclusively fatty acid lipospheres. Moreover, long-term stability neither of the aqueous dispersions nor of the dried or freeze-dried storage formulations have been published yet. This is, however, of importance, since even freeze-dried suspensions might undergo particle fusion on storage (cf. Section 111). In addition, studies on possible changes during freeze-drying or storage with respect to particle shape, degree of crystallinity, or polymorphic transitions of the stearicacid which might influence drug distribution within the carrier and drugrelease have not yet been reported. C. Lipospheres-Lipid Particles Used for Sustained Drug Release

An injectable lipid-delivery system for controlled release, referred to as Lipospheres, has been reported by Domb and Maniar[34]. The liposphere system is described to represent a particulate dispersion of solid spherical particles consisting of a hydrophobic core compoundsuch as triglycerides or fatty acid derivates surrounded by a layer of phospholipids. Preparation of Lipospheres can beaccomplished either by a melt or by a solvent technique in which the core compound is melted or dissolved in an organic solvent and is agitated in an aqueous medium at a temperature above the melting point of the core compound until a homogeneous dispersion is obtained. Lipospheres are formed when the dispersion is rapidly cooled down or thesolvent is evaporated, respectively [34]. Both production meth-

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ods are basically known from the preparation of wax microspheres [e.g., see ref. 221. The storage stability of Lipospheres seemsto be very limited. Domb and Maniar [34] designate a 7-day storage stability of submicronsized tristearin lipospheres (based on particle size analysis) as ‘‘exceptionally stable.” Lipospheres in the micrometer size range Seem to be more stable. The particle sizes and distributions were found to besimilar to the original formulation after 30 days [34],and for oneformulation, Domb and Maniar [34] observed no signs of aggregation for 10 months stored at 4°C. Results from our own laboratory indicate, however, that such liposphere formulations tend to sedimentation (unpublished data). Considering the physicochemical properties of the matrix material used for theexamples in the Lipospheres patent application [34], the spherical shape of the particles, in particular those in the nanometer size range, might not be preserved on storage. Additional changes in particle properties such as the modification of the lipids on storage as has been reported for sprayed micropellets [24] or the degree of crystallinity of the matrix have not yet been discussed for Lipospheres by the investigators despite their possible relevance for drugrelease. In a strict sense, liposphere preparations do not represent colloidal carrier systems owing to their micron size. The average particle size of a and Lipotypical liposphere formulation is between 1 and 20pm[35], spheres are thus suitable for intramuscular or subcutaneous injection but not for intravenous administration. Since there are, however,a number of in vivo studies performed using these solid lipid-based drug carriers, the results will be briefly reported below to illustrate the potential of the use of solid lipids in sustained drug release and drugdelivery. It has to beconsidered, however, that theproperties of the colloidal solid lipid particles might be significantly different from those of microparticles. It is therefore not unambiguouslypossible to conclude drug release behavior of colloidal lipid carriers from results obtained on microparticulate systems. The extended release of drugs from microparticulate liposphere formulations has been demonstrated by in vitro release studies. Drug release was observed over several days, although there is an initial burst of drug related to the first few hours [34], indicating that parts of the drug are not incorporated in the particle core but areassociated with the carriersurface; for example, in the lecithin layer, from where the drug is fast released. The in vitro data on sustained drug release could be substantiated by in vivo studies, and sustained delivery of local anesthetics, antiinflammatory agents, insect repellents, and antibiotics was observed in different animal models by monitoring the therapeutic effect over time after liposphere administration [34,35]. The use of liposphere preparations as vaccines in immunization by incorporating antigens in the lipid carrier has also been

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demonstrated. Immunization with malaria antigen encapsulated in Lipospheres was shown to increase the IgG antibody titer in rabbits [34]. No significant differences were observed whether the antigen was incorporated in the Lipospheres from an aqueous buffer solution or added as a lyophilized powder to the lipid components in the first step of liposphere preparation [34]. Comparison of the animal immune responseusing Lipospheres or liposomes as carriers of the same antigen revealed that the antibody titers in animals injected with Lipospheres weremuch higher than in those injected with liposomes, whereas antigen alone or in the presence of adjuvants produced virtually no titer [34]. The drug loading into theliposphere core is highly dependent on the method of preparation. Incontrast to melt-dispersed Lipospheres, formulations prepared by the solvent method showed that a significant amount of the drugis trapped in the phospholipid layer [36]. These results might be of relevance to drugrelease from the particles. D. Colloidal Lipid Suspensions Prepared by Precipitation in Emulsified Organic Solvents

Sjostrom andBergenstdhl have presented a method to prepare submicronsized particles of cholesteryl acetate by precipitation in lecithin-stabilized O W emulsions [37]. Cholesteryl acetate and lecithin were dissolved in an organic solvent, and the organic solution was emulsified in an aqueous phase containing a cosurfactant by high-pressure homogenization to yield a submicron-sized OiW emulsion. On removal of the organic solvent by evaporation under reducedpressure, cholesteryl acetate nanoparticles precipitated in the emulsion droplets. Inspired by our laboratory, Sjostrom and BergenstHhl employed, among others, a blend of phospholipids and bile salts as emulsifiers. It turned out that only with a blend of phosphatidylcholine and sodium glycocholate particles as small as 25 nm could be obtained. The storage stability as monitored over a period of 30 days by repeated size measurements depended onthe lecithidbile salt ratio. With increasing ratio, the stability was found to be reduced. It was not possible to prepare stable emulsions of the cholesteryl acetate containing organic solvent in water using pure phosphatidylcholine without the addition of cosurfactant. Transmission electron microscopy (TEM) of freeze-fractured replicas of a selection of cholesteryl acetate nanoparticles freshly prepared by precipitation in solvent emulsions revealed that the majority of particles had a smooth surface and were spherical [38]. However, larger particles, in particular those above 80-100 nmin size, displayed a nonspherical particle shape and a layered structure already in fresh preparations (Fig.

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Fig. 1. TEM pictureoffreeze-fracturedreplica of cholesteryl acetate particles prepared by precipitation in a phospholipidlsodium glycocholate-stabilized cyclohexane-in-wateremulsion. The bar corresponds to 128 nm. 1). In a subsequent investigation, the morphology of cholesteryl acetate particles was identified as disk-shaped platelets of uniform thickness by cryotransmission electron microscopy [39]. The conclusions drawn from TEM observations of freeze-fractured replicas and from cryopreparations are controvertial, and they might result from particle aging, polymorphism of particle shape, limited number of sample preparations, or misinterpretation of observations. For particles as small as 10 nm, it is, for example, difficult to distinguish between a disk shape and a platelet shape of a particle. TEM studies on freeze-fractured replicas of aged cholesteryl acetate particles are still outstanding. Although cholesteryl acetate particles were designed as a model of drug nanoparticles, itis possible to use these kinds of colloidal dispersions as drug-camersystems as well as by incorporating lipophilic drugs intothe cholesteryl acetate coreor by substituting cholesteryl acetate on account its artherogenic potentialby other camermaterials such as triglycerides. In an attempt to transfer the precipitation method described by Sjostrom and Bergenstihl [37] to the manufacture of tripalmitate nanoparticles, we succeeded in preparing aqueous tripalmitate suspensions with mean particle sizes ranging from 25 to 120 nm depending on the lecithin-cosurfactant blend, with sodium glycocholate beingthe most effective coemulsifier [40].

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Physicochemical characterization of these tripalmitate suspensions by TEM revealed that, in contrast to cholesteryl acetate, tripalmitate precipitates as anisometrical nanoparticles of predominantly plateletlike shape with a layered structure (Fig. 2). The particles resemble those prepared by meltemulsification (cf. Section III.C), although they are smaller. According to investigations byx-ray diffraction and differential scanning calorimetry (DSc), the recrystallization tendency of precipitated tripalmitate nanoparticles is similar tothat of melt-emulsified suspensions (cf. Section 1II.B). These results indicate that the physicochemical properties of lipid nanoparticles seem to depend predominantly on the matrix material with less importance being placedon the production method. The storagestability of precipitated tripalmitate nanoparticles is, however, poorer than that of melt-emulsified systems. Precipitated tripalmitate suspensions tend to grow in particle sizewithin several weeks or a few months.Residual amounts of organic solvent could be detected by 'H nuclear magnetic responance spectroscopy and DSc. Apart fromtoxicological problems arising from residues of organic solvents, a clear limitation of the precipitation method is the low lipidconcentration in the suspension after evaporationof the solvent. In the case of tripalmitate precipitated in chloroform emulsions, the maximum lipidconcentration achievable was no more than2.5% (w/w)owing to the limitedsolubility of the triglyceride in the organic solvent. Much higher triglyceride concentrations can be achieved by the melt-emulsification method described below (cf. Section 1I.E). E. ColloidalLipidSuspensionsPrepared by Melt-Emulsification Inthe beginning of the 199Os, two patent applications on solidlipid nanoparticles emerged from the laboratories of Muller [41] and Westesen [42]. Both patent applications are based on a melt-homogenization technique favoring the use of high-pressure homogenizers. Whereasthe patent application by Muller and Lucks[41] does not give any precise details with respect to thestability and the physicochemical properties of the disclosed colloidal lipid particles, that by Westesen and Siekmann [42] provides a detailed description of the novel systems, in particular regarding the stabilization and the long-term stability. Siekmann and Westesen were the first to report on melt-emulsified aqueous lipid suspensions with a storage stability of more than 12 months [H]. The reported melt-homogenization method circumvents the use of organic solvents. Solidlipids, for example, long-chain glycerides, are melted, and the melt is dispersed in an aqueous phase by sonication or preferably by high-pressure homogenization with the aid of emulsifying

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Fig. 2. TEM pictures of freeze-fractured replicaof tripalmitate particles prepared by precipitation in a phospholipidsodium glycocholate-stabilized chloroform-inwater emulsion. (a) The bar corresponds to 150 nm. (b)The bar corresponds to 185 nm.

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agents. Emulsification by high-pressure homogenization is more effective and yields more uniformlysized particles than sonication. Thelatter method also bears the disadvantage of considerable metal contamination from the sonication probe. Metal contamination can be avoidedusing highpressure homogenizers with ceramic valves. Moreover, high-pressure homogenization can beused for virtually any batch size from small- to largescale production. The mean particle size of melt-emulsified lipidsuspensions is typically in the range between 50 and 300 nm determined by photon correlation spectroscopy (PCS), and it depends on the homogenization parameters as well as on the compositionof the suspensions; for example, on the matrix constituents, the volume fraction of the dispersed phase, and the type and amount of emulsifying agents. It was observed that the mean particle size of lipid nanoparticles increased with increasing melting point of the matrix constituent, indicating an effect of the melt viscosity [NI. At a constant matridemulsifier ratio, the mean particle size increases with increasing concentration of the dispersed phase. With increasing energy input, for example, by increased homogenizationpressure or by prolonged homogenization times, the meanparticle size isgenerally decreasing until it reaches a minimum and then levelsoff. In some cases, a reverse effect, that is, increasing particle size, can be observed with too high-energy input, a phenomenon referred to as overemulsification. In summary, the effects of composition and homogenization parametersare basically similar to those found for oil-in-water emulsions [43,44], and they thus reflect the emulsionlike state of the dispersed molten lipids during emulsification. The effect of the type and amount of emulsifying agents is very pronounced with respect to both the particle size distribution and the storage stability of melt-emulsified solid lipidnanoparticles [45]. Glyceride dispersions stabilized by phospholipids only in concentrations analogously to the stabilization of commercial parenteral emulsions, that is 12 W% lecithin related to the fatphase, display a broad and bimodalsize distribution with relatively high mean particle sizes. Moreover, these systems were observed to exhibit a consistency change during storage and form semisolid, ointmentlike gels. Increasing the amountof lecithin to 60 W%(related to the fat phase) results in a decrease of the meanparticle size and in a more homogeneous size distribution, but it cannot effectively prevent gel formation. The prevention of gel formation is, however, possible by the addition of highly mobile coemulsifying agents to theaqueous phaseof phospholipid-stabilized glyceride dispersions. These coemulsifiers also decrease the mean particle size of the lipid nanoparticles, and lecithin-coemulsifier-stabilizedglyceride dispersions generally display monomodal, homogeneous size distributions with mean sizes well below those of commercial lipid emulsions (Fig. 3).

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PL (12O/o)/SGC PL (60%) PL (6O%)ISGC lntralipid 100/0

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? C

xi* a2

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0 0

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300 400 Particle size (nm)

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Fig. 3. PCS particle size distribution(bynumber) of differently composed glyceride nanoparticledispersionsusing the suppository base Witepsol W35 as matrix constituent, compared to a commercial lipid emulsion for parenteral nutrition (Intralipid 10%).PL, phospholipids, SGC, sodium glycocholate.

Both ionic and nonionic surfactants such asthe anionic bile salt glycocholate [l81 or thenonionic polymer tyloxapol[46] proved to be efficient coemulsifiers with regard to particlesize distribution andlong-term storage stability (>l8 months), indicating that electrostatic or steric barriers can counteract gel formation. The use of steric stabilizers such as tyloxapol or thenonionic block copolymers of polyoxyethylene and polyoxypropylene, that is, poloxamers, require relatively high amountsof surfactants for effective stabilization, which can be attributed to theirspecific stabilization mechanism. Steric stabilization requires that thestabilizer molecules form a sufficiently thick layer on the particle surface and adsorb to the interfacein a specific steric arrangementwith the ethyleneoxide chains protruding into the continuous phase so as to build up an effective steric barrier toward particle attraction. The numberof these high molecular weight molecules required for effective stabilization is therefore relatively high as expressed on aweight basis. The efficiency of lecithin-nonionic polymer blends was shown to depend on the phospholipid/surfactant ratio in the case of tyloxapol [46]. Nonionic polymers can be used as stabilizers by themselves without being combined with lecithin, but it was observed that glyceride dispersions stabilized by poloxamers only tend to form gels when beingsubjected to shear forces; for example, by pressing the dispersion through the needle of a syringe. The same phenomenon was observed when the dispersions were

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stabilized by highly negativelycharged phospholipids [45]. Steric or electrostatic barriers can thus not be theonly prerequisite for preventing gel formation. A modelof the mechanism of gelformation andits prevention based on further experimental datawhich suggestthat the phenomenonis related to the crystallization of the dispersed phase is discussedin Section 1II.C. After publication of the first paper on the preparation, stability, and physicochemical characterization of melt-emulsified colloidalglyceride dispersions by Siekmann andWestesen [M], Muller et al. disclosed the preparation of solid lipidnanoparticles with particle sizes between 200 and 400 nm by high-pressure homogenization of a preemulsified lipidmelt in a conference abstract. However, no information concerning the long-term stability and physicochemical properties of these systems were given [47]. In a recent publication, optimization of homogenization parameters with the aim to obtain formulations suitable for injection according to the United States Pharmacopeia (USP) XXII monograph entitled “Particulate Matter” was reported [48]. These investigators also came to the conclusion that highpressure homogenization is superior to sonication, and solid lipid nanoparticle dispersions produced under optimized homogenizationconditions were regarded as suitable for intravenous injection, since their content of particles >5 pm is similaror below that of commercial parenteralfat emulsions [48]. The study was conducted on melt-homogenized dispersions of trilaurate (Dynasan 112) stabilized by either soya lecithin or poloxamer using an emulsifier concentration of 12 or50 W% related to the fatphase. The number of large particles depended on homogenization parameters and was considerably affected by the type and amount of emulsifier. Qpical stability problems described for dispersions stabilized by phospholipid mixtures rich inphosphatidylcholine such as gel formation werenot reported by the investigators. It has to be considered, however, that melt-emulsified trilaurate displays a pronounced supercooling effect, as was found by thermoanalytical and x-ray investigations from our laboratory[49]. We observed that trilauratenanoparticle dispersions stored for more than3 months did not display any thermal transition in differential scanning calorimetry (Fig. 4), indicating that the melt-dispersed lipid does not recrystallize but remains in thestate of a supercooled melt. Recrystallization of meltemulsified trilaurate nanoparticles requires cooling to below 0 “C. It can therefore be argued that trilaurate nanoparticles are liquid at room temperature and thusrepresent O W emulsionsaccording to the International Union of Pure and Applied Chemistry (IUPAC) definition [50] instead of solid lipid particles as claimed by Muller et al. [48]. Since stability problems of lipid suspensions seem to be predominantly related to the crystallization of the dispersed phase (cf.Section III.C), gel formation of supercooled trilaurate dispersions cannotbeexpectedand may explain the good stability of

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325 330 TEMPERATURE ( K 1

Fig. 4. DSC thermogram of (a) a 10% trilaurate (Dynasan 112) dispersion stored for 3 months at room temperature and (b) the thermally untreated raw material Dynasan 112. Heating rate: 10 Wmin. phospholipid-stabilized trilaurate dispersions. It thus has to be considered that in a strict sense such trilaurate nanoparticles do not represent solid lipid nanoparticles. In a conference contribution, an 8-monthlong-term stability study on melt-homogenized solid lipid nanoparticles was reported by the group of Muller [51]. The composition of the investigated systems was, however, not described, impeding the reader’s evaluation of the physical state of the colloidal lipid particles. The investigators confirmed our previous results that the use of a combination of surfactants (a complex of emulsifiers) proved to be optimal for long-term storage [Sl]. They also reported that certain solid lipid nanoparticle dispersions may acquire a gel-like consistency during storage, resulting in solidification of the systems [52]. It was found thatespecially hightemperatures andlight exposure causedsolidification within a few days. In conference contributions, spray drying [53] and lyophilization [54] were suggested to generatea dry storage formulation of

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solid lipidnanoparticles to circumvent storage problems. However,data on the stability of the lyophilizates on long-term storage with respect to initial particle size were not reported. Results from our own laboratory indicate that lyophilization is not necessarily a suitable technique for improvement of long-term stability, since it turned out that the lyophilized powders of melt-homogenized triglyceride nanoparticles displayed an impaired redispersibility and exhibited a pronounced particle growth after storage for 12 months, which can tentatively be attributed to sintering processes during storage [S]. Surface modification of colloidal drug carrier systems represents a promising approach to reduce carrier uptake by the reticuloendothelial system (RES). In order todivert colloidal particles away from theRES, the surface of the particles should be hydrophilic and noncharged [17]. In preliminary experiments, we established that melt-emulsified glyceride nanoparticles can be surface modified by adsorption of nonionic block copolymers [45]. Lecithin-stabilized tripalmitate nanoparticles were incubated directly after preparation with poloxamer 407 and poloxamine 908 solutions. Polymer adsorption wasconfirmedby measurements of the zetapotential employing laser Doppler anemometry (Table 2). The surface potential of tripalmitate nanoparticles is considerably reduced by the adsorption of the polymers as compared with the phospholipid-stabilized dispersion. In a conference abstract MaaBen et al. [56] described that thein vitro uptake of surface-modified solid lipid nanoparticles into humangranulocytes was significantly reduced compared with surface-modified polystyrene nanoparticles. The investigators concluded from the distinct reduction of phagocytic uptake in vitro that surface-modifiedsolid lipidnanoparticles possess the potential to prevent RES uptakein vivo.

Table 2. Zetapotential of Surface-Modified Solid Lipid Nanoparticles After Incubation (24 h) with Nonionic Block Copolymers Composition (% [w/w])

l”

PL (%)

Polymer

5 5 5

2 2 2

3% P4W 3% P 908

-

Zetapotential (mV) -29.6 -1.9 -2.9

Abbreviations: l”, tripalmitate (Dynasan 116); PL, phospholipids (Lipoid S 100); P 407, poloxamer 407 (Phonic F127); P 908, poloxamine 908 (Tetronic 908).

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111. PHYSICOCHEMICALCHARACTERIZATIONOF MELT-EMULSIFIED COLLOIDAL LIPID SUSPENSIONS

The idea behind the use of solid lipids as a carrier material is that the emulsified molten lipids resolidify on cooling, creating a solid matrix in which the mobility of incorporated drug is drastically reduced. The advantages of the solid state of the carrier have already been outlined in Table 1. Although solid lipid-based carrier systems are produced from crystalline raw materials, it has to be considered that recrystallization from the dispersed lipid melt might be different from that of the bulk phase. The high dispersity and the small particle size and the presenceof emulsifying agents and drugs as well as the high surface-to-volume ratio of melt-emulsified lipids may account for differences in the crystallization behavior, the degree of crystallinity, and the polymorphism of the nanoparticulate lipids compared with the bulk materials. Investigations by Skoda and van den Tempe1 [57] revealed that the crystallization temperature of a tristearate solution in paraffin oil which wasemulsified in anaqueousphase waslower thanthat of the bulk triglyceride. Crystallization in the dispersed state thus requires a higher degree of supercooling, which can onthe one hand be attributed to fact the that crystallization can only take place in small, discrete, and limited volumes. Whereas crystallization in the bulk can proceed quickly over the whole material once it has started in one location, crystallization in dispersed systems thus has to start individually in each droplet independently of the stateof crystallization inother droplets. Such a crystallization behavior can be easilyvisualized by polarizingmicroscopy for lipid-in-water dispersions with droplet sizes inthe micrometer size range (Fig. 5). On the other hand,it has to be considered that crystallization in the dispersed state may be affectedby emulsifying agents. It was shown before that thepresence of emulsifiers has an effect on the polymorphism and the crystal structure of bulk triglycerides [58]. Crystal nucleation in crudely emulsified triglyceride solutions was observed to depend onthe molecular structureof the employed emulsifying agents [57]. Owing to these effects,alterations of the crystallization behavior in the dispersed state can be expected and may lead to liquid, amorphous, or only partially crystallized systems.The physicochemical characterization of glyceride nanoparticles was therefore first of all focused on the question whether melt-emulsified glycerides recrystallize in the dispersed state. In addition to the studies on the crystallization behavior, it should be investigated why solid lipid nanoparticles stabilized by phospholipids only, that is, analogously to the preparation of submicron-sized lipid O N emulsions,

,

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Fig. 5. Polarized light microscopy picture of a crude dispersion of tripalmitate stabilized by phospholipids and sodium glycocholate (magnification: X 160): The dispersed lipidmelt forms spherical, isotropic droplets. The excess of phospholipids in the system forms large multilamellar vesicles which exhibit maltese cross-like structures in the polarized light microscope. Crystallizationstarts from the particle to surface and proceeds over the whole particle volume. Larger droplets tend recrystallize first.

tend to form semisolid gels and how gel formation can be prevented by certain stabilizers and coemulsifying agents (see above). A. Investigations on the Physical Stateof Colloidal Lipid Dispersions Preparedfrom Crystalline Glycerides

Long-chain triglycerides which are solid at room temperature in the bulk exhibit a complex polymorphic behavior and can crystallize in three basic polymorphic forms, termed alpha(a),beta prime (g'), and beta (g). Polymorphic transformationsare monotropic and takeplace from a to p' to p, with the latter being the thermodynamically stable polymorph [59]. For complex glyceride mixtures such as the hard fat suppository base Witepsol W35, an intermediatecrystal form (pi)between p'and p has been described

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[60,61]. The different polymorphic forms can be unambiguously distinguished by their x-ray spacings, and x-ray diffraction should thus provide a direct method to characterize the physical state of glyceride nanoparticles. The glyceride carrier systems under investigation represent, however, dilute dispersions with lipid concentrations typically of approximately 10 W%. Furthermore, it has to be taken into account that dispersed the glycerides are present as nanoparticles with sizes as small as 50 nm. Considering thatthe long spacings of the investigated glycerides typically range from approximately 3.5 to 5 nm, such small particles can consist of only 10-15 crystal layers. In view of the low glyceride concentration and the low number of repeating units in the nanoparticles, only broad low-intensity diffraction peaks characteristic of poorly ordered systems can be obtainedusing a conventional x-ray source, ashas been demonstrated by investigations using a Debye-Scherrer camera[62]. The necessarily long exposure times of several hours furthermorepreclude a systematic investigation, and time-resolved xray diffraction measurements during temperature scans are not possible. In contrast, synchrotron radiation x-ray diffraction provides an efficient wayof performing such measurements even in dilute dispersions of submicronsized glyceride particles [62], as is illustrated below. Synchrotron radiation small and wide-angle x-raydiffraction measurements were performed on melt-homogenized glyceridedispersions employing tripalmitate and hard fat (Witepsol W35) as carrier matrices and lecithin, bile salts, and nonionic polymers (poloxamer, tyloxapol) as emulsifiers. The dispersions had beenstored for several weeks prior to x-ray measurements. Systems stabilized by poloxamers tendedto form semisolid gels.Small-angle x-ray (SAX) diffraction revealed that all investigated carrier systems were crystalline at room temperature. The reflections in SAX diffraction patterns of the lipid dispersions were generally broader thanthose in the corresponding raw materials. This can be attributed to a particle size effect. The SAX spacings of tripalmitate dispersions correspond to thep modification of the raw material (Fig. 6). Lecithin-bile salt-stabilized hard fat dispersions (Fig. 7b) did not display the double peakcharacteristic of the pimodification of hard fat(Fig. 7a). Whereas for the semisolid hard fat gels this characteristic double peakcould be found(Fig. 7c),the broad SAX diffraction peaks of the hard fat dispersions did not permit a clear distinction between p'and pi forms. In contrast, the crystal modifications of all carrier systems could be unambiguously identified by wide-angle x-ray (WAX)diffraction and indicated that bothtripalmitate and hardfat nanoparticles are in the p modification at room temperature(Fig. 8a andsa). Hardfat-based carriers displayed particularly weak and broadmaxima at wide angles. Obviously,the crystal lattice of hard fatnanoparticles is highlyperturbed, since the complex com-

Biodegradable Colloidal Drug Carrier Systems

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Fig. 6. Synchrotron radiation small angle x-ray diffraction patterns (intensity Z vs scattering vector S): (a) bulk tripalmitate (raw material) and (b) 10 W% colloidal dispersion of tripalmitate stabilized by 1.2 W% soya lecithin and 0.4 W% sodium glycocholate (mean PCS particle size: 167 nm). The intensities of raw material and dispersion are noton the same scale.

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Fig. 7. Synchrotron radiation small angle x-ray diffraction patterns (intensity Z vs scattering vectorS): (a) bulk hard fat (raw material); (b) 10 W% colloidal dispersion 0.4 W% sodium glycocholate of hard fat stabilized by 1.2 W% soya lecithin and (mean PCS particle size: 92 nm); and (c) l 0 W% colloidal dispersion of hard fat stabilized by 1.8 W% Pluronic F-68 (semisolid gel). The intensities of raw material and dispersionsare not on the same scale.

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position of hard fatprecludes the formationof a true lattice. Moreover, the accumulation of lattice imperfections may be attributed to the small crystallite size in the dispersions. WAX diffraction at body temperature indicated that tripalmitate nanoparticles are still crystalline (Fig. 8b), whereas hard fatcarriers do not give any diffraction peaks (Fig. 9b), suggesting that the carrier material is melted at body temperature. The different physical state of the carriers at body temperature may influence their biopharmaceutical properties. The liquid hard fat nanoparticles give rise to fast drug release, whereas tripalmitate carriers may be suitable as sustained-release delivery systems. Being crystalline at room temperature, both carriersystems may provide an enhancedin vitro stability and a decreased risk of drug leakage onstorage as well as directly on administration owing to adsorption of drug to thewall material of syringes, containers with infusion media, or tubes. Preliminary investigations on the recrystallization behavior of melthomogenized glyceride dispersions were performedby time-resolved x-ray diffraction studies during temperature scans [62]. Hard fat nanoparticle dispersions were heated above the melting point of the carriermatrix, and the molten camer was cooled down stepwisemonitoring the WAX diffraction pattern. The raw material was exposed to a similar temperature program. The measurements indicated that recrystallization of the hard fat nanoparticles is different from that of the bulk material. Whereas the dispersed glycerides recrystallizein the a form, thebulk lipids recrystallize in the p’modification and transform rapidly into the piform. Furthermore, recrystallization of the bulk lipids starts at a higher temperature than the dispersed glycerides. The results indicate that hard fat can represent a supercooled melt when dispersed as nanoparticles dependent on storage temperature and storage time (cf. Section 1II.B).

B. Investigations on the Crystallinity and the Recrystallization Processof Melt-Emulsified Glyceride Nanoparticles Preliminary time-resolved x-ray studies already indicated differences regarding recrystallization from the melt of dispersed lipids compared with their bulk materials. In a next step, the recrystallization process should be systematically investigated. Owing to the limited access to synchrotron radiation x-ray sources, differential scanning calorimetry (DSC) was established as an alternative method to study the recrystallization behavior, the time course of polymorphic transitions, and the degree of crystallinity of melt-homogenized glyceride dispersions [63]. Knowledge of the crystallinity isof special relevance to evaluate the incorporability of lipophilic drugs

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Fig. 8. Synchrotron radiation wide-angle x-ray diffraction patterns of a 10 W% tripalmitatedispersionstabilizedbylecithinkodiumglycocholate(intensity Z vs detectorchannel): (a) recorded atroomtemperatureand (b) recordedatbody temperature.

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Fig. 9. Synchrotron radiation wide angle x-ray diffraction patterns of a 10 W% hard fat dispersionstabilizedby lecithidsodium glycocholate (intensity Zvs. detector channel): (a) recorded at room temperature and (b) recorded at bodytemperature.

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into the glyceride matrix. The chances to include foreign molecules, in particular those with a structurecompletely different from the matrix glycerides, are expected todecrease with the increasing order of the crystal lattice of the fatmatrix. Therefore, the timecourse of recrystallization and polymorphic transitions may correlate with the time course of expulsion of incorporated drug fromthe crystal lattice. The degreeof crystallinity of colloidal glyceride crystals is influenced by two factors:

1. The coexistence of recrystallized and amorphous (supercooled) particles, possibly as a result of a size effect as well as the crystallization probability 2. distortion of the crystal lattice by lattice imperfections owing to the high surface-to-volume ratio and the low periodicity within the nanoparticles

An estimate of the crystallinity index can be obtained from thermoanalytical data by relating the heatof fusion of the glyceride nanoparticles to that of the bulk glycerides. DSC heating scans of tripalmitate nanoparticle suspensions indicate an effect of the emulsifying agent on thethermograms. Tripalmitate dispersions stabilized by phospholipids only form ointmentlike semisolid gels directly after preparation. Thesegels display a sharp endothermal peak at around 56-57°C. In the presence of a coemulsifier such as the bile salt sodium glycocholate, tyloxapol, or poloxamers, tripalmitate dispersions do not gel. In thecorresponding thermogramsof freshly prepared systems, the p melting peak is at 54°C. The melting temperatures found for the dispersed triglycerides correspond to the melting point of the p’polymorph of bulk tripalmitate (Table 3). X-ray studies demonstrated, however, that thedispersed systems were in the p modification. In freshly dispersed, coemulsifier-stabilized dispersions, an additional endothermalpeakat around 40°C was detected which can be assigned to the a modification. In general, the a phase is transformed intohigher melting polymorphs within days or weeks. Obviously,phospholipids accelerate these polymorphic transitions, which isretarded in the presenceof cosurfactants. A similar effect of the coemulsifying agent was observed for hard fatsuspensions. Tentatively, this phenomenoncan beinterpretated as an epitaxical effect of the phospholipids due to structural similarities of the hydrocarbon chains of phospholipids and triglycerides. It was generally observedthat the melting point of dispersed glycerides was decreased by up to 13°C compared with the bulk material. This excludes unambiguous interpretation of DSC thermograms. As indicated by Table 3 and already outlined above, thermoanalytical data suggest

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Table 3. Melting Temperaturesof Glyceride Polymorphs and Melt-Emulsified Glyceride Dispersions Form a polymorph

p‘polymorph

Pipolymorph P polymorph

54-56 Dispersion (fresh) 56-59 Dispersion (stored)

Hard Fat (“C)

Tripalmitate (“C)

20-22 32-33 38-39

44-45 56-58

-

12-32 31-33

-

65-67

that the dispersed glycerides are present in the p‘polymorph. Synchrotron radiation x-ray diffraction clearly demonstrated, however, that the lipids were in the stablep or pimodification, respectively. Exact interpretation of DSC thermograms and unambiguous assignment of polymorphic forms thus require a priori knowledge about the crystal modifications, or else interpretation of DSC transition peaks might lead to erroneous assignment of polymorphic forms. Knowledge of the existing polymorph is, however, the basis for deriving the crystallinityindex of solid lipid nanoparticles from thermoanalytical data, since different polymorphs have a different heat of fusion. This aspect has not been taken into consideration by Muller et al. [52], who calculated the crystallinity index of glyceride nanoparticles from thermoanalytic data directly without assuring correct assignment of crystal modifications. The melting point depression observed with melt-emulsifiedlipid suspensions can be explained by the colloidal dimensions of the glyceride nanoparticles and their high surface-to-volume ratio. The excess pressure or solubility, respectively, experienced by the colloidal solid as predicted by increases its chemical potential. As a consethe Kelvin equation [M] quence, small crystals melt at a lower temperature than the bulk. The melting point, T, of small crystals of radius, r, can be derived from the modified Thomson equation [64], where To is the melting point o f the bulk material, y represents the interfacial energy, V the molar volume, and AHrus is the molar heat of fusion:

Furthermore, the thermoanalytical studies revealed that induction of recrystallization of both hard fat andtripalmitate dispersions requires more supercooling than in bulk. The supercooling amounts to approximately 20°C below that of the bulk glycerides inboth cases; that is recrystallization

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Biodegradable Colloidal Drug Carrier Systems

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TEMPERANR (K)

Fig. 10. DSC thermograms of a 10 W% hard fat dispersion stabilized by lecithin/ sodium glycocholate: (a) day of preparation, recorded from 20 to 60°C; (b) day of preparation, recorded from5 to 60°C; and (c) stored for 2 weeks at 4"C, recorded from 20 to 60°C. Heating rate: 10 Wmin.

from the melt starts at a much lower temperature in dispersed fat nanoparticles than in the bulk glycerides. As a result of the higher supercooling required for nucleation, isothermal recrystallization of hardfatnanoparticles is considerably retardedatroomtemperature (Fig. 10). DSC scans from room temperature to 60°C on the day of preparation indicate that hard fat nanoparticles have not yet recrystallized. Recrystallization can be induced by recording the thermograms from5°C onward. A broad peak with several maxima is then obtained pointing to the presence of different polymorphs. After 2 weeks of storage at 4"C, the thermogram recorded from room temperature displays a relatively sharp peak corresponding to thepipolymorph. The peak shoulderindicates the presence of the residual p'form. Investigations on the time course of polymorphic transitions indicate thatpolymorphic transitions generally proceed faster in the dispersed glycerides than in bulk. Recrystallization of phospholipid-cosurfactant-

242 Siekmann

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stabilized tripalmitate nanoparticles can already be detected 2 hs after preparation. Most of the triglyceride is present in the p modification with minor amountsof the CY form. After several days to a few weeks of storage, the CY polymorph has been completely transformed into the stablep modification. Incontrast, this transformation takes several monthsin meltcrystallized bulktripalmitate. In the nanoparticles, the heat of fusion of the p modification increases continuously until a plateau around 20 J/g is reached after approximately 2 months of storage (Fig. 11). The plateau value corresponds approximately to10% of the heat of fusion of the bulk material, indicating that the crystallinity of stored tripalmitate nanoparticles is similar to thatof the bulk. The initial increase during storage can be interpreted as a progressive annealing of the crystal lattice of the nanoparticles. The crystallinity index of glyceride suspensions calculated as the ratio of the heat of fusion of the nanoparticles compared with that of their bulk materials thus depends on the storage time and canincrease to up to 100%. Apart from the storage time, the crystallinity index of glyceride nanoparticles is affected by the type and amountof emulsifier, the presence of foreign compounds, andthe matrix material [63]. The crystallinity index of hard fat suspensions is generally lower than that of similarly composed tripalmitate systems. This difference is also reflected in the corresponding

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Fig. 11. Heat of fusion determined by DSC at different time intervals of a 10 W% tripalmitatedispersionstabilized by 2.4 W% soyalecithinand 0.4 W% sodium glycocholate.

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x-ray diffraction patterns and can be attributed to thelower crystallinity of the complex crystal lattice of hard fat which is more distorted than that of tripalmitate. Blending tripalmitate with monostearate (5% related to tripalmitate) results in a decrease of the crystallinity index compared with pure tripalmitate suspensions. Moreover, the monostearate apparently exhibits an a phase-preserving effect. X-ray data point to theinclusion of monostearate in the tripalmitate lattice without changes in the lattice dimensions. The creaYed lattice imperfections reduce the crystallinity of tripalmitate. The a polymorph is stabilized in these systems, because the presence of monostearate impairs the configurational changes of the tripalmitate molecules associated with the polymorphic transition. This observation is also described in the literaturefor bulk triglycerides [65]. The differences regarding the recrystallization behavior, the time course of polymorphic transitions, and the degree of crystallinity of melthomogenized glyceride nanoparticles and their bulk materials as revealed by thermoanalysis can be attributed to thedispersed state, thepresence of emulsifiers or otherforeign compounds, and thehigh surfaceholume ratio of the nanoparticles [63]. The results are of relevance to evaluate the (storage) stability of a composition andthe incorporability of drugs into the glyceride camer matrix, and to explain the possible expulsion of drugs from the crystal lattice.

C. Mechanism of GelFormationinPhospholipid-Stabilized Tripalmitate Suspensions

t.

Colloidal lipid suspensions display striking similarities to submicron-sized lipid O M emulsions concerningthe chemical composition of these dispersions. Furthermore, suspensions of solid lipidnanoparticles can be manufactured by a melt-homogenizationprocess which is in principle similar to the production of parenterallipid emulsions. The obvious similarities between lipid emulsions andlipid suspensions with respect to chemical composition and production methodmight lead to theconclusion that lipid suspensions could be regarded as“lipid emulsions with solidified droplets.” However, in contrast to lecithin-stabilized lipid emulsions, the preparation of similarly composed lipid suspensions, that is, by employing phospholipids as theonly stabilizer, can result in the formationof semisolid gels[18,52]. The observed instability of lipid suspensions containing the same type and amount of emulsifiers as parenteral lipid emulsions thus indicates basic physicochemical differences of these twotypes of colloidal dispersions. As already outlined above, the prevention of gel formation is possible by the addition of highly mobile cosurfactants such as bile salts [B]or the nonionic polymer

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tyloxapol[46], but results obtained with different types, amounts, andcombinations of emulsifiers indicated that electrostatic repulsion or steric stabilization do notrepresent the exclusive prerequisites to prepare stablesolid lipid nanoparticles [45]. Results from the production of glyceride nanoparticles further point to that gel formation might be attributedto recrystallization of the dispersed lipid, since gel formation was generally not observed above the melting temperature of the glycerides [66]. X-ray studies did, however, not display basic differences regarding the crystalline state of the semisolid gels and theliquid suspensions [62]. In order tostudy the physicochemical processes underlying the gel formation phenomena, structuralinvestigations of tripalmitate nanoparticle dispersions and gels were initiated by transmission electron microscopy of freeze-fractured specimens (FF-TEM) and of frozenhydrated specimens (cryo-TEM) [67]. FF-TEM of lecithidcosurfactant-stabilized tripalmitate supensions revealed that the emulsified glycerides recrystallize as anisometrical particles of predominantly plateletlike shape (Fig. 12a). Larger particles clearly display a layered structure with terraces, steps, and kinks (Fig. 12b). The prevailing existence of platelets indicates the preferential formation of certain crystal faces. The crystal shape of tripalmitate nanoparticles resembles melt-crystallizedmicro- and macroscopic tripalmitate crystals of the /3 modification [68,69]. Owing to these similarities, the prevailing crystal face of tripalmitate nanoparticles could be identified as the (001)-face. As a consequence of the characteristic order of molecules in the crystal lattice, the different crystal faces exhibit different physicochemical properties such as polarity and surface tension. In melt-crystallized macroscopic tripalmitate crystals, the (001)-face, which contains the methyl end groupsof the hydrocarbon chains, is nonpolar. However, owing to its preferential formation in the aqueous medium, it can be deduced that the (OOl)-face of tripalmitate nanocrystals is hydrophilic. This was confirmed by decoration of the fractures with water, which condenses preferably on the large (001)-face [67]. The detected hydrophilicity of the (001)-face points to the association of surfactant molecules with this nanocrystal face. FF-TEM of tripalmitate suspensions stabilized by phospholipids only which form semisolid, ointmentlike gels displays a coherent network of Fig. 12. Transmissionelectronmicrographs of melt-homogenizedtripalmitate. (a) Top: 10 W% tripalmitate suspension stabilized by 1.2 W% soya lecithin and0.4 W% sodium glycocholate (thebarrepresents230 nm). (b) Middle: 10 W% tripalmitate suspension stabilized by 1.2 W% soya lecithin and 0.4 W% sodium glycocholate(thebarrepresents 150 nm).(c)Bottom:10 W% tripalmitate gel containing 1.2W% soya lecithin (the bar represents 250 nm).

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lamellar crystal platelets with a thickness corresponding to that of tripalmitate nanocrystals in suspension [67]. From the view onto the network plane it seems that thegel structure is formed by individual platelets which have agglomeratedto a cardhouselike structure (see Fig. 12c). It is striking that aggregation of the platelets occurs via the lateral crystal faces and not via the (001)-face, which also represents the prevailing crystal face in the gels. Owing to its large expansion, it can be deduced that the (001)-face exhibits a low surface tension, pointing to preferential phospholipid adsorption on the (001)-face. From the electron microscopic pictures, the tripalmitate gels can be characterized as ultrafine three-dimensional networks formed by aggregated tripalmitate crystals. The aqueous phase interpenetrating thenetwork is completely immobilized inthese structures. The physicochemical processes during cooling can be directly monitored by polarized light microscopy of crude lecithin-bile salt-stabilized tripalmitate dispersions [45]. The dispersed lipid melt formsspherical, isotropic droplets which transform into anisometrical, optically anisotropic particles when the melt recrystallizes (Figs. 5 and 13). Crystallization starts from the particle surface, pointing to an epitaxical effect of the emulsifier molecules present at the droplet interface. Beside the fat particles, the typical Maltese cross-like structures representing multilamellar vesicles (MLV) can beobserved. The high number of MLV indicate that a substantial portion of phospholipids is bound in vesicles despite the fact that the emulsifier was dispersed in the melted lipid phase prior to the emulsification process. The existence of small unilamellar vesicles in colloidal tripalmitate suspensions was confirmed by cryo-TEM [67]. The numberof vesicle structures in tripalmitate suspensions is, however, smaller than in commercial parenteral O/W emulsions, althought the lecithin concentration was higher in the suspensions. The microscopic observations oncrudeand submicron-sized tripalmitate dispersions clearly demonstrate that recrystallization of meltemulsified tripalmitate is associated with the formation of anisometrical particles, thereby inducing a substantial increase in particle surface. The newly generated surfaces need to be stabilized immediately in order to avoid particle aggregation and the formationof two-phase gel systems. Phospholipids belong to the group of insoluble swelling amphiphilic lipids. In aqueous media,they spontaneously formvesicles. Sincephospholipid molecules do not tend to leave the bilayer, their concentration in the aqueous phaseis very low. Moreover, phospholipids only exhibit a moderate mobility inaqueous media.Owing to this phase behavior, lecithin molecules are not expected to be sufficiently mobile to stabilize newly generated tripalmitate surfaces immediately. The use of phospholipids as the only emulsifier, even in excessamounts, thus results in gelformation. In contrast,

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Fig. 13. A polarized light microscopy picture of a crude dispersion of tripalmitate stabilized by phospholipids and sodium glycocholate after termination of the recrystallization process (magnification: X 160). The suspended particles are optically anisotropic and exhibit anisometricalshapes.

the employed cosurfactants are micelle-forming agents. Micelles are highly dynamic structures and representa reservoir of surfactant molecules from which the emulsifier can be recruited when the emulsion droplets turn into anisometrical suspension particles. Owing to thehigh solubility and mobility of the cosurfactants in the aqueous phase, they are able to stabilize the freshly generated surfaces immediately. Based on the experimental results from electron microscopy and on theoretical considerations regarding crystal structure andmolecular mobility, a model as illustrated in Fig. 14 has been developed to describe the instability phenomena of colloidal tripalmitate suspensions. The recrystallization of the colloidally emulsified tripalmitate results in an increase in particle surface area due to thechange in the overall particle shape and a sudden local lack of emulsifier in the particle surfaces. The time, cl, corresponds to the time which the recrystallization process takes. During the emulsification process, the excess of phospholipids has formed small vesicles (SUV) which are randomly distributed in the continuous phase and

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Suspension

Emulsion

Bile salt micelles

Fig. 14. Model of the gelformationmechanism in phospholipidstabilized triglyceride dispersions(SUV, small unilamellar vesicle). which exist in a metastable state after finalization of the emulsification which the phospholipid process. The time, t2, corresponds to the time molecules would need to leave the vesicles and to diffuse to the newly created insufficiently stabilized particle surfaces. The time, tz, is, however, significantly longer than the time, C,,resulting in an instability of the dispersed state andgelation via the lateral faces of the crystals. In contrast, dispersions which contain an admixture of a soluble, micelle-forming component such as bile salts are metastable; that is, the dispersed state can be maintained over a considerably long timeof storage. The soluble, micelle-forming component is able to cover thenewly created tz, for the surfaces in a time, t3, which is significantlyshorter than the time, phospholipid molecules and which is short enough that no gelation of the dispersed particles, that is aggregation via unprotected crystal faces, can occur (see Fig. 14). Furthermore, it has to be considered that phospholipids and cosurfactants adsorb to different nanocrystal faces. The TEM results suggested that phospholipids are preferentially adsorbed to the(001)-face, so that the lateral faces are insufficiently stabilized and aggregation can occurvia these faces. It can be assumed that the lateral faces are covered to a greater extent by the cosurfactants. Moreover, it seems likely that phospholipids-

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in contrast to thehydrophilic cosurfactants-are at least partially incorporated into thetripalmitate crystal lattice owing to similarities in molecular geometry. Crystal growth can thus proceed over adsorbed phospholipid moleculesbut is interrupted where cosurfactants are adsorbed tothe nanocrystal surface. From the results outlined above, it can be concludedthat lipid suspensions do not simply represent “lipid emulsions with solidified droplets,” since the solidification of the dispersed phase is associated with basicdifferences in the physicochemical behavior of lipid emulsions andsuspensions as illustrated by the more complex and complicated stability aspects of the latter. It should be noted that theparticle shape of melt-homogenized lipid nanoparticles might be considerably different from thatof microparticles of the same lipid matrix, which may displaya predominantly spherical particle shape. The differences can be attributed, among others, to the different production processes. Lipid microparticles are generally prepared by methods which include a rapid cooling step forcing the lipid to solidify immediately in the amorphous stateor in an unstable polymorphic modification of low crystalline order with a high concentration of lattice imperfections. In this case, the low-ordered, polycrystallinic crystal lattice does not exert a strain on the particle shape, and surface minimized; that is, almost spherical particles can be formed. In contrast, tripalmitate nanoparticles represent extremelysmall single crystals. Sincethe production process does not include a rapid cooling step, the colloidal lipid particles can crystallize slowly, so that the stable polymorph can be formed usually exhibiting a highly ordered crystal lattice whichis reflected by the characteristic anisometrical crystal shape. It is thus not possible to transfer results obtained on lipid microparticles to theproperties of lipid nanoparticles. It has to be taken into consideration, however, that not alllipid materials will form colloidal particles with anisometrical shape as for tripalmitate. Colloidal suspension particles of complex lipid mixtures, such as WitepsolW35, exhibit a spherical shape [l81 in the dispersed state, and the structure of the particle cores is significantly different from thatof the tripalmitate nanoplatelets. Whereas the internal structure of colloidal tripalmitate particles resembles a stack of cards or lamellae, the core of colloidally dispersed Witepsol W35 particles seems to be polycrystalline with low order within each crystallite. D. incorporation of Drugs into Solid Lipid Nanoparticles Solid lipid nanoparticles have been demonstrated to represent crystalline colloidal particles [62,63]. Colloidal systems withthe dispersed lipid existing

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in the solid amorphous state have not yet been described in the literature. The crystalline state of the particle matrix has to be taken into account when estimating the potential of incorporating drugs into these camer systems, since the inclusion of foreign compounds intoa crystalline lattice is generally limited. To a certain extent, foreign molecules can be included in lattice imperfections. It has to be considered, however, that on the one hand the the inclusion capacity is number of lattice defects is limited and consequently low. This inclusion mechanism can thus only be taken advantage of with highly active compounds which do not requireadministration of high therapeutic doses. On the other hand, it was shown that the crystallinity of glyceride nanoparticles is increasing with storage time owing to annealing of the crystal lattice [63].As with the kinetics of polymorphic transitions, annealing is accelerated in the colloidal state and drug molecules located in lattice imperfections can beexpelled from the lattice with increasing annealing, leading to stability problems onstorage. Results from our laboratory [45] suggest that the incorporability of lipophilic compounds into melt-homogenized glyceride nanoparticles depends predominantly on whether the drug can be included in the crystal glyceride lattice without strongly disturbing its structure, that is, that the molecular structureof the drug fits into thelattice, or whether the drug and the carriermatrix can forma crystal lattice together. If this is not possible, the drug is expelled from the camer matrix on recrystallization or, respectively, polymorphic transitions of the glycerides, and it precipitates in the aqueous phase as was found, for instance, for menadione and cortisone. There are, however, a number of drugs which have been demonstrated to be incorporable in lipid suspensions. In a conference contribution, Schwarz et al. [70]reported that they succeeded in incorporating tetracaine and etomidate in solid lipidnanoparticles without severe reduction of the physical stability. We were able to show that, in contrast to cortisone, the structurally related compound prednisolone could be incorporated into tripalmitate nanoparticles, and these drug-loaded camers proved to be stable for several months [45].This observation indicates that even small structural changes, for example, the steric arrangement of functional groups, can be of relevance for incorporation in the glyceride lattice. The incorporability of retinol and ubidecarenone suggests that long, apolar side chains of the drug compound might be beneficial for inclusion in the glyceride matrix [45].The use of lipophilic prodrugs, for example, estersof long-chain fatty acids, might thus represent an approach to improve the incorporability of drugs. Another approach to increase the drug-incorporation capacity is controlled solidification of melt-homogenized glyceride nanoparticles as lowordered crystals of the a phase or even in amorphous form. Because of

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their completely disordered state, amorphoussolids exhibit a high incorporation capacity for foreign compounds. It has to be considered, however, that thestabilization of amorphously solidified triglycerides is energetically unfavored because of the extremely low activation energy required to transform amorphoustriglycerides into thea modification. It might therefore be more promising to concentrate effortson the stabilization of the a phase. Owing to thelower crystalline order in the metastablea polymorph as well as to the rotational motion of the hydrocarbons which reduce the crystal packaging in the a form, theincorporability of foreign compounds intothe a phase is increased compared with the stable p modification. An attempt to stabilize thea phase by the use of asymmetrical glycerides or cocrystallizing mixtures of different matrix lipids is currently being investigated in our laboratory. Considering the observed acceleration of polymorphic transitions for colloidal dispersions of tripalmitate as well as of hard fat, longterm stabilization of the a modification in colloidal dispersions might be extremely difficult to achieve. Recent results demonstrate that on the other hand the drug can affect the recrystallization of thecamer lipids. Recrystallization studies on ubidecarenone-loaded tripalmitate nanoparticles by DSC and time-resolved x-ray diffraction indicated that incorporationof the lipophilic drug can cause a significant reduction of the recrystallization temperature of the camer [71]. Moreover, in drug-loaded camer systems, anaccelerated polymorphic transition into the stable p modification was observed, since the a polymorph of the camer matrix could not be detected in ubidecarenonecontaining tripalmitate nanoparticles. These results furthermore demonstrate the increasing complexity of solid lipid nanoparticles when loaded with drugs. The situation is even more complicated by the fact that drugs differ in their physicochemical properties. In addition, the therapeutic dose depends on the drug requiring different payloads. Therefore, drugconcentration has tobe considered as an additional complicating factor. IV. SUMMARY AND FUTURE PROSPECTS

Colloidal dispersions based on biodegradable solid lipids have been found to be potential drug-carriersystems of complex physicochemicalproperties as expressed by their recrystallization behavior and their tendencyto form semisolid gels, which have prevented theirsuccessful establishment as drug carriers forseveral decades. The results regarding the gel-formation mechanism reported hereemphasize the importanceof physicochemical characterizationforproduct development. Despitetheir complexity, solid lipid nanoparticles represent an interesting alternative to conventional drugcarrier systems such as polymer nanoparticles, liposomes, and lipid emul-

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sions, since it has been demonstrated that stability problems are associated with the crystalline state of the dispersed lipids and can be circumvented by a proper choice of matrix constituents, emulsifying agents, and production parameters. Manufacturing of solid lipid nanoparticles can avoid the use of potentially harmful additives, so that toxic residues from the production process as often associated with the preparation of polymeric nanoparticles can be circumvented. Owing to the biodegradability of physiological lipids, accumulation of the camermaterial on repeatedadministration is not expected. In a comparative in vitro cytotoxicity study using human granulocytes, it was found thatpolyester nanoparticles exhibited severe cytotoxic effects at 0.5% concentration, whereas no cytotoxic effects were observed with solid lipid nanoparticles [72]. In vivo toxicitystudies of solid lipid nanoparticles, however, have not yet been published. The possible advantages of solid lipid nanoparticles compared with lipid emulsions or liposomes is their stability against coalescence and the reduced mobility of incorporated drug molecules preventing drug leakage from the carrier. As outlined, careful physicochemical characterization is, however, required to ensure thesolid physicalstate of the lipid in the colloidal state. A satisfactory long-term stability of lipid suspensions can be achieved considering the results obtained regarding the mechanism of gel formation (cf. Section 1II.C). A general limitation of crystalline solid lipid nanoparticles is, however, represented by the limited inclusion capacity for most drugs, whereas others can be incorporated in high amounts [71]. Drug incorporability might be increased by chemical modificationsof the drug,by controlled crystallization and stabilization of low-ordered crystal polymorphs of the carrier material,or by solidification of the lipid melt in amorphous form, as has already been discussed (cf. Section4II.D). Colloidal solid lipid dispersions represent complex colloidal systems which may contain different types of coexisting colloidal structures such as solid lipid particles, nonsolidified particles of supercooled melt, micelles, mixed micelles, and/or vesicles. Therefore, it has to be considered that the distribution of drug molecules might be influenced by the type and concentration of coexisting structures butalso by the incorporation process, physicochemicalcharacteristics of the drug, and theemulsifier. The extremely small particle size of solid lipid nanoparticles, which can be less than 50 nm, might be beneficial with respect to drug targeting. Small carrier size generally favors reduced uptake by the RES. Moreover, intravenously injected particles smaller thanthefenestrations of the endothelial wall, that is, below 150nm, might be ableto leave the vascular compartment through these fenestraein the sinusoids of the liver, spleen, andbone marrow orat locations where the basal membrane of the

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endothelium is damaged; for example,at sites of inflammation or in tumor tissues. Drug targeting might also be possible by surface modification of solid lipid nanoparticles (cf. Section 1I.E). The drugrelease from solid lipid nanoparticles is probably not diffusion controlled as in lipid emulsions owing to thereduced mobility of drug molecules incorporated in the solid matrix. It can be expected that the ratelimiting step of drug release is the degradation of the lipid carrier matrix. Drug release from lipid suspensions should therefore sustained be compared with lipid emulsions. In contrast to polymeric nanoparticles, lipid nanoparticles can be degraded by lipases inthe blood. Enzymatic degradationby lipases depends, among others, on the hydrocarbon chain length of the lipids. Hence the degradation rate and drug release can be controlled to a certain extent by the use of differently composed carrier matrices as well as by the choice of emulsifier. Moreover, as has been demonstrated by x-ray diffraction studies [62], it is possible to compose the carrier matrix so it is solid at storage temperature but liquid at body temperature, giving rise to fast drugrelease similar to thatfrom lipid emulsions. Besides parenteral application, solid lipid nanoparticles are also suitable for other routes of administration and might be an interesting carrier system forthe peroral administration of poorly water-soluble drugs with a low peroral bioavailability. An advantage of the emulsified lipid particles might be their improved wettability in gastrointestinal fluids. Whereas the poor absorption of certain drugs can be related to their poor wettability, incorporation into solid lipid nanoparticles provides completely wettable carriers. The lipid particles will probably bedigested similarly to food lipids. Because of their high dispersivity, solid lipid nanoparticles exhibit a high specific surface area for enzymatic attack by intestinal lipases. Enzymatic degradation of the lipids leads to therelease of incorporated drugs in molecularly dispersed form or solubilized in micellesformed by degradation products and bile salts facilitating their solubilization in the intestine and subsequent absorption. As a concluding remark, it should be emphasized that the use of colloidal drug carriers based on solid lipids is at present-compared with other colloidal systemssuch as polymer nanoparticles, liposomes, and lipid emulsions-at its early phase of the development, although the conceptional idea is not new. The authors have tried their best to present a comprehensive review of what is state of the art regarding solid lipid delivery systems based on own experimental data and experiences as well as on published material of other research groups working inthe field. We would like to draw the reader’s attention to the fact that much data have unfortunately been published exclusively as conference abstracts. These abstracts generally lack substantial scientific data, since they often only contain a

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brief introduction to the field of research, hypothetical advantages of the proposed delivery systems, and a short statement of what the conference contribution is dealing with. It is very difficult to interpret results from a conference abstract if not even the composition of the studied systems is disclosed. Nevertheless, we have tried to take into account the results extractable from conference proceedings as far as possible. Since in many cases comprehensive scientific publications are yet outstanding, conference abstracts represented theonly source of information. Itis hoped that these data will soon be available to thescientific communityas full-length papers in order toelucidate further thepossibilities and limitations of solid lipidbased drug-delivery systems. REFERENCES 1. K. J. Widder, A. E. Senyei, and B. Sears, Experimental methods in cancer therapeutics,J. Pharm. Sci., 71:379 (1982). 2. P. K. Gupta, Drug targeting in cancer chemotherapy: A clinical perspective, J. Pharm. Sci.,79:949 (1990). 3. E. Tomlinson, Microsphere delivery systems for drug targeting and controlled release, Int.J. Pharm. Tech. Prod. Mfg., 4:49 (1983). E. Doelker,Drug-loadednanoparticles4. E.AWmann,R.Gurny,and Preparation methods and drug targeting issues, Eur. J. Pharm. Biopharm., 39:173 (1993). 5. B. Kante, P. Couvreur, D. Dubois-Krack, et al., Toxicity of polyalkycyanoacrylate nanoparticles. I. Free nanoparticles, J. Pharm. Sci., 71:786 (1982). J. Carpenter,andD.R.Alford,Spontanousfusionof 6. B.R.Lentz,T.

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l0 Pharmaceutical Aspectsof Liposomes: Perspectives in, and Integrationof, Academic andIndustrial Research and Development Rimona Mat'galit and Noga Yerushalmi Tel Aviv University, Tel Aviv, Israel

I.

Introduction

11. Liposomes: Definition, Needs for, and Outline of Their Advantages and Drawbacks A. Liposomes: A Family of Structurally Related Microparticles B. Deficiencies of Treatment with Free Drugs that Justify the Pursuit of Drug Delivery Systems and Consideration of Liposomes for the Task 111. Selection of Liposome me/Species: Views and Criteria from Academic and Industrial Research and Their Proposed Integration A. Liposome m e , Size, and Production Issues B. Lipid Composition IV. Targetedmodified Liposomes: An Interesting and Exciting Scientific Tool, But Can They Be Made into Products (Especially Immunoliposomes)? A. Targeting: Definition, Stages, and Status B. The Means Through Which Liposome Targeting Has Been Attempted and Their Feasibility for Liposomes as Pharmaceutical Products

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V. Liposomes as a Sterile, Pyrogen-Free System with Pharmaceutically Acceptable Shelf Life, Stability, and Dosage Sterile, A. Pyrogen-Free 277Liposomes B. Shelf Life, Stability, and Dosage Forms

VI. Liposome Characteristics (Percentage of Encapsulation, Kinetics of Release, Biological Activity)-in Basic Research and in Quality Assurance A. Efficiency of Drug Encapsulation B. Kinetics Release of Drug C. Biological Activities of Liposome-Encapsulated Drugs

VII. Summary and Future Prospects

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INTRODUCTION

The scientific literature is rich withcomprehensive reviews of liposomes as drug-delivery systems (DDS) (see, e.g.,refs. 1-11). Each review presents a unique and specific point of view, whichcan range from thestrictly physicochemical, through various biological levels, to the clinical. Furthermore, stemming from the application for which theyare reviewed, the prospectof liposomes as pharmaceutical products is inherently presentin these reviews even when not specifically identified and addressed. Onthis background, it was deemed essential tofirst address aquestion that might be raised by the reader, which is why yet another manuscript on liposomes as pharmaceutical products. Liposomes as DDS are among research topics that are being vigorously investigated in both academic and industrial laboratories,with different outlooksbut common goals and endproducts.Addressing fundamental questions while being continuously innovative are preceived to be among the responsibilities of the academician in this field. Addressing developmental issues such asscale-up, shelf life, dosage forms, long-term stability, and cost-effective production are perceived to be among the responsibilitiesof the industrial scientists. Yet the R&D of liposomes as DDS encompass industry-oriented issues that should already be taken into account at the basic academic stage, as well as questions that arise at the industrial stage that have distinct “basic science” components. Stemming from this premise, the objectives of this chapter are to address those issues and questions and attempt to provide an integrated view that would (1)be of use and contributeto themutual understandingof

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the topics most critical to investigators in both industry and universities and (2) serve as guidelines for newcomers to thefield. It is hoped that, through this, this chapter will become a modest addition to, rather than a repetition of, the existing literature. In order tolay out a common groundfor the discussion of liposomes as DDS, the first part of this chapter provides a brief definition of liposomes and discusses the unmet therapeutic needs that justify their pursuit as DDS and their advantages and drawbacks.The subsequent partsof this chapter attempts toprovide the proposed integrated approach, addressing the following issues: (1) types of liposomes, with respect to scale-up, production, and cost; (2) preparation and production of sterile and pyrogenfree liposomes; (3) shelf life, stability, anddosage forms; (4) targeted (mostly surface-modified) liposomes as exciting research tools and as pharmaceutical products; and (5) characterization of liposome-drug systems (such as encapsulation efficiencies, drug-release kinetics, and biological activity) for academicstudies and for industrial quality assurance. The final part will offer conclusions, taken from theposition of an academician, but hopefully relevant for all liposome investigators. The authors are well aware thatthe line of division between academic and industrial liposomal R&D is quite often indistinct. Investigators in academics research could be engaged in questions that bear on liposomes as pharmaceutical products, whereas, more so, investigators in industry could be engaged in basic liposome research. In order toavoid any confusion or resultant grievances (however unintentional), it is stressed here that throughout this chapter the termsacademic and industrial research will be used to indicate the nature and tendency of the research (i.e., basic or applied) and not the physical definition of the environment or community where it is performed. II. LIPOSOMES: DEFINITION, NEEDS FOR, AND OUTLINE OF THEIR ADVANTAGES AND DRAWBACKS A. Liposomes: A Family of Structurally Related Microparticles

Liposomes, frontline microparticulate carriers investigated and developed for the task of drug delivery, are artificial microscopic and submicroscopic particles made of lipids and water alone that were developed thirty years ago. [l].A variety of liposome species with respect to shape, type, size, and composition have been developed in the course of the last three decades, extending this type of particle into a family of structurally related delivery systems. For the novice in the field, it is worth noting that in the

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early days of liposomes, there was an ongoing debate between the terms vesicle and liposome, the legacy of which ispresent in the names, abbreviations, and acronyms used for various liposome types. Multilamellar vesicles (MLV) are the oldest liposome species, and they have been described frequently enough [1,2,5-7,12, and references therein] that only a brief summary is merited here. They are composed of concentric shells of lipid bilayers, with water between shells and an inner aqueous core. Liposomes of this type form spontaneously on the proper interaction of lipids with water, provided the right choice of lipids and technical conditions have been met. A typical MLV will have 8-15 concentric shells, and run to sizes that can range from (roughly) 0.5 to several micrometers in diameter. Consequently, a typical MLV preparation will be quite heterogeneousin terms of liposome sizes. Unilamellar vesicles (ULV) are composed, as their name indicates, of a single lipid bilayer and an inner aqueous core[1,2,6,7,8,10,11,13,14, and references therein]. Dependingon the method of preparation, ULV can be made in various size ranges, from as small as 20 nm to several micrometers in diameter, with a significantly smaller size distribution within a preparation comparedwith MLV. Both MLV and ULV can be made from a wide (although not infinite) range of lipid and lipid mixtures [l-161 and can accommodate hydrophilic and hydrophobic drugs in their aqueous and lipid compartments, respectively. Furthermore, with careful selection of liposome type, encapsulation of matter that is as small as the lithium ion up to macromolecules as large as genetic material (of several hundred thousand daltons) can be achieved. Scanning the threedecades of liposome literature makesit quite clear that besides the abbreviationsMLV and ULV, there aremany more names and types of liposomes the distinction among and the classification of which are frequently based onmethodsand/or devices of preparation with/ without contribution of the resultant size-related properties. In fact, this “horn of plenty” can be a sourceof confusion to thenovice in the field and a burden on both veterans and newcomers, as exemplifiedby the two cases outlined below. The first case concerns liposomes that have been classified as large unilamellar vesicles (LUV). Oneof the major methods forproducing such liposomes is the reverse-phase evaporation technique[13]; such liposomes have been abbreviated REV and are usually classified as LUV. A REV preparation can have a relatively wide size distribution reaching the range of 600 nmin diameter. Subjected,as frequently recommended, to postpreparation fractionation steps, the size distribution of an REV preparation can be quite narrow together with a reductionin the dominantsize. Yet, regard-

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less of whether such fractionation has or has notbeensubjected to postpreparation steps, these sytems will all be referred to as REV and considered to be LUV. Other approaches for producing unilamellar liposomes are based on extrusion devices, such as the LIPEX extruder [18]. Liposomes produced with this device have been named LUVET and are also referred to as LUV;typical preparations of this type have a relatively narrow size distribution. Sizewise, they can run from30 to several hundred nanometers in diameter, depending on the pore sizes of the filters used. The second case concerns SUV, which is the the acronym for small unilamellar vesicles as well as for sonicated unilamellar vesicles. SUV with a size range of 20-nmdiameters are usually obtained throughthe use of probe sonicators and are one of the oldest types of liposomes. One also finds the acronym SUVin usefor unilamellar liposomes thatare in sizeranges of 40, 60, and even up to100 that have been made by a variety of methods other than probesonicators. Yet another device for the production of unilamellar liposomes is the Microfluidizer [19], which yields (depending on operating conditions) liposome preparations with a narrow size distribution that (de0 0to pending onoperating conditions) can range in diameter from under1 several hundred nanometers.Named MEL, for microemulsified liposomes, such liposomes canqualify as either SUV orLUV. Besides the multiplicity in the field with respect to liposome names and classifications that these few examples illustrate, there is also an inherent weakness in using methods and devices for liposome preparation as major criteria for classification. Clearly some methods and devices can become obsolete and othershave not yet been invented. In an attempt to ease the current liposome namehype situation, and atthe sametime offer an informative classification, a simplified and straightforward two-tier approach thateliminates references to preparation methods and/or devices ishereby proposed.This two-tier approach is used throughout this chapter with the hope thatit will find favor with and be of use to both veterans and novices in the field. The first tier, which is the primary classification, is based on a major structural difference between liposomespecies which is simplythe number of lamellae. The line of division is set between one and more than one lamellae. On this premise, there areonly two majortypes of liposomes, for which there is no need to use names other thanthe traditional terms MLV and ULV. Liposome preparations, whether MLV or ULV, might also contain a (usually small) share of liposomes denoted oligolamellar that have few, such as two to three, lamellae. These are considered to be a minor type of liposomes, as to date there has been neither reports on applications for which such liposomes are specifically sought, nor procedures for their preparation as the major dominant particle.

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The second tier, which willbe illustrated below by several examples, is a subclassification according to thedominant size range and/or size distribution, whichever is the more relevant. For example,in a reported study, the investigated liposome type would be simply classified as ULV 80 f 30 nm. These details would allowthe reader infer to (1) that this preparation is quite homogeneous and truly unilamellar and (2) that the majority of these liposomes are sufficiently smallso that, if endowed by proper receptoraffinity, they canalso be endocytosedby nonphagocyticcells through the coated pit mechanism [7].Identifying an investigated liposome preparation ULV as 650 f 200 nm wouldbe a clear indication to the reader that this preparation probably contains not only unilamellar but also olgiomallelar liposomes (i.e., liposomeswith more than one lamellae but less that the number designated todefine MLV). A preparation identified as MLV 0.5-2.5 pm would clearly indicate that this is a rather heterogeneous liposome preparation that has probably not beensubjected to any fractionation procedures and contains particles that candiffer fivefold in diameter. Besides offering simplification and being informative, it is argued that this classification will leave descriptive terms such as small or large to the personal inclination of the investigators and the readers, and it will help clarify differences between liposomes and other lipid-based particles that are under development as delivery systems. The latter are, for the most, proprietary technologies such as plurilamellar liposomes that are reported 0 0pm [20]; to contain well over 100 lamellae and to range in size up to 1 multivesicular liposomes (DepoFoam),also referred to as MLV, each particle of which consists of several wall-sharing large unilamellar liposomes, with a size that can reach 100 pm [21]; solvent dilution microcamers (SDMC) [22-231; and transferosomes [24-251 developed specifically for the transdermal administration to intact skin. B. Deficiencies of Treatment with Free Drugs that Justify the Pursuit of Drug Delivery Systems and Consideration of Liposomes for the Task

Whether viewed from the clinical, the physiological, the financial, or any other of the many aspects involved, treatment with drug-loaded delivery systems is substantially more complex that with free drug. Therefore, investment in research and developmentof liposomes* has to bejustified, on one hand, on theexistence of therapeutic needs that aretruly unmet with *Although most of the issues discussed with respect to the needs for liposomes apply to otherdrug-deliverysystem also, the discussion will refer to liposomes alone, asthey are the topicof this chapter.

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the free drug and, on the other hand, by the ability of the liposomes to provide significant improvements in clinical outcomes that can be developed into established treatment modalities. Tracing the fate of a drug administered in its free form, dissolved or dispersed in an appropriatevehicle, can provide a glimpse of those unmet needs thatjustify the pursuit of liposomes as DDS. Exampleswill be given below for two cases, from which the general picture can be inferred, not only for systemic but also for nonsystemic (such as topical and regional) administrations. The first example is taken from tumor treatment by chemotherapy, where the deficiences of treatment with free drug come fromthe effects of the drug on the biological environment andvice versa. It is well knownthat conventional chemotherapeutic drugs, such as doxorubicin, vinblastine, vincristiine, 5-flourouracil (5-FU), and many others [26] are virulent poisonous substances, a property which is an asset with respect to drug effects on malignant cells but is disastrous when the drug interacts with any other (healthy) cells. Most frequently, such drugs are administred in their free form systemically. Lack of targeting to the tumor sites results in the indiscriminate distribution of the drug over the whole body, reducing the efficacy of the treatment and at the same time leading to undesirable side effects and toxicity. Free-form administration of drugs exposes themto the elements of the biological environment, and they are often prematurely cleared, which is especially critical for drugs that act at a specific stage of the cell cycle. The drugs (in free form) are also susceptible to inactivation and/or degradation and to scavenging by endogenous carriers such as serum albumin and serum lipoproteins, which can aid as well as abet drug access to the target zones. The second example concernswound healing, where growth factors, such as epidermal growth factor (EGF), fibroblast growth factors a and b (FGF-a), (FGF-b), platelet-derived growth (PDGF), transforming growth factors (U and P (TGF-(U,TGF-P), andmany others, are under development as topical therapeutic agents for the acceleration and stimulation of the self-healing process in wounds and bums [27-351. Current dosage forms used in both basic and developmental growth factor studies that are appropriate for the future treatmentof patients center on free growth factor in vehicles such as solutions of saline or other physiological buffers (poured onto thewound or soaked into a gauze dressing), cellulose gels, and collagen sponges[27-381. The use of such vehiclesand procedures corresponds to theimmediate (or almost immediate) exposure of the total growth factor dose to the wound. As in the previous example, the mutual effects of the drug and of the biological environment underminesuccessful therapy with the freedrug.

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Growth factors are especially vulnerable to the biological environment, which is enzymatically hostile to polypeptides. In addition to the enzyme-catalyzeddegradation, growth factors that are free in the wound are subject to continuous clearance from the wound area and can be scavenged through binding to thevarious particulate and soluble wound fluid components. Owing to their susceptibilityto theproteolysis, the growth factors are also prevented (for the most part) fromexerting any self-targeting within the wound area toward their sites of action (i.e., their specific receptors). The small share of the dose that doesreach the target form is often too low to affect a significant difference, especially as growthfactors are agents that act at a specific stage of the cell cycle [39-411. Hence, effective therapy also requires a continuous supply of active growth factor near enoughto its sites of action for a sufficient duration [29,31,40]. The other side of the coin, namely, the detrimental effects of growth factors on the biological system, are seen on attempts to increase the bioavailability at the target by dose escalation. It is well known, and not only for growth factors but for hormones in general, that substantially high (local) concentrations, even if transient, can result in adverse effects and toxicity [31,39]. In order towrest some benefit from the application of growth factors despite the deficiencies described above, current treatment regimens use multiple dosing at frequent intervals; for example, twice daily for 70-100 days [41]. Such regimens prevent the implementation of state of the art (1)minimal interferapproaches to good wound management that advocate ence with the wound; (2) maintenance of a moise wound environment, which wasfound to be most conducive to theself healing; and (3) the use of occlusive dressings that are changed once every few days [43-501. Obviously, treatment regimens that interfere with the healing will detract from any benefit that a medication for heating can provide. Although the deficiencies of free-drug administration, defined above, clearly call out for ways and means to overcome them and in principle a delivery system is among the most viable solutions, such systems need to meet particular attributes in order toqualify for the task. These attributes can be divided into three categories. The first two categories have been amply discussed, are considered to be within the realmof basic science, and comprise qualities that the carrier needs in order to function (denoted category I) and in order to avoid the situation of replacing one set of problems by another (denoted category 11). The third (denoted category 111) comprises qualities required in order to develop the drug-carrier systems into pharmaceutical products that can then be implementedestabinto lished treatment modalities. Properties included in category I are (1) the ability to provide mutual protection of the carried drug and thebiological environment; (2) effective drug targeting, including the ability to reach and access the targetzone; (3)

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stability on route from the site of administration to the site(s) of drug action; and (4) the ability to perform as a sustained-release drug depot. In order to meet the attributes make that upcategory 11, the carrier should be biocompatible, biodegradable, nonimmunogenic, and nontoxic. A major share of the qualities that are dominant in category I11 have to dowith the production of the drug-carrier systems. The carrier should be of the type that its preparation prccedures(including drug loading) can be scaled up, yield a sterile and pyrogen-free product, and meet the requirements of quality assurance. Another major part of category I11 properties concerns postproduction requirements, such as dosage forms that can provide longterm shelf life and high stability. Needless to say, in order to ensure that as wide as possible patient population could have access to this new therapy, all cost-effective issues should be considered within category 111, be they the cost of production itself, sources and cost of raw materials, shelf life length, expenses of storage, and others. Among drug-delivery systems that can address a major share of the unmet needs encountered when treatments arewith free drug,liposomes are frontline candidates. Their advantages for the task have been well documented and include the following: Liposomes can meet that required mutual drughiological environment protection. Liposomes are biocompatible and biodegradable and, to a significant extent, are also nonimmunogenic and nontoxic. Their versatility in terms of type, size, and lipid compositions allows the encapsulation and delivery of a wide range of drug species, and liposomes can act as sustained-release depots. Despite all these qualities, significant problems in the implementation of therapies with liposomes still exist, particularly with respect to targeting; theresponses of the biological system to liposomes in the blood stream; the stability of both particle and drug in vivo;the in vivo rates of drug release; shifts in rather than prevention of drug toxicity; and long-range effects of chronic liposome administration. The advantages of liposomes as well as these unresolved issues are addressed in the following sections, taking into consideration not only these category I and I1 aspects but also those of category 111that arecritical for the evolution of liposomes from entities underresearch and development into entities that are pharmaceutical products. SELECTION OF THE LIPOSOME TYPUSPECIES: VIEWS AND CRITERIAFROM ACADEMIC AND INDUSTRIAL RESEARCH AND THEIR PROPOSED INTEGRATION 111.

A. Liposome Type, Size, and Production Issues

One of the initial decisions that need to be made on venturing into the development of a liposomal drug-delivery system for a specific therapeutic

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case centers on the type or types of liposomes that will be explored. Whetherthat selection will bemadefrom the abundance of existing liposome species or theinvention of new liposome species is contemplated, the criteria and considerations that are used to make that decision constitute oneof the majorpoints of diversion between investigators in academic and industrial research. The selection of liposome types in industry will follow, in general, guidelines for pharmaceutical products, and would be relatively free from the limitations of accessibility to devices for liposome production. The main concerns would be production and product issues. As discussed under category I11 (see Section II.B), a critical criterion would be a liposome production method/procedure thatcould be scaled up and executed under GMP rules and regulations. Other critical criteria would stem from the requirements for a product that would have batch-to-batch reproducibility andbestable, sterile, and pyrogen free. As also discussed above, the liposome production method had to be cost effective, also taking into consideration sources, supplies, and cost of quality raw materials. Cost and environmental considerations would also take into account the issue of waste disposal, in particular for those methods of liposome productionthat involve the use of organic solvents. Needless to say, the selection of the liposome type would also take into consideration the need for acceptable shelf life and dosageforms. The academician is perceived to have a considerably higher level of freedom, being able to choose, in principle, from all types of liposomes for which procedures of preparation are in existence. Physiological and clinical aspects the drive selection criteria. These include the specific therapy in question, drug properties, the route of liposome administration with all its inherent issues, and the mutual effects of the drug and the biological system. However, as exemplified by the following case, other criteria often become deciding factors. Liposomes as delivery systems have become common andprevalent enough that the situation in which an investigator with expertise and a primary interest in using a particular drug-for example, a 6000-d polypeptide-seeks liposomes asa delivery system for it. Screening the “horn of plenty” with respect to liposome types and species, certain dogmas in the field might drive that investigator to exclude from consideration MLV, although these liposomes might well fit his therapeutic objectives and the other requirements. Concerns of stability and retention of the native active conformation could lead that investigator to also exclude ULV made by methods that bring the encapsulated polypeptide in contact with organic solvents or detergents. If costly devices for making ULV of sufficiently large size range are unavailable because of financial limitations, the investigator might end up selecting small ULV (20 nm in diameter) made with a

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probe sonicator. This selection can posesevere limitationson theefficiency of encapsulation and on other properties of the system that can result in low efficacy of treatment that would discourage the investigator fromany further pursuit of the liposomal approach. Even if those small ULV would prove to be satisfactory with respect to physicochemical properties and successfully functional in vitro and/orin vivo, the questionof whether they could be developed into apharmaceutical product would stillbe faced. The same situationwould have to be faced withnumerous otherliposome types that are currentlyused in academic research, such asthose madeby reverse phase evaporation, ethanol injection, detergent dialysis, French press, detergent removal through gel exclusion chromotography, and others [2,3,57,12-14,18-191. It is stressed that for objectives other than drug delivery, especially those where liposomes serve as modelsof cell membranes or as a solubilization media for lipophylic matter, most of the limitations discussed in this section are of no consequence. However, cases in which efforts and resources have been invested in developing liposomal drug systems that prove to be successful in animal model studies and even in clinical trials, with liposome types that cannot be produced as pharmaceutical products, are not far fetched. For such cases, the progress of that system into an established treatment modality would be (at best) significantly delayed until similar positive results would be obtained with another, productsuitable liposome type. Needless to say, a major shareof the already done studies would have to be repeated. The clear conclusion that emerges from the issues discussed in this section is that, in the selection of liposome type, the burden of change in current considerations is mostly on the shoulder of the academician. It is proposed that the academician integrate the industrial criteria into his or her initial selection process, even if the requirements of a planned study could be accomplished withthe use of small-scale, nonsterile, preparations that would be discarded at the end of a day’s work. A few examples of implementing the integrated view are offered below. A comprehensive list of selected liposome types is deemed to be beyond the scope of this chapter. Thisis due not only to the abundanceof liposome production methods but also to the realization (already introduced in the previous section) that this is a dynamic field where new species and/or production methodologies are expected tobe continuously invented. The first example concerns MLV,* that have fallen out of favor with

*Other propertiesthat might favor MLV will be discussed in subsequent sectionsof this chapter, aswill be the already mentioneddrawback due to thesue heterogeneity and others issuesin this vein.

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many investigators. Granted, there definitely are treatments and applications for which MLV wouldnot do. For treatmentsof tumors andinfectious diseases in which the targets are outside the RES and where intravenous administration with long-term retention of the liposomes in circulation is required. On the other hand, MLV can be eminently suitable for some therapies that require nonsystemic administration as well as for therapies that requireintravenous administration and fast uptake of the liposomes by phagocytic cells in circulation. MLV can be easily made in the laboratory and canalso meet thecriteria for pharmaceutical products. The latteris not meant to imply that all industrial issues with respect to MLV production have beenresolved, but thattheir resolution is feasible. A pertinent example of a current problem that needs resolution is the already mentioned issue of organic solvents, such as methanol,chloroform, dichloromethane, Freon, and similar ones, that are routinely used in MLV production. The use of such organic solvents is currently critical for proper MLV production. They are applied for the dissolution and optimal mixing of the different lipids that make up the formulation and of those drugs that are introduced through thelipid phase. They also make a major contribution to the attainment of the form anddispersion of dry lipids for optimal contact with the swelling solution. On the other hand, the specific solvents used present a problem ontheir own, both in the industrial and the academic laboratory level, as regulatory authorities phase more and morepossible solvents out of use. Owing to the environmental and economic issues involved, waste disposal of these solvents, which could accumulate to substantial quantities in industrial-scale MLV production, is another cause for concern. Clearly, if MLV or any of the many types of ULV.for which MLV are the source material are to be used, other meanswould be required to still produce the MLV, but with an altogether elimination of the need for organic solvents or their replacementby lesshazardous materials that could be used with much lower quantities. Based on the issues discussed above, we propose that MLV should berevisited. The second example is the case in which, based on the therapeutic objectives, MLV are ruled out and it has been determined thatthe optimal ULV should have a dominant size of 200 nm diameter. The many ways by which such liposomesare produced can bedivided into twocategories: The first is device oriented, using as source material MLV that are subject to extrusion, homogenization, or microemulsification.devices in order to transform them into thedesired ULV. The second category includes methods such as reverse phase evaporation, ethanol injection, detergent dialysis, detergent removal through gel exclusion chromotography, and others (2,3,5-7,12-'14,18-19). In this category, the start materials are usually multicomponent systems that include lipids, drug, water, and an organic

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solvent or detergent.Despite the relatively ease and distinct lower cost of producing liposomes by methods of the second category, even assuming the contact between drug and organic solvent or detergent is not a limiting factor, it is argued here that the first category is the better choice. The ability to scale-up procedures of the second category is questionable and,at best, might need investment for itsdevelopment while scaled-up models of the first category devices are already in existence. The issue of organic solvents and waste has already been discussed above for MLV, but unlike the case of MLV, it is questionable if the critical roles of the organic solvents/detergents in methods of the second category can be eliminated or even reduced in quantity (especially relative to the other components of the system), The recommendation here for the selection of ULV made by the device-oriented methodsis not meant to ignore the cost limitations. Rather it is argued that ULV of a specified size range that are producedby either type device should be sufficiently similar, especially as all use the same source material (i.e., MLV). With the right choice, this should make it possible to perform thebasic stages of a study with one device and proceed to theindustrial R&Dlevels withanother without significant changes in the liposomal properties. For example, it should be possible for the academician to use small relatively inexpensive extrusion devices (as well as design and build one) and use for the industrial stage amicroemulsifier operated under conditions that would give the same size liposomes under similar temperature, buffer, and other experimental parameters.

B. Lipid Composition The source and cost of lipids have been concerns for liposomologists in both academic and industrial research.For the major run-of-the-mill lipids usedinmost liposome formulations, such as phosphatidyl choline, the increase in available sources has somewhat eased those concerns. This trend can be expected to continueand grow as more liposomes will successfully complete clinical trials and move into the arena of products, making the supply of high-purity lipids a viable business venture. The situation is different when it comes to the more rare lipids. The motivation for incorporating relatively rare lipids into a liposome formulation comes from the need to endow the liposomes with unique properties that will take into account the natureof the drug in question and address specific demands of the intended therapy. The perceived freedom of the academician in the selection of liposome types might also be extended into selection of lipid composition, but in reality, the paucity of source materials for uncommon lipids together with their high cost mightnegate theiruse at the pharmaceutical product stage. Thus,even at the basic stage, a balancewould have to

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be struck between the liposome formulation and the properties to be endowed by specific lipids.In general, the integrated approach would dictate giving up that singular lipid and finding alternative means to endow the it should be liposomes with the samedesired attribute. On the other hand, borne in mind and taken into consideration that basic studies might offer unanticipated solutions to particularly difficult pathological situations that would justify even exceptionally high costs. The evolution within the Stealth (a registered trade name of Liposome Technology, Inc., Menlo Park, CA)liposomes is an instructive example. The unique feature of these liposomes is their ability to delay/evade uptake by the reticuloendothelial system (RES) for longer periods than regular liposomes of similar size range. This allows for prolonged retention in circulation and through that higher shares of the dose can reach the non-RES targets. The first generation of Stealth liposomes required a specific ganglioside in their formulation [51]. Subsequent generations of Stealth liposomes have seen themove from that lipid to the relatively less rare regular and hydrogenated phosphatidyl inositol and specific hydrogenated phosphatidyl choline-cholesterol mixtures [52], ending up in formulations that do not require rareand/or modified lipids at all. In the current version of these liposomes, the “stealth” property is achieved through surface modification by polyethylene glycols [53]. The extent to which elimination of the dependence on rare lipids has been a driving force in this evolution has not been specified byits developers. Yet, even if considerations of lipid raw materials were not a major driving force for this particular evolution, this case shows that such considerations can be accommodated without giving up on desirable formulation-dependent liposomal features.

IV. TARGETED/MODIFIEDLIPOSOMES:ANINTERESTING AND EXCITING SCIENTIFIC TOOL, BUT CAN THEY BE MADE INTO PRODUCTS (ESPECIALLY IMMUNOLIPOSOMES)? The ability to be targeted to the sites of drug action, and to them alone, is among the critical properties required of liposomes. It is well known that despite enormous investigative efforts, the achievement of targeting that would be highly effective in vivoand applicable to a wide range of therapeutic objectives is still an elusive goal. As will be discussed below, complete targeting has not been achieved even in some of those cases defined as “passive targeting,” where the targets are within the RES [54]. Despite the frequent appearanceof the term targeting in liposome research and literature, its very prolific use has somewhat blurred it into meaning different

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things to different people. Owing to this current state of affairs, it was deemed advantageous to offer here a brief discussion and definition of targeting prior to a discussion of the means by which liposome targeting is attempted, and their evaluation from both the academic and industrial points of view.

A. Targeting: Definition, Stages, and Status It is offered here that liposome (or any other drug-delivery system) targeting can beviewed as a two-tier process, the first of which takes place on the introduction of liposomes into a living system, and thus is termed here “targeting at theorgan level.” The objectives and efforts within this tier are focused on getting the drug-loaded liposomes from site(s) the of administration to the orgadtissuehicinity where the state(s) of disease reside. The second tier, termed “targeting at the cellular level,” is a form of “fine tuning.” It centers on pinpointing the drug-loaded liposomes, on their arrival at thatlocation, as close as possible to theactual sites of drug action. It should be emphasized that in order toavoid interference with the therapeutic activity, the liposomes might need to be placed close to-but not at-the sites of drug action. Thus, targeting at thecellular level can involve two distinct types of sites: sites at which the liposomesreside, which willbe termed “camer sites,” and those at which the drugbinds in order to initiate its therapeutic effect, which will be termed “drugsites.’’ The targeting capabilities demanded of liposomes for therapies that require systemic administration depend to a large extent on the locations of the sites of drug action. For ultimate targeting to locations outside the RES, the liposomes need capabilities to address the needs of both targeting tiers. For targeting to locations within the RES, the liposomes need to provide the second tier alone, as thefirst one is provided forby the biological system itself. For therapies thatdonot require or are preferably achieved by nonsystemic drug administration, the situation is somewhat similar to RES targets; that is, the liposomes haveto provide only for the second tier of targeting. In contrast to theprevious case, in the nonsystemic therapies, the first tier is achieved (or one could say eliminated) by the mere selection of the route of administration. Included in such therapies would be topical drug administration in the treatment of wounds, bums, and ocular conditions and regional drug administrations such as intraperitoneal infusions and aerosol inhalations for treatment of disorders in the peritoneal cavity and in the lungs, respectively [55-581. Liposomal specifications for the achievement of the first tier of targeting in systemic applications for non-REStargets have been comprehensively and extensively reviewed together with the inherentphysiological and ana-

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tomical obstacles [2-12,51-54, and references therein]. In an effort to refrain from redundancy, the reader is referred to these referenced sources and the discussion below is focused on the less-addressed specifications for achievement of the second tier. The basic premise is that once the liposome arrives at, oris introduced into, theorgadtissueAocation where the state(s) of disease reside, effective targeting at the cellular level requires that the liposome adhere with high affinity to its designated carrier sites and be retained there, despite cellular and fluid dynamics,for a sufficient duration in order torelease an effective drug dosefor the time spandictated by the specifics ofdrug anddisease. Ideally, the liposome should also be instrumental in aiding drug access to sites of drug action. Tailoring liposomes to this end should be done with care andis rational for those cases where detailed knowledge of the cellular or tissue location of the drug site is available. Obviously, for intracellular sites of drug action, liposome internalization would be beneficial, but it is not an absolute necessity. For sites of drug action that areextracellular, such as certain membrane-embedded receptors or matrix-residing bacterial colonies, liposome internalization would defeat the purpose. B. The Means Through Which Liposome Targeting Has Been Attempted and Their Feasibility for Liposomes as Pharmaceutical Products

The main efforts for endowing liposomes withtargeting abilities have been focused on liposomal features. Other avenues that explore treatment regimens such as liposome dose size and frequency of administration or pretreatment with “blank” liposomes have enjoyed less attention. With respect to liposomal features, the two main avenues explored were liposome specifications such as size, type, and lipid composition and liposome surface modification. The attempts to confer a targeting ability on liposomes through manipulation of their specifications, provided they do notstray from the integrated criteria for selection of liposome types discussed in the previous section, will continue to result in liposomes that can be developed into pharmaceutical products. Some success has been found in this avenue of exploration, as shown recently by preferential accumulation of small daunomycin-encapsulatingliposomes in solid tumors, althoughthe operating mechanism has yet been elucidated. [59]. It has been recently shown that small stealth liposomes accumulate, apparently by entrapment, in the intracellular spaces within solid tumors [60]and are retained within those spaces for a sufficient duration to allow drug to diffuse from the liposomes .into the tumor cells.

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The attempts to confer targeting ability through liposome surface modification are an entirely different matter. Antibodies are, by far, the most extensively investigated and attempted class of targeting agents that have been attached to the liposomal surface [2,3,7-8,12,61-641 for both tiers of targeting. A comprehensive discussion of these liposomes, for which the termimmunoliposomes has been universally accepted, is beyond the scope of this chapter and the reader is referred elsewhere [e.g., see refs. 1-12,51-54-55, and those cited therein]. The discussion here will touch briefly onthestate in the fieldwith respect to targeting with immunoliposomes andtheir feasibility as pharmaceutical products. A fair share of the studies pursuing targeting with immunoliposomes have been conductedin cellcultures. Taken as studies aimed at thefirst tier of targeting, cell cultures are not the best choice for a model system, especially for systemic applications. Taken as studies aimed at the second tier of targeting, cell cultures are quite suitable models even if the goal of the investigators was to test the first tier rather than the second. Animal model studies have shown, to date, that despite being such an elegant approach, successfulinvivo targeting withsystematically administered immunoliposomesis quite limited and still far from beingthe overall solution to liposomal targeting. Moreover, the question of whethere it might ever be a wide-range solution is still open. On the other hand, in vitro studies with immunoliposomes, which have shown specificity in binding and/or improved a biological response that was restricted to cells carrying the appropriate antigen, clearly demonstrate that it should be possible to achieve the second tier of targeting with this approach. Obviously, this speaks fora realistic potential of immunoliposomesin nonsystemic applications where, as already discussed, the first tier of targeting is achieved (or the need for it eliminated) by the selection of the administration route. Theoretically, targeting of intravenously administered immunoliposomes could also become possible for a wide range of applications if the means could be found to endow the immunoliposomeswith the ability to provide the first, without compromising their ability to provide the second, tier of targeting. Because they have a future with at least some targeting applications, the current situation with immunoliposomes makes the question of their feasibility as a pharmaceutical productrelevant and valid. This is a rather complex question comprising issues, to be discussed below, that concern the antibodies themselves as well as the antibody-liposome systems. It is stressed that thediscussion isfocused only on theissues that are related to the question at hand;namely, the feasibility of immunoliposomes aspharmaceutical products. Physiological and clinical advantages and drawbacks of immunoliposomes, are not addressed here.

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Obviously, to be a pharmaceutical product, immunoliposomes are required to meet all criteria that bear on antibody production, stability, retention of biological activity (for targeting, at least with respect to the antigen-antibody interaction), and appropriate shelf life. Another major factor in the making of an antibody into a pharmaceutical product (or part of) is the combination of specificity and antigen abundance. The antibody has to be specific enough for its designated use (whether as a targeting or as a therapeutic agent) yet the antigen shouldbe general andabundant enough so that theproduct would be applicable to a wide enough range of the patientpopulation. Viewed as futurepharmaceutical products, the process of modifying a liposome into an immunoliposome is wrought with obstacles that start from the need to meet industrial criteria of the type previously defined for the selection of the liposome type and thelipid composition and continueinto the modification process. The most frequently reported surface modification approaches employing agents such as SPDP and similar ones [64-651 are multistep processes. First, the antibody and a lipid undergo, eachseparately, chemical modification and purification procedures, including masking of active residues for molecular protection. Next come liposome formation (incorporating that modified lipid into theformulation) and the unmasking of an active residue on the antibody, after which the antibody-liposome association process takes place. The process is completed by further purification steps. Performed ona a large scale, as would be neededfor a product, it is easy to see thatthis could turn into a monumental endeavor. Suffice to consider the many steps along the way where significant liposome, drug, and antibody losses might occur and the issue of sterility in the final product. Considerations of raw materials, waste disposal, stability, retention of encapsulated drug, and drug stability would all add to thedifficulty, as wouldthe need for an overall cost-effective process. On this basis, it seems clear that although immunoliposomeswill continue to bea useful research tool, their development into pharmaceutical products, even for cases where they can provide targeting, will have to be decided case by case, and taking into account not only the obstacles listed above, but also those that couldarise from physiological and clinical aspects. Other types of liposome surface modifications for the purpose targeting that are notbased on theimmune system might be more feasible as future pharmaceuticalproducts. The case of polyethylene glycolated stealth liposomes for systemic administrations has already been discussed. Other potential opportunities are bioadhesive liposomes, which have beendeveloped in recent years for nonsystemictopical and regional applications [56581. These arebased on theuse of ubiquitous surface-bound agents such as collagen, hyaluronic acid and growth factors, and on surface modification procedures thatare less complexthan those discussed above forantibodies.

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With respect to targeting, the first concern of an investigator developing a drug-liposomesystem for a specific therapeutic objective is to find the means to achieve targeting. Together with this concern, it is proposed that the investigator should weigh his or her options along the lines discussed above. If the therapy in question requires addressing both tiers, then, unless the investigator intends to embark on the development of new targeting agents, at present he or she might be better off attempting targeting with nonmodified liposomes, exploring liposome specifications. If the therapy requires the second tier alone, targeting opportunities are offered by surface-modified liposomes, with current ornewly developed agents. In all cases, regardless of whether targeting is pursued with current or new surface-modifying agents, the feasibility for future pharmaceutical products, as exemplified above for immunoliposomes, should be taken into account. In closing this section on targeting, two additional points are offered. It is stressed that thediscussion above should not be taken asa proposal to discount immunoliposomestotally. Rather it is meant to propose extreme caution in the pursuit of antibodies as liposomal targeting agents. Recent studies with immunoliposomescarrying an antimyosin antibody are a pertinent example [66].These liposomes have been shown to. be capable of targeting in vivo, and this antibody meetsthe combined criteria of specificity and antigen abundance for sufficiently large patient populations, as discussed above. For such a case, overcoming other hurdles with respect to immunoliposomes as pharmaceutical products, could be well worth the efforts. Finally, despite “the heat of the battle,” it is imperative not to lose sight of the critical objective of targeting: It is the drug which should be targeted, with the liposome a means to that end, even though it is often experimentally wiser to first pursue targeting with drug-free liposomes. Even if successful targeting of a given liposome system has been achieved, and theresultant liposomes can meet the criteria of becoming pharmaceutical products, the work is not yet done. Completion requires experimental verification that those liposomes canreach the target while still carrying a drug load that is sufficient for effective therapy. V. LIPOSOMES AS A STERILE, PYROGEN-FREE SYSTEM WITH PHARMACEUTICALLY ACCEPTABLE SHELF LIFE, STABILITY, AND DOSAGE FORMS A.Sterile,Pyrogen-FreeLiposomes

In basic research, an investigator can forego the need for sterile, pyrogenfree liposomes provided the experiments conducted are &ch that the lipo-

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somes do not come into contact with living matter at all or that such contacts, bethey in vitroor in vivo,are of short duration. For a pharmaceutical product, such qualities are critical. The steps thatshould be takenin order to obtain pyrogen-free liposomes are not essentially different from those implemented for other pharmaceutical products [5,67].Securing the sterility of liposomal products isan entirely different matterand has the makings of a major obstacle [67].The major approaches in use, such asheat sterilization and y-irradiation, are not suitable the for end product [5,67].The two feasible approaches aresterilization by filtration and an aseptic production process. Of the two,the lattercan potentially be applied to all species and types of liposomes, regular aswell assurface modified. The sterile fill is restricted, obviously, to liposomes small enough to pass the selected pore size of the filter, usually 0.2 pm, provided obstructions such as filter clogging and filter adsorption (especially for surface-modified liposomes) do not turn into insurmountable complications or lead to significant losses inmatter. Consequently, the issues surrounding the production of sterile liposomes, by being of mutual interest and need to liposome investigators in both academic and industrial research, constitute an area where cooperation will be neededin order to arrive atacceptable solutions.

B. Shelf Life, Stability, and Dosage Forms It is well known that the formation of liposomes from dry lipids suspended in an aqueous media is driven by thermodynamic stability [1,68].Hence, to survive and function as liposomes, these particles need to be surrounded by water, particularly at the start and end of the road; namely, at the endof a liposome production process and when called to action within the biological milieu. During thetime interval between production and in vivoperformance, the liposomes have to be stored for an acceptable period that, for pharmaceutical products, usually extends beyond 1 year. For refrigerated (but not frozen) aqueous suspensions and lyophilized powders, defining the majorrisks of long-term storage and reviewing, from the point of view of the academician, the advantages and drawbacks of each form of storage and for administration, are thetopics of this section.

1. Shelf Life and Long-Term Stability Breakdown in sterility, chemical destablization, formation of undesirable degradants and loss of therapeutic activity are among the majorrisks run in long-term storage of any pharmaceutical entity. When it comes to liposomes, guarding against and prevention of such risks are inherently more difficult. More than one chemical entity is involved: lipids, drugs, and for modified liposomes, the surface-anchored agents. It also involves the pres-

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ervation of particle integrity and the retention of the encapsulated drug within the liposome. The major advantages of liposomal storage in the form of aqueous suspensions are (1) the retention of the original type and specifications of the liposomes selected and produced for the designated therapeutic task and (2) their “ready to use” mode. The former is of particular importance when unilamellar liposomes are concerned, especially when substantial efforts have been invested in their production and where the effective therapy is dependent on a specific size and narrow size distribution of the liposomal preparation. A critical component of the ready to use mode involves the distribution between encapsulated and unencapsulated drug. Obviously, effective therapy in which the liposomeswill make a substantial difference compared with free drug requires that a sufficiently highshare of the drug in the administered dose be encapsulated. This is particularly important fortoxic drugs (such as chemotherapicagents) where protection of the biological environment from the drug is a major motivation for liposomal delivery. In such cases, even a relatively small share of unencapsulated drug could defeat that objective. The considerations outlined above favor the storage of liposome preparations that have been cleaned from unencapsulateddrugs together with the retention of this state throughout the storage. Removal of unencapsulated drug, which constitutes a displacement of the equilibrium distribution of thedrugbetweenthe liposomes and the external aqueous phase, will inevitably create anelectrochemical gradient, driving the drug outof the liposomes [56-58,691. As will be discussed in greater detail in a following section, the rate of this diffusion will depend on several factors, with drug properties being foremost among them. Evaluation of the release of kinetics of the drugin question and of the effects of already recognized tools for implementationof satisfactory storage conditions, as well as the development of new tools for the task, start at the basic research level. The risks of loss of encapsulated drug onlong-term storage of liposomes thathave undergone separation from unencapsulated drug and are therefore in a nonequilibrium state are particularly high for small molecular weight drugs stored under those nonequilibrium conditions. For suchcases, storage at high lipidconcentrations [69], the introduction of additional intraliposomal barriers such as theammonium sulfate gel [67], and utilization of remote-loading procedures [70] could be used to reduce therisk of that loss. For othercases, even thoughit might take away from the ready to use advantage, the only recourse would be to store the preparation underequilibrium conditions (i.e., without the removal of the unencapsulated share) anduse it as it is or institute a separation procedure immediately prior to administration.

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Besides beingeconomically more attractive, storage of liposome-drug systemsin dry form, as freeze-dried powders, seems the better choice, since it leads to significant reduction in the risks discussed above. On the other hand, excepting selected situations (to be discussed in the next section), the dry powder is not the readyto use form, and liposomal reconstitution through rehydration would be required prior to use. Needless to say, reconstitution has to be performed under sterile and pyrogen-free conditions. The main concerns and drawbackswith this dosage form center on the nature of the liposomal species and on thesituation of drug encapsulation that are obtained on reconstitution. Studies aimed at theseissues have shown that unless specific steps-mainly the introduction of cryoprotectants into theliposomal formulation-are taken prior to lyophilization, the reconstituted systemswill revert to MLV, independent of the original liposome species [71-731. Although the feasibility of this approach has been proven and these exipients are considered safe, potential adverse effects of those cryoprotectants on the properties of the drug in question and (where relevant) on the surface-boundagent cannot be ignored. Independent of liposome species, less is known about the fateof drug distribution in the reconstituted system between the liposome and the (new) aqueous medium. If the share of unencapsulated drug is beyond the level of acceptance for the system in question, this will require correction through actions such as revisions in conditions of lyophilization and/or reconstitution and postreconstitution purification steps. For those liposomes thatare surface modified, the distribution of that agent should be also considered. If big enough, steric hindrance might favor its relocation on reconstitution to the liposomal surface, but the situation could arise where the share of reconstituted liposomes having that agent inside is not negligible. This could result in problems that range from the mere loss in the fraction of targeted liposomes to thehazards of liposomes carrying toxic drugs circulating indiscriminately in the body. The characteristics of the reconstituted liposome, including preservation of original structures through the use of cryoprotectants, the possible adverse effects due tothose additives, and the distributions of the drug and the surface-bound agent in the reconstituted systems, have been explored [71-731. However, it is argued here that whether such investigations have been conducted with drug models or with specific drugs, they need to be done anew for eachspecific liposome-drug system investigated and developed for a particular therapeutic objective. Undoubtedly, such questions have strong components that put them within the domainof basic research. 2.

Dosage Forms for Administration Suspended in an aqueous suspension form, regardless of the original storage form, the liposomes are ready for application utilizing not only the

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intravenous route but other routes of administration as well. They couldbe applied topically to the eyes, to wounds, and to burns. They could be injected subcutaneously or intramuscularly or introduced into the peritoneal cavity through injection or infusion. Other routes of administration requirefurther processing of the aqueousliposome suspension, which touch on basic and industrial issues that havenot been addressedyet in this chapter. A most prominent case isprocessing into aerosols of dru'gencapsulating liposomes (regular and surface modified) that would be useful for pulmonary applications. The making of liposomal aerosols for drug delivery opens new questions such as theeffects of the aerosolization process (where relevant) aerosolization components on (1) all aspects of stability (drug, lipids, surfacebound agents), (2) the retention of encapsulated drug, (3) the retention of (liposome) particle integrity and size, and (4) the retention of therapeutic activities, without which success inthe retention of the other propertiesis immaterial. Liposome aerosols have been prepared andcharacterized with respect to some of these properties [74-821, and theresults attest to both the feasibility of this dosage form and to thestrong dependence of its success on the drugand the liposome species explored. Consequently, the data that has accumulated using drug models will not save an investigator developing a drug-liposomesystem aerosol form fromthe need to invest in basic research of the stability and retention issues listedabove. The physical features of the aerosol and their optimization with respect to the designated therapeutic goal would also be required, utilizing cascade impingers and similar devices [74,77-781, as well as a means for aerosol collection and analysis. VI. LIPOSOME CHARACTERISTICS (PERCENTAGE OF ENCAPSULATION, KINETICSOF RELEASE, BIOLOGICAL ACTIVITY)-IN BASIC RESEARCH AND IN QUALITY ASSURANCE

In this chapter, much has already been said about basic liposomal traits such asthe efficiency of encapsulation, kinetics of drug release, and retention of biological activity.The investigation of these traits is inthe realm of basic research, and among other objectives is intended to serve in system optimization and in the development of in vitro models for pursuit of the liposomal invivo fate. Once these encapsulation, release, and activity specifications are set for a given liposome-drug system, they can be used within the industrial environment in the evaluation of the retentionof the liposome properties in the scaled-up production and as part of the battery of criteria for quality assurance. In consideration of the common ground these propertiesconstitute for investigators from both academia and industry, several comments andproposals are offered below.

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Efficiency of Drug Encapsulation

Throughout the three decades of liposome research, several definitions have been introduced as measures for this property. For example, the fraction of encapsulated drug from thetotal in the system, the ratioof drug to lipid, and the volume (pergiven lipid quantity) of the internal aqueous phase. For moleculesserving as modelsof encapsulated matter, such asCF (carboxyfluorescein) and similar derivatives, elegant procedures have been devised in order todetermine the efficiency of encapsulation [2,6,12]. One of their most favorable features is the elimination of the need to separate the drug-encapsulating liposomes from the media containing the unencapsulated drug. For most real drugs, irrespective of the efficiency parameter pursued, the experimental process for determination of encapsulated matter will require such a separation step. Although simple in concept, in continuatibn of the discussion inSection V.B.l abovewith respect to cleansing from unencapsulated drug, most methodologies contain steps that can lead to overestimation or underestimation of the quantity of liposomeencapsulated drug, especially for small molecular weight drugs. Separations that makeuse of semipermeable matrices such as exhaustive dialysis, various filtration devices, or gel-exclusion column chromatograpies all run the risk of dilution of the external medium in the course of separation. This creates a driving force for efflux of the encapsulated matter, which would be most pronounced for small molecular weight drugs. The encapsulation level determined at the end of such proceduresmight be an underestimation of the initial level, with the deviation depending not only on drug propertiesbut also on thespecific experimental conditions used for the separation. This could becomea problem notonly withrespect to accurate evaluation of encapsulation efficiencies. Batch-to-batch reproducibility and implementation of encapsulation efficiency as a quality assurance criterion would also be adversely affected, unless extensive care is taken in duplicating the separation process from batch-to-batch. Separation by centrifugation is supposedly free from the risks of external medium dilution. However, if a single cycle is used, there could be overestimation of the encapsulated fraction due toadsorption of unencapsulated drugto theliposomal pellet. Additional cycles of washing drug-free in buffer could reduce that error, but they need to be used withcaution in order toavoid the risk of dilution, which can lead (as discussed above) to underestimation. How then can thelevel of encapsulation be accurately assessed without the interferences that occur as a result of the separation steps? One. approach exemplified by the aforementioned CF is to develop assays that can distinguish (quantitatively) between encapsulated and unencapsulated matter in the same system. Such assays would be strongly dependent on

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drug properties and probably would be useful for a limited number of cases. Another possibility, to be discussed in the following section, is the determination of encapsulation efficiencies through studiesof drug release. B. Kinetics of Drug Release

Irrespective of the level of organization at which they are performed, studies of drug release/diffusion from liposomal systems are directed toward issues that are relevant to the in vivo as well as to the non-in vivo arenas. For liposomes in the in vivo arena, the drug-release studies are expected to yield data and understanding that will lead to (1) minimizing the loss of encapsulated drug on route from the site of administration to the site of drug action and (2) the ability to match the rate of release (once the liposomes arrive at the target) to the requirements of the therapy. The objectives of drug release studies that concern the non-in vivo arena are (1) physicochemical characterization of the systems, including liposomes processed into aerosols or reconstituted from freeze-dried powders; (2) various aspects of system optimization such as the selection of liposome type, lipid composition, and parameters of shelf life; and (3) criteria for quality assurance. In order to derive relevant data from such studies, the experimental conditions should be set to fit the specific objectives, especially with respect to the extent of liposomes and drug (each, separately) dilutions that the system is anticipated to undergo. Detailed discussed of the in vivo objectives are outside the theme of this chapter, anddiscussion of the impact of such dilutions has been discussed elsewhere [55]. The nonin vivo objectives are within the subject of the present discussion and will be addressed below. Drug release from intact liposomes into the media within which the liposomes are placed (or suspended)is essentially a process of diffusion in a heterogeneous system; the latter madeof several phases of water andlipid bilayers, the number of which willdepend on the liposome type andon drug solubility properties. At thevery least, two phases are involved: a lipophylic drug embedded within the lipid bilayer of a unilamellar liposome. In principle, initiationof drug diffusion simplyrequires setting the driving force (i.e., an electrochemical gradient of the drug from its liposomal location to the external medium) afterwhich the process will proceed until equilibriumis established (or reestablished). When the starting material is a drug-liposome system at equilibrium with respect to drug distribution, reduction in the drug concentration at the external medium suffices to trigger the releaseprocess. Several means can be used togeneratethatreduction:precipitation, enzyme-catalyzed degradation, and dilution of the liposomal system into the medium of choice such asbuffer or body fluid(real or simulated). The latter

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is, by far, theeasiest and most general means to generatesuch a drug concentration reduction that is suitable for a wide range of the objectives listed above. If, as discussed above with respect to encapsulation efficiencies, the liposomal system has been previously subjected to “cleaning procedures” in order to rid it of unencapsulated drug, the electrochemical gradient is already in existence and the diffusion process already operative prior to the specific kinetic experiment. Independent of the state of the system on initiation of a particular experiment (i.e., equilibrium or nonequilibrium of drug distribution), it is proposedthat sustaining a unidirectional fluxof the drugfrom the liposomal system into the bulk medium during the entire experiment is a key element. Data from experiments done under such conditions represent the worst-case scenario for drug loss from the liposomes. This knowledge can then be used to design shelf life conditions under which drug loss is minimized [56-58,691. Data of this type can also be used to set up,in the process of liposome surface modification, the conditions that will prevent depletion of the encapsulated drug. Once the diffusion has been initiated (or is already in progress), the experiment itself consists of quantitative evaluation of drug accumulation in the medium and/or drug loss from the liposomes at designated periods. Critical to thesuccessful execution of such studies is the quantitative separation of liposomes and theexternal medium in aliquots withdrawn fromthe reaction mixture at designated time points. Classic means forthe quantitative separations of particles from their suspension medium, such as centrifugation and filtration, are often compromised by drug loss to the filtration matrix or by centrifugation procedures that aretoo slow withrespect to the time that has elapsed between samplings. Additional experimental constraints might be encountered in drug and liposome assays. The dilution which is essential for the onset and maintenanceof the gradient can result in lipid and druglevels in the separated fractions of the withdrawn aliquots that fall below the limits of detection and/or quantitation. Elimination altogether of the needfor separation together with significant reduction of the risks of falling belowdetection limits is offered by the use of appropriate dialysis set-ups [56-58,691. In this approach, a liposome preparation is enclosed within a dialysis sac that is immersed in a drug-free medium. The wide variety among available semipermeable dialysis membranes makesit possible to select, according to specifications of the investigated system, a membrane that will not adsorb the drug and will not be a barrier to the drug but at the same time be a complete barrier to the liposomes. Analysis of the raw data, rather than remaining at the phenomenological level, can not only give insights into the mechanism(s) but can also

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aid in identifying factors that are instrumental in gaining control of the release. Among the theoretical frameworks available for the analysis of diffusion data, two properties make the theoretical approach developed by Eyring particularly suitable for drug diffusion from liposomes. It allows drug release from homogeneous (unencapsulated drug) and heterogeneous (liposome-associated and liposome-encapsulated drug) systems to be dealt with simultaneously. It yields parameters that allow the direct determination of the fraction of encapsulated drug and of the half-life of drug release. In the long run, such parameters are especially useful for systems that are destined to serve as therapeutic entities, in defining optimization criteria, andfor designing dose ranges and treatmentregimens. Studying the kinetics of drug release from liposomal (regular and surface-modified) systems, using theexperimentaland data processing approaches discussed above, the authors were able to sortseveral underlying principles that are general to these release processes [56-58,691: It was found that the release kinetics can be described as a series of parallel first-order processes, each representing a drugpool that exists in the system at time = 0. One pool represents the drug that is unencapsulated and all others drug thatis liposome associated. The mathematical expression for this type of mechanism is given by [see refer. 69 for additional details]:

f=

2 &
e+>

(1)

j= 1

where t represents time, the experimental independent parameter, and f represents the dependent experimental parameter, which is the cumulative release, normalized to the totaldrug in the system, at time = 0. The total number of independent drug pools is given by n, fi is the fraction of the total drug occupying the j’th pool at time = 0, and kj is the rate constant of the diffusion of the drug from the j’th pool. For data derived from liposome-drugsystems in whichthe drug was at equilibrium distribution at the onset of the experiment, this form of data analysis will yield the percentage of encapsulation through the magnitudeof fi for the encapsulated pool. It is proposed that, althought extracted from kinetic data, this is the most accurate evaluation of this thermodynamic parameter, since it is free of the limitations imposed by separation procedures (see Section VI.A above). Being diffusion processes, it was anticipated and verified experimentally [56-58,691 that theproperties of the drug are the primary parameters dictating the specifics of drug release. For a given drug, liposomal properties such as liposome type, lipid composition, and liposome concentration

286

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constitute means for some modulation drug release. Among those liposomal parameters, liposome concentration was found to be the most useful tool for thetask [56-58,691. It has been shown that the rateconstant of the release of the encapsulated drug generally decreases with the increase in liposome concentration [69]and that the phenomena and trend are indepe dent of drug and of liposome species, whereas the actual magnitudes are system-specific. Besides providing an understandingof the liposomal system of interest, how then can such studies be used for the attainment of those non-in vivo objectives that are among the concerns of the industrial scientist? With respect to shelf life, if the selected dosage form is liposome suspensions, storage should obviously be at high liposome concentrations. Furthermore, the kinetic parameters determinedfor the system of interest become product specifications (or “fingerprints”) that will constitute critical input into the data base on which the decision of whether to storewith or without unencapsulated drug will rest. If the selected dosage form is a freeze-dried powder of drug-encapsulating liposomes, these fingerprints can be useful in defining the optimal conditions for reconstitution and in verifying that the reconstituted system has retained its original properties. Regardless of the dosage form selected for storage, retention of the same , magnitudes of the kinetic parameters can beincluded within the batteryof quality assurance tests. When it comes to surface-modified liposomes, the processes of drug release add some concerns that are of interest to both the liposome investigators in both academic andindustrial research. A particular concern is the risk of drug (encapsulated) loss that can occur in the course of the modification itself, as well as in the subsequent proceduresof separation andpurification. Kinetic studies of the type discussed in this section can be used to determine whether such losses are significant at all and to evaluate their extent. For thesystems at risk, inclusion of drug in the wash buffers could eliminate the problem. Whether the modification is done on preformed drug-encapsulating liposomes oron a singlelipid component prior to liposome formation, such studies can also address the extentto which (if at all) the modification interferes with drug release and the optimal conditions for minimizing that interference. In conclusion, studies of drug release from liposomes that can be categorized as basic research and thereforeinherently within the domain of the academician are also an essential part of many aspects and needs of product development and maintenance that are delegated to thedomain of the industrial liposomologist. Such studies are needed anew for eachdrugliposome system, conducted with the specific drug of interest rather than with models.

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C. Biological Activities of Liposome-Encapsulated Drugs The obviousness of the need to retain the therapeutic activity of the liposome-encapsulated drug is so clear and unambiguous that it does not necessitate, besides this statement, any further discussion. The context in which this issue is discussed here centers on basic and product-oriented pre-in vivo studies that are deemedcritical for theevaluation and application of liposome-drug systems. Monitoringand ensuring the biologicalactivity of the liposomeencapsulated drug has the highest priority whether the studies are conducted at the basic research level, at the stage of produce development, or in the course of routine production of a pharmaceutical product. The span of systems studied varies among these objectives, and are the most extensive at the basic stage, where it is attempted to best understand the system, optimize it, and elucidate the operating mechanisms. Even for productswherethe finaldecisionmight favor the administration of a liposomal system that contains both encapsulated andunencapsulated drug, the basic studies should explore both the complete system and a preparation free of unencapsulated drug. This is to verify that the biological activity is indeed that of the encapsulated drug, since if not, it defeats the objective of using liposomes. In light of the discussions in the previous sections, the time span between separation and initiation of the activity experiment should be selected according to the properties of the system at hand, with the goal of minimizing the loss of the encapsulated drug to thenew drug-free medium. In addition to the obvious controls of free drug and vehicle, further verification that it is the encapsulated drug whichis responsible for all (or at the least most) of thetherapeutic activity, drug-free liposomes suspended in the vehicle and in free drug should also be tested. Where modified liposomes are concerned, at least part of those studies should be duplicated for the (control) unmodified liposomes also. For those cases in which the basic studies have ensured that the designated product (i.e., the encapsulated drug in the specified liposomes) has retained a satisfactory level of biological activity,the product development stage can forego controls of drug-free liposomes suspendedin vehicle and in free drug. Similarly, if theproduct ismodified liposome, nonmodified liposomes can be abandoned. Yet the weight of separately evaluating, the contributions of the system at equilibrium (i.e., containing both encapsulated and unencapsulated drug) and at nonequilibrium (i.e., containing encapsulated drug alone) statescannotbe ignored. Such data should beespecially sought for the final storage and administration dosage forms, be those aqueous suspensions originally, freeze-dried powders that

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are rehydrated, or aerosols. Evidently, once the liposomal system reaches the stage of an established pharmaceutical product, the biological studies can be limited to continuous monitoringof that system alone. It is indisputable that the ultimate preclinical level of organization at which the therapeuticactivity of the liposome-encapsulated drug shouldbe studied is that of the whole animal. Nevertheless, for the objectives listed above, it is well worth the effort to precede with in vitro studies. If done properly in relevant systems, such studies can significantly reduce the investments (such as time, efforts, and resources) needed at the in vivo stage without compromising the quality and significance of the results and their conclusions. Furthermore, for quality assurance of established products, the in vitro studies might suffice. As will be illustrated by several examples below, selecting and implementing in vitro systems for evaluation of the in vivo designated therapeutic activity is an attainable task. In vitro testing of liposome-encapsulating chemotherapeutic drugs for tumor treatment has been extensively studied and reported, anda newcomer to thefield can scan the liposome literature to select the systems suitable to his or her objectives. A similar situation exists with respect to liposomes encapsulating antibiotics for the treatment of intracellular infections of phagocytic cells.l b o additional examples that have not been as extensively reported, and therefore are addressed here in more detail, concern wound healing and the treatment of extracellular bacterial infections. The wound-healingeffects of several growth factors is exerted mainly (if not solely) through their stimulation of cell proliferation. This opens the doorto in vitro testing of the biologicalactivity of suchliposomeencapsulated polypeptides through the selection of suitable cell lines. Basically, two types of cell lines can be used as thetest system: those that have been specifically developed to have absolute dependence on a particular growth factor and those that can be made to have such dependence through the use of specific experimental conditions. For EGF, which is in the first line of growth factors in development for wound healing, the first type is represented by the cell line BALBMK, whichis the classic for EGF bioassay[83]. The well-known NIH3T3 line, growninverylow serum (0.125%), is a representative of the second type [Yerushalmi and Margalit, in preparation]. For both cases, the first task is to determine the doseresponse curves to free EGF, in order to determine the range in which stimulation of proliferation is measurable yet stillfree from adverse effects [83, Okon, Nunez and Margalit, in preparation; Yerushalmi and Margalit, in preparation]. Next, the liposomal systems including the various partial and control groups discussed above can be tested. Implementing suchstudies with both lines, it was found that EGF does indeedretain its biological

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activitywhen encapsulated in regular as well as in surface-modified liposomes. Cultures of the relevant test organism canserve as the in vitro systems for testing liposomes that are designated for infection treatment of extracellular organisms, irrespective of the route of administration or anatomical locations. Adapting the paper disk version of the growthinhibition assay of free [84] to liposome-encapsulated, antibiotics, this approach was satisfactorily implemented for the following cases: liposome-encapsulated ampicillin[G-861 and liposome-encapsulated cefazolin [I. Schumacher and R. Margalit, unpublished data]using Micrococcw Zuteus and Staphylococcus aureus as the respective test organisms. Furthermore the adaptation to liposomal systems is general enough that it can be easily extended to other antibiotic-liposome systems and additional test organisms. In all cases in which the in vivo designated therapeutic activity is tested in vitro, it is imperative to distinguish quantitatively between the total drug dose in the liposomal preparation and the actual dose that has become available to the cells during the time course of the experiment. Treatment with equal total doses of free andof liposome-encapsulated drug where the latter system yields lower response need not be interpreted as liposome-associated loss in drug activity. Nor should similar levels of response to freeand to liposome-encapsulated drug be takento indicate that the liposomal system has no potential to improve clinical outcomes. In both cases, it might simply be that in actuality the comparison was between unequal doses, with the liposomal one beingthe lower. This situation could arise when not all of the encapsulated drug is released and thus made available to the cells during the experiment. Two approaches can be taken in order todistinguish between cases in whichthe equal or lower response of the liposomal treatment is an artifact of comparisons at unequal doses andwhen it is truly due to the loss of activity and/or inadequacy of liposomes for that specific treatment. Both approaches require previous data and analysis of the drug-release profile, especially at the level of dilution used in the bioassay. For the question of whether the liposomeencapsulated drug has retained its activity, such data can then be used to design bioassay conditions under which all of the encapsulated drug 'is made available. For the question of whether the liposomal systems have the potential to improve clinical outcomes, such data can be used to estimate the actual dose to which the cells have been exposed. Obviously, caution should be exercised in making quantitative use of drug-release profiles derived under conditions devoid of living matter. Determination of the kinetics of drug release in the cellular system tested under the conditions of bioassay, when possible, would augment the available data and increase the accuracy with whichthe dose-response curve of the liposomal

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system can beshifted to its true place. Finally, in closing,it is noted that a similar awareness of differences between total and actual liposomal doses is needed for dose-response comparisons in vivo, especially whenthe objective is to evaluate if liposomes can improvethe clinical outcomes.

VII. SUMMARY AND FUTURE PROSPECTS It is the opinion of the authors that any communication on liposomes as delivery systems cannot be concluded without a reminder of the current reality. Awareness of the substantial hurdles to the implementation of liposomes asdrug-delivery systems inestablished treatment modalities cannot be escaped. Nor can the current state of affairs where, despite three decades of research, the number of systems that have matured intoclinical trials and/or have becomeproducts on themarket is quite modest. Yet the need for drug-delivery systems is as acute as ever and the potential that liposomes hold, although somewhat tarnished, has not been substantially diminished. On this background, it is proposed thatat least some of those hurdles can be overcome and substantial strides be made in advancing liposomes from the laboratory to theclinic through attention and mutual awareness of the issues faced by liposome researchers in the academic and industrial environments and through increased collaborations. It is hoped that this chapter will make a contribution, however modest, in these directions. REFERENCES 1. A. D. Bangham, Liposomes: The Babraham connection, Chem. Phys. Lipids,

64:249 (1993). 2. G. Gregoriadis, Liposomes as Drug Carriers, Recent Trends and Progress, Wiley, New York, 1988. 3. T. Sat0 and J. Sunamoto, Recent aspects in the useof liposomes in biotechnology and medicine, Prog. Lipid Res., 31:345 (1992). 4. D. C. Litzinger and L. Huang, Phosphatidylethanolamine liposomes: Drug delivery,genetransferandimmunodiagnosticapplications,Biochim.Biophys. Acta, 1113:201 (1992). D. D.Lasic,Stericallystabilizedliposomes,Biochim. 5. M.C.Woodleand Biophys. Acta, 1113:171 (1992). 6. J. H. Senior, Fate and behavior of liposomes in vivo: A Reviewof controlling factors, CRC Crit.Rev. Therapeutic DrugCamer Sys., 3:123 (1987). 7. P. Machy and L. Lesserman, Liposomes in Cell Biology and Pharmacology, Libbey, London, 1987. 8. G. Gregoriadis, Overviewof liposomes, J. Antimicob. Chem., 28:39 (1991). 9. J. A.Karlowskiand G . G. Zhanel,Concepts on the use of liposomal

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49. C. K. Field,andM.D.Kerstein,Overviewofwoundhealinginamoist environment,Am. J. Surg., 176:2S (1994). 50. C. S. Burton, Venus ulcers, Am.J. Surg., 176:37S (1994). into 51. T. M. Allen and A. Chonn, Large unilamellar liposomes with low uptake the reticuloendothelial system, FEBS Lett., 223:42 (1987). 52. A. Gabizon and D. Papahadjopoulos, Liposome formulations with prolonged circulation timein blood and enhanced uptake by tumors, Proc. Natl. Acad. Sci. USA, 85:6949 (1988). 53. T. M. Allen, C. Hansen, F. Martin, et al., Liposomes containing synthetic lipid derivativesof poly(ethy1ene glycol) show prolonged circulation half-lives in vivo, Biochim. Biophys. Acta, 1066:29 (1991). 54. G. Poste, Liposome targeting in vivo: Problems and opportunities, Biol. Cell, 47:19 (1983). 55. R. Margalit, Vesibes as topical drug delivery systems, in Vesicles, Surfactant Science Series (M. Rossof, ed.), Marcel Dekker, New York, 1994 pp. 527-560. 56. R. Margalit, M. Okon, N. Yerushalmi, andE. Avidor, Bioadhesive liposomes fortopicaldrugdelivery:Molecularandcellularstudies, J. Control. Rel. 19:275 (1992). 57. N. Yerushalmi and R. Margalit, Bioadhesive, collagen-modified liposomes: Molecular and cellular level studies on the kinetics of drug release and on binding to cell monolayers, Biochim. Biophys. Acta, 1189:13 (1994). 58. N. Yerushalmi, A. Arad, and R. Margalit, Molecular and cellular studies of hyaluronic-acid modified liposomes as bioadhesive carriers of growth factors fortopicaldeliveryinwoundhealing,Arch.Biochem.Biophys.313:267 (1994). R. T. Proffitt, Selective in vivo localization 59. E. A. Forssen, D. M. Coulter, and ofDaunorubicinsmallunilamellarvesicelsinsolidtumors,CancerRes. 52:3255 (1992). 60. A. A. Gabizon, Selective tumor localization and improved therapeutic index of anthracyclines encapsulated in long-circulating liposomes, Cancer Res., 52:891 (1992). 61. J. Senior and G. Gregoriadis, Dehydration-rehydration vesicle methodology facilitatesanovelapproachtoantibodybindingtoliposomes,Biochim. Biophys. Acta, 100358 (1989). 62. A. Mori,A. L. Klibanov,V. P. Torchilin, and L. Huang, Influence of the steric barrier activity of amphipatic poly(ethyleneglyco1) and ganglioside GM1 on the circulation time of liposomes and on the target binding of immunoliposomes in vivo, FEBSLett., 284:263 (1991). ofliposomesin 63. S. Toshinoriand J. Sunamoto, Recent aspects in the use biotechnology and medicine, Prog. Lipid Res., 31:345 (1992). P. Machi, in Vesicular 64. L. Lesserman, C. Langlet, A.-M. Schmitt-Verhulst, and Transport, Part B (A.M. Tartakoff, ed.), 1989, pp. 447-471. 65. L. Huang and X. Zhou, Targeted delivery of DNA by liposomes and polymers, J. Control. Rel., 19:269 (1992). 66. A. A. Bogdanov, L.V. Gordeeva, V. P. Torchilin, and L. B. Margolis, Lectin-

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bearing liposomes: differential bindingto normal and to transformed mouse fibroblasts, Exp. Cell. Res., 181:363 (1989). S. Amselem, R. Cohen, and Y. Barenholtz, In vitro tests to predict in vivo performance of liposomal dosage forms, Chem Phys. Lipids, 64:219 (1993). M. K. Jain and R. C. Wagner, Introduction to Biological Membranes, Wiley, New York, 1980. R. Margalit, R. Alon, M. Linenberg, et al., Liposomal drug delivery: therJ. Control.Rel., 17:285 modynamicandchemicalkineticconsiderations. (1991). L. D. Mayer,M. B. Bally,M.J.Hope,andP.R.Cullis,Uptakeof antineoplastic agents into large unilamellar vesicles in response to a membrane potential, Biochim. Biophys. Acta, 816:294 (1985). M. Friede, M. H. V.Van Regenmortel, and F. Schuber,Lyophilizedliposomes as self items for the preparation of immunogenic liposomes-peptide conjugates, Anal. Biochem., 211:117 (1993). L. M. Crowe, C. Womersley, J. H. Crowe, et al., Prevention of fusion and leakage in freeze-dried liposomes by carbohydrates, Biochim. Biophys. Acta, 861:131 (1986). L. M. Crowe, J. H. Crowe, A. Rudolph, et al., Preservation of freeze-dried liposomes by trehalose, Arch. Biochem. Biophys., 242:240 (1985). I. W.Kellawayand S. J.Farr.Liposomesasdrugdeliverysystems to the lungs, Adv. Drug. Del. Rev., 5:149 (1990). P. K. Gupta and A. J.Hickey, Contemporary approachesin aerosolized drug delivery to the lungs,J. Control. Rel.,17:129 (1991). H. Schreier, R. J. Gonzalez-Rothi, and A. A. Stecenko, Pulmonary delivery of liposomes, J. Control. Rel.,24:209 (1992). S. J. Farr, I. W. Kellaway,and B. Carman-Meakin, Comparison of solute partitioning andefflux in liposomes formed by a conventional and aerosolized method, Int. J. Pharmaceut., 51:39 (1989). K. M. G. Taylor, G. Taylor, I. W. Kellaway, and J. Stevens, The stability of liposomes to nebulization, Int.J. Pharmaceut., 5857 (1990). R. W. Niven and H. Scherier, Nebulization of liposomes. I. Effects of lipid composition, Pharmaceut. Res., 7:1127 (1990). R. W. Niven, M. Speer, and H. Scherier, Nebulization of liposomes.11. The effects of size and modeling of solaute release profiles, Pharmaceut. Res., 8:217 (1991). R. W. Niven, T. M. Carvajal, and H. Scherier, Nebulization of liposomes. 111. The effectsof operating conditions and local environment, Pharmaceut. Res., 9515 (1992). S. B. Debs, R. M. Straubinger,E. N. Brunete, et al., Selective enhancement of pentaminide uptake in the lungs by aerosolization and delivery in liposomes, Am. Rev. Respir. Dis., 135731 (1987). C. Carpenter and J. Zendegui, A biological assay for epidermal growth fact uorogasterone and related polypeptides, Analyt. Biochem., 153:279 (1985). Y. Akimoto, Y. Mochizuki, A. Uda, et al., Concentrations of ampicillin in ,

70. 71. 72. 73. 74. 75. 76. 77. 78. 79. 80.

81. 82. 83.

84.

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humanserumandmixedsaliva following a single oraladministration of lenampicillin, J. Nihon Univ. Sch. Dent., 32:14 (1990). 85. I. Schumacher and R. Margalit, Chemical andbiological assays for liposomeencapsulated antibiotics using ampicillin as the test drug, submitted (1995). 86. I. Schumacher and R. Margalit, Physicochemical and biological characterization of liposome-encapsulatedampicillin, submitted(1995).

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11 Stealth Liposomes Danilo D. Lasic* Liposome Technology,Inc., Menlo Park, California

I. Introduction 11. Liposomes in Drug Delivery

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111. Stability of Liposomes A. PhysicalStability B. Chemical Stability C. Colloidal Stability D. BiologicalStability

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IV. Sterically Stabilized (Stealth) Liposomes A. Theoretical Understanding of Steric Stabilization of Liposomes B. An Extended DLVO Interparticle Potential

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Coupling of Polyethylene Glycol to Lipids

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VI.

Some Experimental Results and Discussion

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Applications of Sterically Stabilized Liposomes A. Medical Applications

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VII.

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~~

*Currentaffiliation: MegaBios Corporation, Burlingame, California.

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Lasic

INTRODUCTION

Liposomes are spheric$, self-closed vesicles of colloidal dimensions with (phospho)lipid bilayers that sequester part of the solvent, in which they are suspended, into theirinterior [l-31. With respect to their size and number of lamellae, one defines small or large unilamellar vesicles or, in the case of many concentric bilayers, large multilamellar vesicles (Fig. 1) [2]. Owing to their structure, chemical composition, and colloidal size, which can be well controlled by preparation, liposomes possess several properties which can be useful in various applications [3]. The most important properties are small, uniform, and controllable size and special membrane and surface characteristics which include bilayer phase behavior, its mechanical properties and permeability, charge density, the presence of surface-bound or grafted polymers, or attachment of special ligands, respectively. Additionally, owing to their amphiphilic character and very large surface to volume ratios, liposomes are a powerful solubilizing system for a wide range of compounds. Apart from these physicochemical properties, liposomes prepared from natural lipids exhibit many special biological characteristics, including biocompatibility,biodegradability, and (specific) interactions with biological membranes and various cells. Because of these properties, liposomes are used in various applications such as solubilizers for difficult-to-dissolve substances, dispersants, sustained-release and delivery systems for microencapsulated substances, stabilizers, protective agents, and microreactors. In addition, because of

unilamellar vesicle

vesicle

Fig. 1. Schematicpresentation of variousamphiphilicaggregatesandphases formed by different amphiphiles at different concentrations in mixtures with water Open sphere presents hydophilic part of the molecule while the hydrophobictail is shown as a wavyline.

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their similarity to thecell membrane, they are used in many studies in cell biology and medicine[2,3]. Bilayers can be made entirely from naturally occurring substances and are therefore nontoxic, biodegradable, and practically nonimmunogenic. This makes them ideal candidates for drug-delivery systems. Apart from well-known endeavors in drug delivery, the industrial applications include liposomes asadjuvants in vaccination, signal enhancers/carriers in medical diagnostics, analyticalbiochemistry, and transfection vectors in genetic engineering. Currently, however, thelargest market is incosmetics, in which liposomes serve as dispersants for various substances and/or as a support matrixfor various ingredients and, according to some reports, as a penetration enhancer [4]. Although some of the claims of cosmetic products are not supported by rigorous scientific tests, one can surely say that liposomes definitely represent an improvement simply because of the fact that hydrophobic substances are suspended in an aqueous and biocompatible matrix as opposedto creams andgels, which use skin-irritating oils, alcohols, and surfactants [3].

II. LIPOSOMES IN DRUGDELIVERY The optimistic expectations of liposomes as drug-delivery systems have not been met mostly because of their stability problems. Later, we shall show that theproblems with the shelf lifestability were in mostcases successfully solved, whereas thestability of liposomes in liposomicidalenvironments of biological systemspresented a greater challenge that only recently has been solved [S-lo]. Numerous experiments have shown that intravenously administered liposomes quickly released the encapsulated or membrane-bound molecules. It was found thatthe leakage can begreatly reduced by the inclusion of cholesterol into the membranesof liposomes. But this did not help much in the fast clearance and sequestration of liposomes by the mononuclear phagocytic (MP) cells of the immunesystem located mostly inthe liver and spleen, which is the major drawback in most of their applications. The first attempts to alter their biodistribution by either surface ligands or membrane composition were undertakenin the late 1970s. The results showed that liposome disposition can be altered, but predominantly within the mononuclear phagocytic system, including the intrahepatic uptake itself. In the absence of an undesired increase of blood circulation times by the presaturation, suppresion, and impairmentof the MP system, blood circulation times were prolonged using neutral and small liposomes composedof rigidlipids and cholesterol. The first substantial improvementswere achieved by the incorporation of ganglioside G,, or phosphatidylinositol at

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1

Secondary amine.linkage

Fig. 2. Structure of thepoly(ethy1enegy1col)-lipidmoleculeN-(carbamyl",poly-(ethyleneglycolmethyl ether)-1,2-distearoyl-sn-glycero-3-phosphoethanolamine), normally sodium salt. Usually molecular weights of PEG are 750, 1000, 2000,3000,5000, and 12000. Carbamate and secondary amine linkages are shown. (Courtest of S. Zalipsky, Liposome Technology, Inc.)

5-10 mol% into the bilayer. The best results, however were obtained by substituting these two lipids with synthetic polymer-containing lipids. The longest circulation times were achieved when polyethylene glycol covalently bound to thephospholipid was used (Fig.2). It seems that intermediate molecular weights from 1500 to 5000 da at about5 mol% in the bilayer give rise to the longest blood circulation times [lo]. Fig. 3 shows bloodclearance profiles of several different formulations. Surface grafting of polymers offers much better control overthe colloidal properties. In the case of liposomes, surface adsorption of polymers normally leads to leakage of the encapsulated molecules or even possible aggregation of liposomes due to bridging interactions. It is believed that theenhanced stability is due to the mrface-attached polymer which represents a steric barrier to the interacting macromolecules. For this reason, they areoften called sterically stabilized [lo] liposomes, whereas commercially they are often referred to as Stealth (a registered trade nameof Liposome Technology, Inc.) liposomes because of their invisibility to the body's immune system (Fig. 4) [ll].

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307

l

v

0

5

10

15

20

I

25

Hours

Fig. 3. Blood clearance profiles of several liposome formulations. From top to bottom: 2000PEG-DSPE/EPC/Cho1, EPC/Chol(2/1), EPG/EPC/Chol. formulations withmolarratio 0.15/1.85/1. Abbreviations: PEG, methoxypoleythyleneglycol; DSPE, distearoylphosphatidylethanolamine;EPC.partiallyhydrogenatedegg phosphatidyl choline (IV 40), Cholesterol; EPG, egg phosphatidyl gylcerol. 100 nm liposomes were labeled with 67Ga-desferal. (From ref. 59.)

In order tounderstand this phenomenon and owing to the confusion in the field, we shall briefly review the stability of liposomes, and on the basis of this understanding, show how to make vesicle preparations with enhanced stability in a particular medium. 111.

STABILITY OF LIPOSOMES

In liposome research and applications, the term stability is used rather vaguely and is often not properly defined. In the broadest sense, liposome stability involves the stability of the particle and its constituents, including the entrappedmolecules in any, includingliposomicidal, environment. This general definition of stability contains various aspects which can bedefined separately but oftensynergistically act together [3]. We shall somehow arbitrary define stability as being a sum and a product of physical,chemical and colloidal constituents. In addition to these inherentstability components, applications of various liposome products require us also to define the stability of liposomes on application, which is often in a liposomicidal (liposome deleterious) environment, and the long-term stability of the product. Most of them are of a pharmaceu-

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Stealth Liposome

(b)

Conventional Liposome

Fig. 4. Schematic presentation of (a) stealth and (b) conventional conventional and Stealth liposome. Polymer chains can be attached on one or both sidesof the vesicle. (Courtesyof Phillip Le Roy Carter, Liposome Technology, Inc.)

tical natureand,thereforeone candefinebiological and shelf life stabilities, which is a cumulative property of several particular stabilities, respectively. A.

Physical Stability

This aspect of liposomes mostly involvespreservation of the liposome structure and entrapment; thatis, the stability of liposome size and shape (size distribution of the sample) and 'the barrier properties of the bilayers for the encapsulated or membrane-bound molecules.

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Both the preservation of the size as well as encapsulation depend on the mechanicalproperties of the bilayers and thermodynamics of the system. Phospholipid molecules in the vesicle membrane are in equilibrium with free molecules in solution at concentration of critical micellar concentration (CMC). The values of CMC for phospholipids are very low, norM, and this practically does not affect liposome properties as mally < encountered in laboratory work. Actually, having in mindthese low values and being aware of the slowness of these processes, one can in principle also question the concept of CMC at all. From the kinetic point of view, this looks rather more like a lipid solubility than a quick, dynamic process as observedin micellar solutions [12].

l . Thermodynamic Stability Helfrich has shown that all single-componentbilayers and symmetrical bilayers increase their free energy oncurving, because the spontaneous curvature of the system, C,, is zero. For large bilayered fragments, one can assume that the bending is elastic and the excess energy due to curvature can be expressed as 1 E = - K (C, + C,- CO)* + k C,C* 2

where C, and C, are the principal curvatures, C, is the spontaneouscurvature, K is the bending modulus, andk is the bending modulusof Gaussian curvature. In the simplest case of bending a plane into a sphere, principal curvatures are equal and reciprocal to the radius of curvature, R. On closure, the radius of curvature equals the liposome radius, and one can write E=81rK+41rk

(2)

Assuming K > > k , we shall neglect the last term. Eq. (2) shows the excess energy of the self-closed system due to the bending [12-141. Because E exceeds thermal energy, that is, E-10-50 kT, where k is the Boltzmann constant and T is temperature, the entropy, which is, according to the Equipartition principle, 3/2 kT per liposome, cannot stabilize the system. unstable From 81rK < kT, it follows that liposomes are thermodynamically [3,12,13]. Neglecting k , Eq. (2) shows that theoretically there are two possible ways to produce thermodynamically stable liposomes: entropic stabilizaC, in the case of tion requires E = kT, whereas E = 0 for C, = C, asymmetrical bilayers [151. K is proportional to thethickness of the membrane. Membrane thinning due to the interdigitation of bilayers or action of cosurfactants, as in

+

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microemulsions, can give rise to K = kT, and entropically stabilized vesicles can be formed. Another way to produce thermodinamically stable liposomes is to create nonzero spontaneouscurvature. This can bedone by asymmetrical bilayers or by asymmetrical conditions across the bilayer, such as different values of pH, concentrations of counterions, or interacting (macro)molecules. Furthermore, it is likely that some of the novel lyotropic phases, such as cubic sponge phase, can spontaneously disintegrate into liposomesolution on dilution owing to the large critical fluctuations and low values of K . Similar intermediate structures may also occur in some liposomepreparation methods, such as in demulsification of particular reverse phases or in some lipid-detergent mixtures during the preparation of vesicles by the detergent-depletion technique. Here, however, the changes are driven by the removalof the nonpolar phase[3]. The entropically stabilized spontaneous vesicles are not interesting for applications, because owing to their existence, they have low values of K,which inherently reduce theirbamer and permeability properties. Alternatively, the existence and stability of these vesicles canbebased on transmembrane gradients which can dissipate. In contrast, asymmetrical bilayers, including some novel amphiphiles, such as bipolar lipids with asymmetrical polar heads, or polyethylene glycol (PEG)-lipids, can make very stable and nonpermeableliposomes with C,, = 0. The stability of liposomes against external perturbations such as dilution can be understood by a very low CMC value of lipids and the large energy required to open a liposome. The energy to create a pore, break upthe liposome, and causefusion or formation of liquid crystalline phase in excesssolvent is [161

E, = 2 m r - IT r2y

(3)

r

where y is the surface tension and the edge interaction at the polarnonpolar interface of exposed bilayer. Contamination of bilayers with single-chain surfactants, which can effectivelyshield the interface, may reduce E,, [3,12]. Normally, however,it still has arather large value ( 2050 kT) and pores, even if formed, quickly reseal. This can explain why vesicles can be relatively stable. All this indicates that liposomes can be kinetically trapped into their structures, because they are formed ina nonequilibrium irreversible process in conditions of high excesses of energy, and the energy needed to open or break them canalso be = 50 kTfor r > R. Moreover, a simple Gedanken (thought) experiment shows the advantage of kinetically trapped systems as opposed to systems at thermodynamic equilibrium for applications such as microencapsulation. A system in

-

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305

equilibrium quickly adapts to the external change, such asdilution, concentration, pressure, the presence of other substances, injection into body fluids, or temperature by a phase change. For instance, micelles or microemulsion droplets quickly disintegrate on dilution (i.e., move in phase diagram into different regions), whereas liposomes made from phospholipids with low values of critical vesicle concentration are stable against dilution. This simple analysis can explain many of the unexpected data from the drug delivery by colloidal carriers. Furthermore, it is interesting to note thatliposomes can be stabilized against some phase transitions, such as freeze-thawing and freeze-drying, by the addition of water-soluble cryoprotectants, such as sucrose. This enables manufacturing of freeze-dried liposome formulations.

2. Mechanical Stability According to the theory of smectic liquid crystals, there are three different ways to deform a thin membrane: area dilationkondensation, surface extension at constant area (shear),and bending. Each deformation is characterized by the appropriatemodulus and the knowledge of material properties of the bilayer can be used to improve the behavior of membranes under stresses; that is, prepare more stable liposomes. Deformation can be a consequence of osmotic imbalances, adsorption, hydrodynamic shears, and other forcesand results in changes of vesicle shapes, membrane tension, and possibly rupture and closure with reequilibration of internal and external solutes, topological changes, such as the formation of buds, invaginations, or daughter vesicles, or simply liposome disintegration. Obviouslymechanically stronger,that is, more cohesive, bilayers resist external perturbations better. Micropipette manipulation [l71 studies have shown that the strongestbilayers consist of equimolar DSPC and cholesterol. Quantitatively this observation was explained by a theory in which cholesterol is regarded as a crystal beaker of the gel phase and inducer of chain ordering in the fluid phase without rigidification of the overall phase. This gives rise to the liquid-ordered phase, which has a high degree of conformational order and no resistance totheshear forces. It was found that liposomes behave as tiny osmometers. However, they can sustain large osmotic imbalances before they break. Experimentally, it is known that, for instance, DSPC/cholesterol (3/2 mol/mol) can sustain osmotic pressure of several dozens of atmospheres without bursting. This can be explained by the above arguments of high mechanical strength of membranes. Often when the pressureis too high, liposomes can transiently open, reequilibrate their contents, and quickly reseal, which

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explain minimal changes in size distribution after osmotic shocks in some systems. Bilayers can be also mechanically stabilized by polymerization [3] or by using lipids with fluorocarbon chains [18]. Despite reduced van der Waals attraction among carbon chains, these lipids form stronger andless permeable bilayers owing to stronger hydrophobic interaction [3,18]. Permeability of bilayers depends onthe natureof permeating species, mostly on their charge, size, hydrophobicity, and hydration, and can be correlated to the phase behavior and mechanical stability (i.e., cohesivity) of the bilayer and structural defects. In addition, it also influences the bilayer stability against chemical degradation. Although the thermodynamic behavior of bilayers and its correlation to permeability has beenknown for quite sometime, the strong correlation betweenthe mechanical properties of the bilayers and the stability of liposomes was realized only recently.With several other surface characteristics, this knowledge allows one tocontrol the stability and interactability of liposomes andrationally design their properties.

B. Chemical Stability Lipids are very sensitive biomolecules and quickly undergo different degradation reactions. The most common degradation pathways are oxidation and hydrolysis. In general, polar heads are much more stable than the hydrocarbon chains. Hydrocarbon chains, especially the unsaturated ones, are subject to oxidation, whereas hydrolysis pinches off a hydrocarbon chain from the backbone molecule. In the case of double-chain phospholipids, it produces a fatty acid and lyso lipid. Ultimately, it produces two fatty acids, glycerol, and a free polar head. Because the reaction is acid and base catalyzed, it exhibits strong pH dependence [19-211. The protective measures include optimization of pH, normallyaround 6.5, and, preferentially, freezing or drying thesample in thepresence of cryoprotectants. Oxidation is a radical reaction which ultimately results in the breakage of chains or, in the case of two adjacent double bonds,the formation of cyclic peroxides. Protective measures which include the useof saturated lipids and the addition of antioxidants, such vitamin E (a-tocopherol) or BHT, results in a substantial reduction of oxidation and samples can be stable (within tight specifications, normally 5% degradation) for years [3,21]. Chemical and physical stability are closely related. Mechanically strong andwell-packed bilayers reduce theaccess of oxidizing or hydrolyzing agents. This in turn reduces the changesin sizedistribution, fusion, and

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307

mechanical properties of the bilayer owing to thereduction of the concentration of the degradation products, which influncethe mechanicalcharacteristics of bilayers such as bending and stretching elasticity, molecular reorientation rate (flip-flop), and permeability [3]. Several synthetic lipids, including the ones with fatty acids linked to the glycerol backbone via ether linkage, are chemically more stable. However, they are not biodegradable by mammalian enzymes, and this limits their usein parenteral drugdelivery, whereas they are often used in chemical or cosmetic applications. The lytic rate of various degradative enzymes can be reduced by using lipids with unnatural chirality. For instance, almost all naturally occurring lipids are levorotary, that is, “left-handed.” Obviously, dextrorotary lipids are less susceptible to the attack of various phospholipases and yield chemically more stable liposomes in biological environments. Of course, chemical stability becomes a much more complex issue when degradable substances are encapsulated. In these cases, the optimal stability is a compromise betweenvarious contributions, and in some cases, it may not be achieved even in a frozen or freezekpray-dried liposomal product. C.ColloidalStability Changes in the particle size distribution in colloidal systems occur mostly via two mechanisms: On the molecular level, this can be an asymmetrical molecular exchange, whereas on theparticle level, this is mostly aggregation, flocculation, and/or fusion. The first process is the so-called Ostwald ripening and the latter, in the case of liposomes, aggregation and fusion. 1. Ostwald Ripening

In this process, larger particles grow at the expense of smaller ones. Smaller particles have a larger surface to volume ratioand, additionally in the case of vesicles, higher curnature gives rise to a higher chemical potential. Assuming that the concentration of exchangeable monomersis equal to the CMC, we see that this size disproportionation is negligible for most M. Forsome synthetic phospholipids, whichhavevalues of CMC < surfactants, however, the CMC can bemuch higher, and this becomes the dominant s u e growth mechanism. In the case of sodium heptylnonyl benzenesulfonate, with a cmc of of 5 mM, it was shown that vesicles grow proportionally to to”;that is, they double their size to 40 nm in40 days [22]. UsingthesameequationsandtheCMC =5 x M of lecithin,onegetsthe kinetics of apparent growth < 5 x 10” nm/day.

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2. Aggregationand Fusion In phospholipid liposome preparations, the increase of the particle size is mostly due to particle aggregation, because the size growth due to Ostwald ripening is negligible. Aggregation is the first step in a vesiclevesicle fusion which depends on the thermodynamicstability of the sample, crystal defects, and the presenceof intrinsic or exogenous fusogenic substances or vectors. Fusion is a practically irreversible process, whereas aggregation can be reversible: Often liposome systems are only loosely aggregated, and mild swirling of the sample redisperses liposomes. Here we shall discuss only the origin of aggregation. First,we shall define different forces acting betweenliposomes, comment on the DLVO theory, and develop a more general interparticle potential. Later, we shall use this concept to simulate the interaction of bilayers with macromolecules. We shall not be concerned with the kinetics of aggregation and subsequent fusion, which was reviewed extensively [23,24]. There are several attractive andrepulsive forces which act between colloidal particles. Ubiquitous attractive forceis van der Waals attraction, whkh acts between all particles separatedby a medium of different polarizing properties. In the case of charged particles, there is a strong electrostatic force. Mixing two samples of liposomes bearing opposite charges results in flocculation.Mostly, however, particles bear the same charge, and this results in electrostatic repulsion,which leads to a strong repulsive force. Theabove-mentioned van der Waals attractionandelectrostatic repulsion forces are thebasis of the DLVO theory of the stability of colloidal particles [25]. In many cases, however, this theory could not explain the stability of liposomes against aggregation. In the case of predicting a too stable colloidal suspension, additional attractive forces, such as hydrophobic attraction or ion-ion correlation attraction, were introduced, whereas to explain the stability of uncharged liposomes and liposomes in high ionic strength media, otherrepulsive forces were recognized. The aggregation of noncharged liposomes is normally prevented by the repulsive hydration force,which is due to the hydration of polar heads. In order tobring two bilayerstogether, onehas to remove water molecules from the polar heads. This results in a repulsive force and so does the vertical motionof polar heads,which causes the so-called protrusion repulsive force. In soft bilayers that can thermally undulate, another repulsive force was recognized. This undulation forceis due to theexcluded volume effect two approaching undulating membranes exert on one another. order In to approach, they have rigidify, to This is entropically unfavored and results in a repulsive force.

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309

These forces can explain the colloidal stability of conventional liposomes and provide a basis for their stabilization. Historically, mostly surface charge density, zetapotential, size distribution and homogeneity,ionic strength, and pH of the medium were found to determine colloidal stability in vitro. In biological systems, however, the behavior is completely different because of the presence of active liposomicidal substances. Also, ionic strength is around 290 mM. Later we shall show additional repulsive interactions originating from the surface-attached molecules. Since Egyptian times it has been known that lyophobic colloids, such as pigments in inks, can be stabilized by natural polymerssuch as arabic gum, starch, or cellulose. This gives rise to the so-called steric repulsion, which willbe discussed in Part IV. D. BiologicalStability

Even the most stable conventional liposomes are rather unstable in the biological milieu. In addition to interactions with various plasma components and enzymes, an active uptake of liposomes by the body’s immune system takes place (Fig. 5 ) . It has been shown that mechanical stability can help reducethe interactions in vitro and reduceleakage in vivo but cannot slowdown appreciably the uptake of liposomes by phagocytic cells on parenteral administration. It was assumed [6] that the predominant mechanisms of liposome clearance from blood were (1) opsonization by immune and nonimmune components in blood, such as immunoglobulins, complement factor proteins, and fibronectin, which trigger subsequent macrophage uptake; and (2) reaction with plasma (lipo)proteins, which can be expressedas

Fig. 5. Interactions of liposome with cells. There are predominantly four possible interactions: adsorption, lipid exchange, fusion and endocytosis. (From ref. 82.)

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L+O===>LO L 0 + R-M ===>

(4)

M-R 0 L E==>M (L)

(5)

and PLP(m) PLP

PLP PLP(n) PLP

U

PLP(1-m-n)

U

L(1) ===> L(1-m) ===

U

L(1-m-n)

===>

0

(6)

where: L is liposome containing 1 lipid molecules, 0 is opsonin, M is macrophage, R is receptor for 0 ,PLP are plasma lipoproteins which bind m,n lipid molecules. Improved mechanical stability can greatly reduce liposome disintegration as shown by eq. (6). In order for effective opsonization to take place, a macromolecule has to bind to the liposome surface. We believe that this adsorption can be electrostatic, hydorphobic, H bonding, or physisorption due to van der Waals forces; thatis, =

west

+

whfo

+

+ wH~ond + wspec

(7)

where the total adsorption energy contains contributions from electrostatic, hydrophobic,van der Waals, hydrogenbonding, and specific interactions. The latter ones occur mostly when some antibodies or haptens are attached. Obviously, an ideal drug targeting would tend to eliminate all contributions toW,ofwith exception of W,,. Probably in some proteins the adsorption or adhesion requires conformational change to trigger subsequent macrophage uptake. Similar conclusions may be applied to interaction with plasma proteins. Obviously more cohesive bilayers exchange lipid molecules with the surroundings moreslowly. Very cohesive bilayers inthe gel state, however, . containtoo many hydrophobic defects to be stable [3]. This model of liposome uptake can explain the above-mentioned results. Clearly, uncharged liposomes should be more stable, because any strong ionic binding is absent (Wuf = 0). Hydrophobic binding, such as penetration of a segment of the macromolecule into thebilayer, should also be reduced in the presenceof more mechanically cohesive bilayers (reduction of Wh,J. Physisorption, however, is not prevented; that is, W,,, and W,,,,,,", are < 0. Later, we shall show that physisorption can be greatly reduced by the presence of a steric shield of an inert, nonimmunogenic (Wspec= 0), nonionic (W-, = 0) polymer. The absence and reduction of chemisorption, that is, electrostatic and hydropohobic binding, does not reduce the H bonding and physical adsorption of macromolecules onto the liposome. This may explain their lower

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31 1

stability in vivo and their macrophage uptake. In contrast, the presence of a steric shield, as exhibited by a noninteractive surface-attached polymer (shown in Fig.4), prevents orreduces all the differenttypes of macromolecular interactions with liposomes. Results show that in the presence of surface-grafted polymer, both fluid and less cohesive or charged liposomes can exhibit prolonged blood circulation times [26]. This property is important in designing liposomes with well-defined leakage characteristics and containing no cholesterol, as well as improving their encapsulation efficiency for hydrophobic molecules. However, the anchoring of the PEG moiety must be optimized; that is, lipid chains should be distearoyl or should match the fluidity of the bilayer. It seems that in contrasttoelectrostaticstabilization, whichwas found to be effective in many in vitro applications, steric stabilizationis a ubiquitous way to stabilize particles in biological systemsand in media with variable salinity. Of course, stericstabilization can be imposed on charged bilayers, andin these cases, in vitro stability may be increased, whereasthe stability is not additive in the in vivo case. A combination of the two, that is, electrosteric repulsion in the case of surface-attached polyelectrolytes, would probably increase the stability of liposomes because of thicker and more rigid and mutually electrostatically repulsive brushes in vitro. However, as was discussed before, it is unlikely that it would increase biological stability, because W,, # 0. Polyelectrolyte chains bearing opposite charge to thebilayer may, however, disrupt the bilayer integrity. IV. STERICALLY STABILIZED (STEALTH) LIPOSOMES One of the major breakthroughs in the medical applications of liposomes was the realization that neither mechanical nor electrostatic stabilization can substantially improve the stability of liposomes in biological environment, especially in the systemic circulation. In analogy with the stabilization of inorganic and polymeric (latex, polystyrene beads),colloidal particles polymer coating was imposed on the surface of the liposomes. Because adsorption of block copolymers (polyethylene oxide-polypropylene oxide) caused leakage, lipids with covalently attached polymers were introduced, predominantly polyethylene oxide. Before we describe some practical applications of these systems, we briefly describe the theoreticalbackground of steric stabilization and of the polymer behavior at the interfaces. Fig. 4 is a graphic presentation of conventional and polymer-coated liposomes.

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A. Theoretical Understanding of Steric Stabilization of Liposomes The repulsive forces between the two particles can be greatly increased by coating the surface of the particle with appropriate molecules. Natural and synthetic polymers are the most common materials used. They must be inert, well solvated, and compatible with the solvent and have polarizability close to water. Steric stabilization can be inducedby surface attachment of various natural or synthetic polymers, either by adsorption, hydrophobic insertion, electrostatic binding or preferentially by grafting via a covalent bond. Nonionic, water-compatible, flexible, and well-hydrated polymers arepreferred. The repulsion between surfaces with attached polymers was shown to bedependent on thegrafting density and the degreeof polymerization. In the low-grafting densities, that is, in the region where the polymer chains cannot interact among themselves, the so-called mushroom modelwas defined which yields steric repulsion at distance, h, as 1271

where

h.=..(%)” and D is average distance between adjacent grafting points, U is the size of the segment, and N is the degree of polymerization. At higher densities where polymers start to interact, this causes their extension and the socalled brush model [27] can be applied. The repulsive force is now

F,?

kT

= - [(h&)”

D’

- (h/hc)3’4]

These equations explain the repulsion in the case of polymers which form both the so-called adsorption and depletion layer. In the case of depletion layer-forming polymer, the conformation probably looks more like an inverse droplet or pear thatalters the density profile but leaves the scaling laws unchanged [28]. Theoretical analysis predicted extension of the chains to be40% longer as opposed to the model above[29]; that is 7/5 R, vs R,, where Flory Radius R, = a M”.

B. AnExtended DLVO Interparticle Potential The stability of colloidal particles against aggregation is normally described by the DLVO theory. This theory is based on theinterplay of the attractive

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313

van der Waals and repulsive electrostatic forces, and the minima in the cumulative potential indicate the stability regions. Obviously, the theory cannot explain the stability of an uncharged liposome suspension. This can be corrected by invoking hydration repulsion. For the most general case, however, we propose that the total potential between the particles should include all the above-mentioned forces, including steric

+ v,, (11) I have also included the hydrophobic attractive force despite that its existence is still controversial. A treatment, analogous to the DLVO for two potentials, shoule be performed to find stability of the system. In a first approximation, potentials could be treated as additive. For rigorous treatment, however, one may expect dependence betweed various terms. It is reasonable to expect the interdependence of electrostatic, hydration, and steric interaction in the case of charged surface and covalently bound polymers due todifferent water structure at thesurface. Inthe case of grafted polyelectrolyte chains, however, the conformation of chains themselves is a function of electrostatic interactions and surface charge, and obviously different analyses should be performed.

v,, = - v,

V.

-

v,,,+ v,, + v,,, +

VU",

COUPLING OF POLYETHYLENE GLYCOL TO LIPIDS

Apart from various adsorption methods, which are prone to cause leakage of the liposome-encapsulated molecules, PEG polymer chains of various lengths can be covalently coupled to lipids using several different coupling agents. Although shorter oligoethyleneoxide spacers have been coupled to many different lipids [3], the only lipid used for the attachment of longer PEG chains was phosphatidylethanolamine with different chain lengths and degreesof saturation because of the reactivity of the amino group.The reactivity of this group is further catalyzed by thedeprotonation by triethylamine. Several researchers also attached PEG chains on cholesterol, butresults in general do not show prolonged blood circulation times. This is not unexpected, because these molecules, such as cholesterol -0(CH,-CH,-O),,-H, are commercially available as surfactants. This means that PEG-lipid molecules can be quickly lost, especially after dilution, from the bilayer owing to their high CMC values. Synthetic chemistry using other phospho- or sphingolipids has not been worked outyet, or the products mayyield rather unstable and hydrolyzable ester bonds. Polyoxyethylene glycols of different chain lengths, including n = 3, 8, 17,45,70, 115, and 270, have been attached. The polydispersity of the polymer depends on thepolydispersity index, p , of the starting material and virtually does not change during the synthesis.

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Commercial-grade material has p , defined as the ratio of weight over number average molecularweight, around 1.1, whereas BioTech grade hasp = 1.05. Different researchers used different coupling strategies, all adopted from the protein-modification field [30].All of them start with methoxyPEG and as a lipid use phosphatidylethanolamine. Three different synthetic routes were employed using succinyl, cyuronic chloride, and carbamate derivatives yielding ester, secondary amine, and uretane linkage, respectively (Fig. 6). Although the rate constant of hydrolysis of the ester linkage produced by the first reaction probably does not contributesignificantly to the liposome uptake, itisless stable than the urethane bond

I mPEG-N-PE H

4

Tresyl-cl 2. PE

0

mPEG-OH 1

1 CD1

mPEG-0

2. PE I TEA

&-PE

2

H

3 1. Succinic anhydride 2. N-hydroxysuccinirnideI DCC 3. PE I triethylamine

Fig. 6. Schematic presentation of four different synthetic routes to link polymer to the lipid. Most frequently used synthesisis the one on the right side. (Courtesy of Samuel Zalipsky, Liposome Technology, Inc.)

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produced by the third reaction. The second coupling method, however, uses a toxic reagent. Owing to the above reasons, we have used the following synthetic route: m-PEG (1) and carbonyl diimidazole in a molar ratio 1:l.l were reacted in dried benzene in nitrogen atmosphere at 75°C for 16 hs, yielding carbamate of mPEG ether (2). Coupling 2 to PE was achieved by refluxing2:3:triethylaminein molar ratio 1:2:2 in benzene at 95°C for 6 hs. The PEG-PE (4) was purified by reverse phase silica gel chromatography by eluting withch1oroform:methanol or water:ethanol mixtures. Because 4 forms micellesin aqueous solutions, this compound can be purified simply by dialyzing micellar solution against water or preferably against saline. PEG-lipids behave asionic surfactants in aqueous solutions and the addition of electrolytes appreciably reduces their CMC values; in the case of 2mPEG-DSPE, CMC decreases from mM to 30 p M on addition of salt [31]. The chemistry of the heterobifunctional PEG chains for the subsequent attachment of antibodies to the far end of the grafter polymer chain was also established [32]. Because all the above syntheses use unreactive mPEG on one side, a heterobifunctional polymer derivative had to be prepared. On oneside, it bears the reactive succinimydil carbonate for the attachment to the amino group of phosphatidylethanolamine and on the other tert-butyloxycarbonyl-protected hydrazide group. Various ligands, including monoclonal or polyclonal antibodies, can belinked to preformed functionalized liposomes containing this conjugate owing to the chemical versatility of hydrazide groups. Recently, several new polymers whichrendered liposomeslong circulating were discovered. Polyethyloxazoline and polymethyloxazoline with a degree of polymerization around 50 and attached to lipid were shown to be equivalent to PEG[33], whereas polyacrylamide and poly(viny1)pyrrolidone prolonged the blood circulation of liposomes to a lesser extent, probably because of a single chain lipid anchor [34]. Preliminary results using several others polymers did not match the prolongation of the blood circulation as achieved by PEG. It seems that they do not combine the same hydrophilicity and flexibility behavior as does PEG or the presence of dipolar interactions. The samewas found true for oligo- and polysaccharides, such as glucuronic acid, dextrans, cellulose derivatives, and copolymers of glucuronic acid with simple sugars. These polymers can form a stronger and denser steric shield, possibly at an increased density of hydrogen-bondingsites but at compromised mobility of the chains. Furthermore, there are manyspecific receptors for various sugars in the body.

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Liposomes containing PEG-lipid can be prepared by using virtually all the standard preparation methods. After thin-film hydration, better dispersal is observed and liposomes show reduced lamellarity (number of lamellae). Unilamellar liposomes, produced by various techniques, are generally characterized by enhanced stability. VI. SOME EXPERIMENTAL RESULTS AND DISCUSSION The hypothesis of increased repulsive pressure above membrane and reduced adsorption on the blood circulation lifetimes was tested by measuring the repulsive pressure between membranes with and without incorporated polymer-bearing lipid. Fig. 7 demonstrates that bilayers containing PEG lipid show much larger interbilayer spacings, and even on strong compression, the bilayers are still 4 nm apart as compared with surface unmodified bilayers, whichshow practical collapse tothe hard wall repulsion [35,36]. These data are comparedwith the theory in Fig. 8. The interbilayer repulsion calculated from the repulsive pressure of surface-attached polymer in a mushroom configuration (broken line), that is, from

P=(

5/2) kTN/D2 a(~/(h/2))~

and for N = 44, a = 0.35 nm, D = 3.57 nm, is in nice agreement with experimental data [36]. For distances > h,, the repulsive pressure is zero. Similar results were also obtained by the surface force apparatus [37]. These results also show increased repulsion with an increasing amount of PEG polymer, whereas biological data find optimal circulation times at lower surface coverages or show a saturationlike behavior at grafting densities which assure complete coating of the surface. Whether this is due to some specific interactions of possibly the flexibility of the chains has not been evaluated yet [lo]. Surface force measurements have foundreversible repulsive force at all separations, and the thickness of the steric barrier was found to be controlled by the amount af added PEG-lipid. At low coverages, Dolan and Edwards' theory of steric forces was found to describe experimental data satisfactorily, whereas at higher coverages, Alexander de Gennes' theory described the data better[37]. At low coverages, in the mushroom regimen, the Dolan-Edwardsexpression for force between two curvedcylindrical surfaces of radius, R , can be described by Pc@)

- 7 2 1 r A k T e x p - -h

"

R

R,

Stealth Liposomes I

31 7

( , , , , I I

-60

-20

20

60

100

140

Distance from Bilayer Center (A)

201%

B

8 OO

0 0 0

l21 10 A V

.

0

20

40

Distance Belween

60

80

100

Bilayer Surfaces dw

l !O

(A)

Fig. 7. (A) Electron density profilesof bilayers with (bottom) and without (top) 2mPEGylated lipid as determinedby S A X S measurements of multilamellar vesicles. ?kro unit cells are shown, and it can be clearly seen that the presence of polymer increases the gap between the bilayers. Furthermore, an increased electron density in the aqueous gap may indicate that the polymer, which is known to form the depletion layer, is stretched away from the bilayer. (B) Pressure versus gapbetweentheadjacentlamellae:solidsquares,stearoyloleoylphosphatidyl 4 mol%of cholinelcholesterol (2/1) bilayers;solidcircles,somebilayerswith 2000PEG-lipid in 100 mM NaCl and in 1 mM NaCl (open circles). For allthe applied pressures, the presence of polymer-bearing lipid markedly increases the aqueous gap betweenthe bilayers. (From ref.36).

Lasic

318

0

20

40

60

80

Distance Between Bilayer Surfaces dw

100

120

(A)

Fig. 8. Theoretical interpretationof the experimental data from Fig. 8. Experimental data of SOPUChol bilayers containing 4 mol% of mPEG-lipid fit into repulsive pressure calculatedfor the mushroom model of steric repulsion (see Eq. [7]). (From Ref. 71.)

where Rg is the radius of gyration of the polymer in a theta solvent and corresponds to the thickness of extending polymer. A is the surface coverage of the polymer. At higher grafting densities, that is, in the brush regimen, the force could bedescribed by

F,(h) - 16 kT T L 35D3 R "

[~($T+~(&T-Iz]

where

L = D ( : ~ The expressions for force between two cylindrical surfaces (F,) and repulsive pressure (F)can be calculated using Derjaguin approximation

Conformational changesin monolayers of PEG-lipids and theirmixtures with phospholipids were measured by film-balance experiments and fluorescent microscopy: twophase transitions were observed-at low coverages pancake-mushroom and at higher mushroom-brush. In the pancake state, phase segregation was observed. Only when a depletion layer was

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319

formed in the mushroom regime, could the lipid molecules intermix with PEG-lipids. Composite polymer-lipid films were deposited on solid substrates and studied by ellipsometry. Disjoining pressures as a function of humidity were studied, and it was found that distance versus pressure curves are governed by steric and electrostatic forces and were in good agreement with surface force apparatusand osmotic stress technique measurements [38]. This simple theory states that higher grafting densities of longer chains result in better stabilization, as experimentally verified above, and presumably longer circulation times. In vivo experiments, however, have shown that the best results were obtained at around 5 mol% of relatively short chains, n = 40-130, and it seems preferably 40-70. This is probably due to several factors. First, in these membranes or surfaces,inclusion of PEG-lipids shows saturation behavior. The overall pressureabovethe bilayer can destabilize liposome by creating hydrophobic defects which then act as sites of attachment for opsonins. Their adsorption may also dependonthe mobility of the polymer chain, which decreasesafter mushroom-brush transition. At higher polymer concentration above the bilayer, the polymer may also exhibit phase transition (collapse of polymer brush) [7].The later hypothesis could be tested by using other polymers which do not have solubility gaps in their aqueous solubility but can still lead to enhancedstability in circulation. The vesicle size dependence of the blood circulation times which decrease with increasing size is still not understood. Although the liver uptake is still low,the spleen uptake increases for 2 R > 200 nm. It is possible that the spleen uptake exhibits size dependence, and such filtering may result in the accumulation of the largersterically stabilized liposomes in the spleen. VII.

APPLICATIONS OF STERICALLY STABILIZED LIPOSOMES

Sterically stabilized vesicles present anew model in studies of polymers and scaling laws, colloid stability, and cell biology. It seems that cells function viaspecific attraction and nonspecific repulsion. Therefore, these liposomes offer a much better model for thestudy of cell function and interaction than conventional liposomes, which can exhibit many nonspecific attractive interactions. This propertyof sterically stabilized liposomes offers several useful applications, mostly in diagnostics and drug delivery. In basic sciences, scaling laws of polymer behaviour near various interfaces can be experimentally verified. This knowledge can be applied to the biomaterialsscience for designing the surfaces which come into contact with body fluids, mostly blood.

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The phase behaviour of various liposomes containing various amounts of PEG-lipids with various chain lengths (well-defined HLB values, HLB = hydrophobic lyophobic balance) can be used to characterize further the liposome formation and stability. A.MedicalApplications

Stealth liposomes provide an entirely new and unique drug-delivery vehicle because of their ability to evade quick clearance by the immune system. Using such liposomes, the goal of specific cell targeting became theoretically possible. Indeed, early studies of Stealth liposomes labeled with antibodies have shown increased delivery of antibody-labeled Stealth liposomes to the targettissue in accessible sites [39]. The accesible sites are in the vascular compartment,at sites of increased vascular permeability (which occurs naturally at the sites of infections, inflammations, tumors, and other traumatic conditions, or it can be induced artificially) on systemic administration and in the peritoneal cavity and in the centralnervous system on localized administration. This was also the case in nontargeted Stealth liposomes, which represent the majority of applications and will be discussed below. A remaining problem is the delivery of the encapsulated drug into the cell, because the majority of cells are not phagocytic and fusion of liposomes and cells is a very rare phenomenon. One approach to overcome this difficulty is to promote fusion by surface-attached fusogenic proteins or polymers, whereas a simpler way is to try to induce leakage of liposomes by changes in temperature or pH or toprepare liposomes with compromised stability. Although this is one of the current hot issues in liposome research, several applications are not affected by the apparent lack of drug bioavailability (see below). These applications include the anthracycline anticancer drugs. The agents can be loaded at very high concentrations (normally 1 g drug/7.5 g lipid; see Fig. 9) and have shown great improvements in therapeuticefficacy on encapsulation into Stealthliposomes. According to current understanding [l,6], the prolonged presence of liposomes in blood allows them to extravasate intosites where the vasculature is leaky. This often happensto be in tumors [ll],and indeed experiments in animals and humans have shown increases in drug concentration in tumors as compared with the administration of the free drug [40-471. In thesecases, it seems that liposomes are trapped in the tissue where they slowly release their contents either by leakage or on degradation by enzymes, such as phospholipases. This coincides with the observation that peak druglevels in tumors are observed 3-4 days after liposome administration as compared with a few hours for the free

Stealth Liposomes

321

Fig. 9. Cryoelectron micrograph of unilamellar vesicles with encapsulated drug. Doxorubicin can be encapsulated at very high lipid concentrations using various gradients. In our case, we have used ammonium sulfate gradient in which weak base NH, with pK = 9.3 exchanges with weak-base doxorubicin HCl, pK = 8.3. When doxorubicin enters liposome its charged form precipitates with sulfate and of doxorubicin until most this molecular sink keeps the chemical potenial gradient of the drug is precipitated. This results in very high concentrations the liposome in interior,andthedrugsolidificationgivesrise to peculiarchanges of liposome shapes. (Micrograph: Courtesy of P. Frederik, University of Limburg, Maastricht).

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drug [40]. At the peak level, the concentration of the drug can be 10-30 times superior in the case of Stealth liposome-encapsulated drug [ll]. Fig. 10 shows the effect of various treatments on the tumor size ina C26 solid tumor mouse model,which is practicallyinsensitive to treatmentwith free adriamycin and treatments with conventional liposomes containing the drug [41]. Placebo liposomes, and their mixture with free drug, ascontrols did not have anysignificant effect over the untreated(i.e., saline-injected) animals. The administration of Stealth liposomes containing Adriamycin (SDox), however, resulted in complete remission of tumors in the case of early treatment and in significant improvements in the delayed treatment (Fig. 10). Practically identical results were obtained with the very similar drug epirubicin [42]. Improvements were also achieved in other resistant tumor models, such as mammary carcinoma. The S-Dox formulation was substantially more effective not only in curing mice with recent implants from various tumors but also in reducing the incidence of metastases originating from these intramammaryimplants [43], as can be seen fromFig. 11. Arrest of the growth of human lung tumor cells wasachieved with the S-Dox formulation [44]. The datashow that atequivalent doses, S-Dox can arrest the growth, whereas free drug and indrug conventional liposomes do 1.1 cm3/week in not. Both only decrease the size growth from approximately untreated animals to approximately 0.5 cm3/week, which is in agreement with previous observations that conventional liposomes are effective mostly in the treatmentof experimental liver metastasis. Treatment with S-Dox also shows a linear dose response[M]. One hundredpercent of mice survivedto week 12 when the formulation was dosed at2 mg/kg weeklyfor 6 weeks. The mice appeared healthy and active with minimalbody weight losses (2-19%) (Fig. 12). The same extravasation mechanism of small liposomes at sites of leaky vasculature also can beused intreatment of inflammations andinfections, which are also characterized by enhanced vascular permeability [3,10]. For instance, 10-fold improvements in the liposome concentration lung infected with Klebsiellapneumoniae [49]. Because many fungal, bacterial, and viral infections do not reside in the liver andspleen, longcirculating liposomes loaded with antifungals, antivirals, and antibiotics may also improve the treatment of some disseminated infections [3]. Furthermore, since it seems that many Stealth liposomes end up in the skin, deep tissue macrophages, and lymph nodes, it is possible that they can eradicate parasites in these sites, which are very difficult to target, and on the other hand, often harbor infections, possibly including human immunodeficiency virus (HIV) andMycobacterium avium. '

Stealth Liposomes

C

c

0

L

-0 -

s

c

a

L

lL

ti

c

x

Y)

9 a

323

324

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325

0 1 2 3 4 5 6 7 8 9 1 0 WEEKS

n

0

.

6.0

g 5.0

-

W

-

v

E

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0

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WEEKS

6.0

-

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5 6 7

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9101112

WEEKS

Fig. 12. Growth ofhumanlungtumorcancerin SCID micewhichwere engrafted with 2 million "L-1 cells and given weekly injections of nothing (O), doxorubicin in conventional liposomes (A),doxorubicin in Stealth liposomes(m). Open squaresrepresentdelayedtreatmentwithStealthliposomescontaining doxorubicin. The two experiments were performed 10 with and 5 animals (A and B, respectively). The fractionof surviving miceafter the treatment is shown in parentheses. (C) The same treatment as in (A) and (B) for the determinationof a dose response: from top to bottom: untreated mice (O), 0.5 mgkg 1 mgkg (A), and 2 mgkg (0)body weight.

(e),

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Although many new drugs are being currently monitored and tested for encapsulation in sterically stabilized liposomes, phase I and 11 clinical trials of adriamycin encapsulated in Stealth liposomes in humans have yielded very encouraging results. It was found that blood circulation times in humans, as measured by the plasma concentration of the drug, are around 45 hs. This results in greatly reduced bone marrow supression, neutropenia, and leukopenia, as well asshifted biodistribution of doxorubicin versus tumors as observed in AIDS patients with Kaposi’s sarcoma [3,11,50,51]. Unfortunately, but notunexpectedly, the altered (and unnatural) biodistribution may give rise to unusual toxicities. In this case, the dose-limiting factor seems to be stomatitis, which was observed in some patients [1l]. Another possibility of long circulating liposomes is their use as a blood substitute. Several hemoglobin-based red blood cell substitutes with desirable physicochemical properties are tootoxic and not safe(hemostasis and thrombosis) forin vivo applications. It was shownthat sterically stabilized liposomes with encapsulated hemoglobin showed a significant decrease in immunotoxicity [52]. Future developments include targeting of Stealth liposomes, spleen targeting by large sterically stabilized liposomes, long circulating microreservoirs or platforms fortherapetic activity (prevention of septic shock), and potential development of oral and topical liposome formulations. In conclusion, polymer-grafted liposomes and their applications show an interplay of various sciences, from theoretical polymer physics, colloid and interface science, organic chemistry, to biology, immunology,pharmacology, medicine, anatomy, and oncology [3,10]. As a result, very stable liposomes in biologicalenvironmentswere designed and showed remarkable therapeutic potential.The next goals are now to improve spatial and temporal control of the release of the encapsulated drug. Targeting can be improved in physically accessiblesites by the attachmentof antibodies to the of the drugwill end of polymer chains. Better controlof the temporal release be tackled by three approaches:physical approaches use temperature, pH, or (target) binding-sensitiveinduced phase change; chemical approaches try to destabilize liposome at a particular location by a chemical change of membrane-forming lipid; whereas biological approaches can use surfaceattached fusogenic proteins and polymers or stimulate internalization. We dare to conclude that sterically stabilized vesicles are an example of how complex and elusive problems can be solved by a multidisciplinary approach while for the continuation of this development even broader interdisciplinary work will be required. I believe that other polymer-liposome systems will

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be developed inwhichpolymerswillhaveadifferentfunction,suchas inducing specific binding, permeability,or fusion characteristics. REFERENCES 1. A. D. Bangham, and R.W. Home, J. Mol. Biol., 8:660-668 (1964). 2. D. Papahadjopoulos ed., Ann. N.Y. Acad. Sci., 308:l-412 3. D. D. Lasic, Liposomes: From Physics to Applications, Elsevier, Amsterdam, 1993. 4. 0.Braun Falco, H. C. Korting, H. I. Maibach eds., Liposome Dermatics, Springer-Verlag, 1992. 5. D.Papahadjopoulos, et al.,Proc.Natl.Acad.Sci.USA,88:11460-11464 (1991). 6. D.D.Lasic, F. J. Martin,A.Gabizon, et al.,Biochim.Biophys.Acta, 1070:187-192 (1991). 7. M. C. Woodle andD. D. Lasic, Biochim. Biophys. Acta, 1113:171-199 (1992) 8. L. Huang (ed.),J. Liposome Res., 57:109-430 (1992). 9. T.M. Allen, Adv. Drug Del. Rev. (in press). 10. D. D. Lasic, Angew. Chemie Int. Ed. Engl., 33:1685-1699 (1994). 11. D. D. Lasic andF. Martin eds., Stealth Liposomes, CRC Press, Boca Raton, FL, 1995. 12. P. Mukherjee, personal communication. 13. W. Helfrich, Zeit. Naturforsch., 28c:693-703 (1973). 14. D. D. Lasic, Biochim. Biophys. Acta, 592501-502 (1982). 15. D. D. Lasic, Nature,35279 (1992). 16. K.H.Klotz,M.Winterhalter,andR.Benz,Biochim.Biophys.Acta, 11471161-164 (1993). 17. D. Needham and R. Nunn, Biophys.J., 58:997-1009 (1990). 18. C. Santaella,P. Vierling, andJ. G. Riess, Angew. Chem., 30567-568 (1991). 19. M. Grit, J. H. deSmit, A. Struijke, and D. J. A. Crommelin, Int. J. Pharm., 50~1-6 (1989). 20. M. Grit and D.J. A. Crommelin, Chem. Phys. Lip., 62:113-122 (1992). 21. F. J. Martin, inSpecializedDrugDeliverySystems (E. v l e , ed.), Marcel Dekker, New York, 1990, pp. 267-316. 22. E. W. Kaler,A.K.Murthy,B. E. Rodriguez,and J. A. N.Zaszadinski, Science, 245:1371-1374 (1989). 23. S. Nir, J. Bentz, J. Wilschut,and N. Duzgunes, Prog. Surf. Sci., 13:l-124 (1983). 24. S. Okhi, D. Doyle, S. Hui, and E. Mayhuew eds., Molecular Mechanism of Membrane Fusion, Plenum Press, New York, 1988. 25. J. N. Israelachvili, Intermolecular and Surface Forces, Academic Press, New York, 1985. 26. M. C. Woodle, et al., Biochim. Biophys. Acta, 1105:193-2OOO (1992). 27. P. G. deGennes, Adv. Colloid. Interface Sci., 27:189-209 (1987).

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28. P. G. deGennes, Personal communication, 1992. 29. K. Hristova and D. Needham, in ref.11, chapter 5,35-50. 30. S. ZalipskyandC.Lee,inPoly(ethy1ene)GlycolChemistry, J. M. Hams (ed.), Plenum Press,New York, 1992, pp. 347-370. 31. S. Zalipsky, in ref.11, chapter 9,93-102. 32. S. Zalipsky, Bioconjugate Chem., 4:296-301 (1993). 33. M. C. Woodle, C. Engbergs, andS. Zalipsky Bioconjugate Chem., 5:493-497 (1994). 34. V.P. Torchilin, et al., Biochim. Biophys. Acta 1195,181-185, 1994. 35. D. Needham, T. J. McIntosh,andD. D. Lasic,Biochim.Biophys.Acta, 1108:40-48 (1992). 36. D. Needham,K, Hristova, T.J.McIntosh, et al., J. Liposome Res.;2:411-430 (1992). J. N. Israelachvili,Biophys. J., 37. T.Kuhl, D. Leckband,D.D.Lasic,and 66~1479-1488 (1994). 38. T. R. Baekmark, G. Elander, D. D. Lasic, and E. Sackmann, Laugmuir (in press). 39. H. F. Dvorak, A. J.Nagy, J. T. Dvorak, and A. M. Dvorak, Am. J. Pathol., 133:95-108 (1988). 40. A. Gabizon, CancerRes. 52:891-896 (1992). 41. E. Mayhew, D. D. Lasic, S. Babbar, and F. J. Martin, Int. J. Cancer, 51:302309 (1992). 42. S. K. Huang, et al., Cancer Res., 52:6774-6781 (1992). 43. J. Vaage, E. Mayhew, D. D. Lasic, and F. J. Martin, Int. J. Cancer, 51:942948 (1992). 44. S. Williams, et al., Cancer Res., 53:3964 (1993). 45. P. Working, et al., J. Liposome Res., 4:667 (1994). 46. J. Vaage and E. Barbera, in ref.11, chapter 14,149-172. 47. D. D. Lasic, in ref11, chapter 13,139-148. 48. T. M. Allen, in ref 11, chapter 16,187-196. 49. I. M. K. Baker-Woudenberg and M. C. Woodle,J. Infect. Dis. (in press). 50. J. Boggner andF. Goebel, in ref. 11, chapter 23,267-278. 51. D. Nortfeld,et al. in ref 11, chapter 22,257-268. 52. R. Beissinger, et al., J. Liposome Res., 3575 (1993). 53. D. D. Lasic, Amer. Sci., 80:20-31 (1992). 54. G. R. Haran, R. Cohen, L. K. Bar, and Y.Barenholz, Biochim. Biophys. Acta, 1151:201 (1993). 55. D. D. Lasic, et al., FEBS Lett., 312:255-259 (1992).

12 Nonionic Surfactant Vesicles Containing Estradiol for Topical Application Don A. van Hal, JokeA. Bouwstra, and Hans E. Junginger LeidentAmsterdam Center forDrug Research, Leiden University, Leiden, The Netherlands

I. Introduction 11. Nonionic Surfactant Vesicles A. Preparation B. Physical State C. DrugEntrapment

330 330 331 333 333

111. Liposomes, Micelles, and Transferosomes

333

IV. Transdermal Application of Vesicles

334

V. Interactions Between Vesicles and Skin A. Interactions Between NSVs and Skin 1. Transport Studies 2. Thermodynamic Activity 3. Penetration Enhancement by Surfactants 4. Increase in Drug Transport by Vesicles 5 . Physical State

VI. Summary

335 336 336 337 337 338 338 339

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330 1.

INTRODUCTION

About 17 years ago, vesicular systems were added to the armamentarium for delivering drugs to or through the skin: nonionic surfactant vesicles (NSVs) [l] and lipid vesicles (liposomes) [2]. In this chapter, the term vesicles stands for both liposomes as well as NSVs. The application of vesicular systems in cosmeticsand for therapeutic purposes may offer several advantages: The vesicle suspension is a water-based vehicle. This offers high patient compliance in comparison with oily dosage forms. The vesicles allowthe incorporation of hydrophilic, amphiphilic, and lipophilic drugs. The vesicles supply both a lipophilic and an aqueous environment for the encapsulation of drugs. The characteristics of the vesicle formulation are variable and controllable. The vesicle characteristics can be controlled by altering vesicle composition, size, lamellarity, trapped volume, surface charge, and concentration. Biocompatible and biodegradablesystems maybe designed. The vesicles may act as a depot, releasing the drug in a controlled manner. The vesicles may protect the drug from degradation. Because liposomes consist mainly of phospholipids, and NSVs are mainly prepared fromnonionic surfactants. Differences in characteristics exist, between liposomes and NSVs, especially since NSVs are prepared from unchanged single-chain surfactants and cholesterol, whereas liposomes are prepared from double-chain phospholipids (neutral or charged). One of the advantages in using NSVs is the higher chemical stability, lower costs of the chemicals, and the large number of classes of surfactants available to design these vesicles on demand.

II. NONIONICSURFACTANTVESICLES NSVs consist of one or more nonionic surfactant bilayers (Fig. 1) enclosing an aqueous space. NSVs consisting of one bilayer are designated as small unilamellar vesicles (SUVs) or large unilamellar vesicles (LUVs). Vesicles with more bilayers are called multilamellar vesicles (MLVs). Since1975[3]vesicleshave been prepared from synthetic surfactants. Various synthetic surfactants that have been used for the preparation of vesicles are listed in Table 1. For vesicles prepared from nonionic surfactants, we prefer to use theterm nonionic surfactant vesicles, or

Estradiol-containing NSVs

33 1 ..

.

.

Fig. 1 Structure ofnonionic surfactant aggregates: L W (left), micelle (middle), MLV (right).

NSVs. The NSVs are alreadyused in cosmeticsby L'Oreal (Paris, France), and they are still being tested for their therapeutic potential. In our laboratory NSVs, have been prepared from two groups of surfactants [22,30,31]: 1. Polyoxyethylene alkyl ether surfactants (GEO,,,),in which m'rep-

resents the number of oxyethylene units and n represents the number of carbon atoms in the hydrocarbon chain. q2EQ, C,,EC4, and G=9E0,0(Brij96) form liquid-statevesicles and G,EQ and q,EQ form gel state vesiclesoat room temperature. 2. . Sucrose ester Wasag-7. Wasag-7. consists of 70% stearate ester and 30% palmitate ester(40% mono, 60% di/tri-ester) and forms gel-state bilayers at room temperature. These surfactants were chosen because they are relatively nontoxic. .. A .thorough description of the formation and properties of these types of vesicles from nonionic surfactants is given by Hoflandet al. [22].and Van Hal [31].

A. Preparation NSVs can be formed using the same methods that are used fur the prepara. .. . tion of liposomes [32]. 1 .." Sohication Method [30,31] The lipophilic components are dissolved in a n apolar solvent,such a s a mixture of chlorofoni and methanol. The solvent evaporates overnight

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et al.

Table 1. Vesicle Preparation from Synthetic Surfactants References

Surfactant

Fatty acids Polyglycerol alkyl etheror esters and polyethoxylated analogues Alkyl glycoside surfactants Sucrose esters Polyoxyethylene alkyl ether surfactants Triethoxylated cholesterol Glyceryl dilaurateor distearate with polyoxyethylene-10-stearyl ether Crown ether surfactants Polyoxyethylene di-alkyl ether surfactants Diphenylazomethine ammonium amphiphiles

394

2,s-l7 18-20 21 22

Ufasomes Nonionic surfactant Vesicles, niosomes AGVs Lipid vesicles Nonionic surfactant Vesicles, NSVs

23

24

Novasome

25,26 27

Niosomes Bilayer membranes, vesicles Bilayer assemblies

28,29

under vacuum, resulting in a surfactant film. The film isthen hydratedwith aqueous solution and the mixture is sonicated at 80°C. The method results in the formation of both LUVs and MLVs.

2. Ether Injection Method [7] The oil-soluble components are dissolved in diethyl ether and injected intothe aqueous phase at 60°C. The method results in the formation of mainly MLVs. 3. HandshakingMethod [7] The apolar ingredients are dissolved in a mixture of chloroform and methanol. The solvent evaporates overnight under vacuum. The resulting surfactant film is hydrated with an aqueous solution and gently shaken at 50-60°C to allow the formation of vesicles. This preparation methodyields MLVs. 4. Reversed Phase Evaporation [l81 The oil-soluble components are dissolved in a apolar solvent such as chloroform. The aqueous phase is added and the mixture is sonicated to form an emulsion. The solvent evaporates overnight under vacuum. The resulting suspension is shaken to form the vesicles. Mainly MLVs are formed.

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5. Method as Described by Handjani-Vila [5] The surfactants are added to the aqueous phase and the mixture is agitated to yield a homogeneous lamellar phase. The suspension is then homogenized by means of agitation or ultracentrifugation.

B. PhysicalState The bilayers of the vesicles are either in the so-called liquid state orin gel state (Fig. 2), depending on the temperature, the type of lipid or surfactant, and the presence of other components such as cholesterol. In the gel state, thealkyl chains are present in a well-ordered structure, andin the liquid state, the structure of the bilayer is more disordered.The surfactants and lipids are characterized by the gel-liquid transition temperature, Tc [33]. In general, the action of cholesterol is twofold: On the one hand, cholesterol increases the chain order of liquid-state bilayers, and on the other, cholesterol decreases the chain order of gel state bilayers [34]. At a high cholesterol concentration, the gel state is transformed to a liquidordered phase. C.DrugEntrapment The NSVs have proven to be able to entrap several model compounds and active substances, such as carboxyfluorescein [7,22,31], sucrose [M], the DGAVP peptide [35], estradiol [30,31,36], and dithranol [31]. The entrapment efficiencies of NSVs are in the same order of magnitude as in liposomes. The exact efficiency depends on the drug, vesicle composition, and preparation method. The entrapment efficiencies and maximum capacities of lipophilic drugs decrease as the cholesterol content increases [31]. 111.

LIPOSOMES,MICELLES,ANDTRANSFERSOMES

Liposomes are vesicles consisting of various types of phospholipids [32]. These amphiphilic molecules can be neutral, ionic, or zwitterionic. Like

Gel state

Liquid state

Fig. 2 Gel state and liquid state of bilayers without cholesterol.

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et al.

NSVs, they consist of bilayers,enclosing-anaqueous core and are capable of encapsulating lipophilic, hydrophylic, and amphiphilic drugs. Liposomes can vary in sizebetween 50 nm and several micrometers., A micelle (see Fig. 1) is another surfactant aggregate that is able to entrap lipophilic molecules in an aqueousmedium. They are much smaller than liposomes andcontain no aqueous coreor lipid bilayers. Transfersomes [37-411 consist of a mixture of a lipidic agent with a surfactant. The bilayers of transfersomes are much more elastic than those of most liposomes. A transfersome is one or two orders of magnitude greater in size than mixed-lipid micelles. Furthermore, transfersomes contain an aqueous core enclosed by bilayers. Although lipophilic and amphiphilic drugs may be incorporated in transfersomes as well, their unique potential is to act as carrier for encapsulated.hydrophi1icdrugs such as, for example, insulin [39]. IV. TRANSDERMAL APPLICATION OF VESICLES The effectiveness of vesicles has been investigated by several research groups (Table 2). Liposomes, in particular, have received a great deal of attention. In several studies the diffusion of a drug was facilitated or achieved certain selectivity into humanor nonhuman skin by vesicleencapsulation. Others studies show that theinfluence of liposomes on drug trans? port is negligible. Our group has investigated the potential of NSVs to increase .the diffusion of estradiol throughhumanstratumcorneum [30,31,36,42,43]. Table 2. Effect of Liposomes and NSVs on the Permeation of Drugs Through in the Skin inVitro and Vivo .. . References

Effect Liposomes No or decreased effect of vesicles Superiority of

liposomes

Transfersomes Superiority of transfersomes NSVs Superiority of NSVs

In vitro In vivo animal In vivo human vitro In studies In vivo animal In vivo human

3,649 50-52 53,s ' 24,56-67 1,56,58,68-82 79,83-92

In vivo animal

38, .39

In vitro' human

31,36,42

,

':

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There are three reasons for the controversy found in the literature data with regard to theeffects that vesicular formulation may achieve: The first reason is the natureof the control formulationin the diffusion studies. The effect of the liposome formulation was compared with aqueous solutions of drug or emulsions, ointments, creams, gels, or suspensions. The second reasonis thefact thatvarious drugs have been used,such as steroid hormones, glucocorticoids, anti-infectives, local anesthetics, retinoids, and minoxidil [M]. NSVs have been studied for the topical administration of pyrrolidone-carboxylate [2]and estradiol [36]. A thirdreason is that liposome formulations with completely different chemical and physical properties have been tested. All of these factors influence the thermodynamic activity of the drug in the formulation, a parameter which determines the driving force for thedrug to diffuse from the formulation to the skin [45]. The thermodynamic activities of the drugs in the tested vesicle . . formulations are therefore very likely to differ. V. INTERACTIONS BETWEEN VESICLES AND SKIN

Increased drug transport through the skin may be achieved by specific interactions between vesicles and skin. Because the vesicles may alter the structure of the skin, a thorough knowledge of the “normal” structure of the skin is important. The vesicles may alter the structure of the skin in such a way that the diffusion of the drug is facilitated. The interaction can be influenced by the chemical characteristics of the vesicles, such as .the amount and type of surfactant,membranestabilizer, or charged compounds. The physical characteristics include the physical state (gel or liquid state) of the bilayers, the number of bilayers, or the size of the vesicles. The interaction may also be influenced by skin characteristics: hydration of theskin, sweat production, skin microflora, sebum,and cellular debris. Three mechanisms bywhich vesicle-associated effects occur have been described so far: (1) penetration of intact vesicles, (2) adsorption of vesicles onto the skin surface, and (3) penetration of vesicle constituents into the skin (Table 3). The so-called transfersomes are also listed in Table 3. Cevc reports [39] ‘that these .vesicles areelastic enough to move through the constrictions of the skin.. .The transfersomes are thought to exhibit their action owing to “hydration forces and osmotic gradients” [37,38]..It is claimed that drugs which are applied by transfersomes are able to reach an almost 100% maximum biological or therapeutic effect, which is comparable to . injectioil [39]. ’

.

a

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Table 3. Overview of the Reported Mechanisms on Vesicle Action Mechanism Penetration of intact vesicles 1. Into the stratum corneum and into the deeper layers of the skin The vesiclesare considered to cross the biologicalbamer and may even target their constituents to specific sites Adsorption of vesicles to the surface of the skin This hypothesis comprises the idea that vesicles adsorb to the surface of the skin. This would: 1. Allow a direct transfer of the drug to the stratum corneum 2. Induce an occlusive effect 3. Result in a gradient of thermodynamic activity Penetration of vesicle constituents This hypothesis comprises the disruption of the vesicles and penetration of molecularly dispersed material or bilayer fragments into the stratum corneum.

A.

VesicletReferences Liposomes Pro: 1,78,90,93 Transfersomes Pro: 38-41

Liposomes Pro: 47,50,55,62,67,73,87, 94-97 NSVs Pro: 31,42,43

Liposomes Pro: 55-57,59,60,63,65,67,68, 72,87,88,95,97,.98-103 NSVs Pro: 31,42,43

Interactions Between NSVs and Skin

l . Transport Studies Our transportstudies using estradiol [30,31,36] have revealedthat the effect that NSVs have on the diffusion of the drug through the stratum corneum in vitro depends on the vesicle composition and the thermodynamic activityof the drug. Incorporation of estradiol ingel-state Wasag-7:CHOL:DCP (CHOL, cholesterol; DCP,dicetylphosphate),Wasag-7:CHOL:DCP, and C,,Eq:

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CHOL NSVs resulted in an increase in estradiol steady-stateflux by a factor of 2.2,2.4, and1.5, respectively [31]. NSVs of G,EQ did not have a significant effect on the estradiol penetrationas compared with the control [36]. The steady-state flux could be increased by a factorof 4.8 by incorporation of estradiol intoliquid-state C,,EO,:CHOL (1:0.43) NSVs [31]. Decreasing the CHOWSURFmolar ratio of G,EO,:CHOL from 0.43 to 0.18 resulted in an increase in steady-state flux. Compared with the control, the flux was increased by a factor of 9.5. The flux increase was even higher in liquid-state Brij96:CHOLand G,EO,:CHOL vesicles, being a factor ofapproximately 40, respectively [36].

2. ThermodynamicActivity The first important parameter that influences the penetrationof a drug is the thermodynamic activity of the drug in the formulation, which is the driving force fordiffusion of the drug into theskin. This so-called push-effect [45] isequal in samples at saturationlevel, where the thermodynamic activity reaches a value of 1. In order to check the differences in thermodynamic activity between the vesicle suspensions and the control solution (estradiolsaturated phosphate buffer saline (PBS)), “reversed membrane’’ experiments were carried ou$ The stratum corneum and Silastic sheeting were mounted intodiffusion cells with the Silasticsheeting facing the donorchamber. The experiments thatwere carried outwith liquid-state G,EQ:CHOL (1:0.43) vesicles and with gel-state Wasag-7:CHOL:DCP vesicles showed that thediffusion of estradiol by means of NSV suspensions was a factor1.82.2 higher than thediffusion of estradiol applied in estradiol-saturated PBS [31]. A possible explanation for the results is a higher estradiol thermodynamic activity (push-effect) in the NSV suspensions as compared with the PBS solution. This may indicate that theNSV suspensions were supersaturated. This corresponds to the meta-stable state of estradiol incorporation [311. 3. Penetration Enhancement by Surfactants In drug transport studies, one has to consider a second mechanism whichis associated with thesurfactants of the vesicles. Animportant mechanism that may be responsible for this increase in steady-state flux is that theliquid state G,EO,:CHOL [31], G,EO,:CHOL, and Brij96:CHOL vesicles [36]also appeared to have a penetration enhancement effect, which is called the pull-effect [45]. This was confirmed by pretreatment experiments. The stratum corneum was pretreated with empty NSVs prior to application of the drug control solution [31,36]. Pretreatment with liquidstate G,EQ:CHOL (1:0.18) vesicles resulted in an increase of the steadystate estradiol flux by a factor of 2.9 [31], which is clearly lower than the

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factor 9.5 that could be achieved by application of estradiol-loaded vesicles. Pretreatment with liquid-state G,EQ:CHOL and Brij96:CHOL also increased the estradiol flux, but with a clearly lower factor, as compared with the- estradiol-loaded formulations ,[31]. Pretreatment with gel-state G,EQ:CHOL vesicles revealed that.gel-state surfactants did not increase the estradiol flux [36]. The increase in f l t q induced bythe liquid-state G=,EO,,,:CHOL vesicles is partly caused by4he action of the surfactants with skin lipids of the stratum corneum. The surfactants can act as penetration enhancers. The addition of estradiol in G=,EO, micelles increased the estradiol flux by a factor of 2 [36]. Probably the G=,,EQ, micelles are able to penetrate molecularly dispersed stratum corneum andlower the barrier function. 4. Increase in Drug Transport by the Vesicles . However,'for the G,EQ/CHOL(1.:0.18) formulation, the penetiation of surfactants is not a valid explanation, as could be deduced frdm ,an experiment that involved.the diffusion of estradiol from G,EQ micelles. The addition of estradiol to this formulation had no effect on the diffusion throughstratum ,corneu'm [31]. A. change in thestructure of stratum corneum incubated with G,EQ:CHOL NSVs could only occasionally be visualized using freeze-fracture electron microscopy (FFEM) when' compared with stratum corneum. that was treated with PBS for 48 h '[31]. However, the observed minor changes in stratum corneum structure cannot explain the increased transport rate of estradiol. The vesicular form of G,E07, which is induced' by the addition of a small amount of cholesterol, appears to be crucial for obtaining a strong increase in estradiol flux. This is also confirmed by the fact that occlusive application of estradiol in a G,EO,:CHOL formulation in phosphatebuffer (PB), which probably contains micelles instead of vesicles, had noeffect on the flux as comparedwith the control situation [31]. To achieve a maximum effect, estradiol has to be associated with the vesicles. The importance of encapsulation of estradiol in NSVs is confirmed ,by the results of Hofland et al. [36]. The penetration,of bilayer constituents into the stratum corneum probably facilitates the simultaneousdiffusion of estradiol when estradiol is . . . associated these to bilayers. . . . 5. Physical State The physical state of the NSVs is another important parameter n i flux enhancement. The results showed that liquid-state .vesicles could.strongly increase the flux [31,36], whereas gel-state vesicles resulted in only a slight flux increase [31] or nomcrease at all [36]. The flexibility.of the' bilayers is decreased by an increase in CHOW .SURF molar ratio andby the use of surfactants forming gel-state bilayers. '

'

,

'

,

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The fact that an increased cholesterol content of C;,EQ:CHOL vesicles decreased the steady-state flux of estradiol confirms this hypothesis. Furthermore, the results show that the increase in estradiol flux issmaller after the application of gel-state bilayers compared with liquid state vesicles [31,36]. The fact that the gel-state vesicles from Wasag-7:CHOL:DCP exert almost the same steady-stateflux in the reversed membrane experimentas in the transport studies [31] shows that the main mechanism behind the steady-state flux increase concerning gel-state vesicles is achieved through differences in thermodynamic activity. These vesicles, and probably also the vesicles from the other gel-state vesicles Wasag-15:CHOL:DCP and C,,EO,:CHOL, do not lead to an increase in drug transport when a direct contact between vesicles and skin occurs. The FFEM results showed that the formation of bilayer stacks occasionally plays a role after occlusive application of NSVs [31,42]. We concluded that the formation of bilayers only occasionally explains. the increased fluxes, especially in the case of liquid-state vesicles. Bilayer sheet formation plays a major role after nonocclusive application of NSV suspensions [31]. The ultimate contact between the estradiol-saturated bilayer sheets and the stratum corneum may increase the flux of estradiol, possibly via three mechanisms: first, the contactallows a direct transferof drug and vesicle constituents to the stratum corneum; second, via the formation of a thermodynamic activity gradient across the stratum corneum [42]; and third, the penetration of the vesicle constituents into the stratum corneum. VI.

SUMMARY

Incorporation,of estradiolinto NSVs may lead to. increased penetrationof the drug throughhumanstratum corneum. The results have shown thatit is possible to increase the estradiol flux by a factor of 50 as compared with estradiol-saturated PBS. These results make NSVs promising for transdermal application of drugs, because with penetration enhancers such as h o n e or oleic acid in propylene glycol, onlyan enhancement factor of 1.4 or 3.6 can be reached [104].. The mechanisms by which the increase in steady state flux can be described are thestate ofsupersaturation of the vesicle suspensions and the intrinsic penetration enhancement effects of vesicle bilayers that are in the liquid state at the experimental temperature. Depending on the of surtype factant, a penetration enhancementmay be caused by the diffusion of the surfactants into the stratum corneum. Furthermore, the elasticity of the bilayers may play an important role [38,41]. Decreasing the elasticity of

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the bilayers by increasing the cholesterol contentor by applying surfactants forming gel-state bilayers results in a decrease in estradiolflux. The formation of bilayer sheets on top of the stratum corneum can only occasionally explainthe increased fluxes obtained afterocclusive application of the NSVs tested in this investigation. After nonocclusive application, largeregions of bilayer stacks couldbe visualized on the surface of the stratum corneum using freeze-fracture electron microscopy.

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74. N.Weiner,K. Egbaria, and R. Chandrasekhasan, Topical delivery of liposomally encapsulated interferon evaluated by in vitro diffusion studies and in a (0.Braun-Falco, cutaneous herpes guinea pig model, in Liposome Dermatics H. C. Korting, and H. 1. Maibach, eds.), Springer, Berlin, 1992, 242-250. 75. V. Masini, F. Bonte, A. Meybeck, and J. Wepierre, In vitro percutaneous absorption and in vivo distribution of retinoic acid in liposomes and in a gel studiedinhairlessrats,Proc.Int.Symp.Control.Rel.Bioact.Mater., 17~425-426 (1990). 76. C. Artmann, J. Roding, M. Ghyczy, andH. G. Pratzel, Liposomes from soya phospholipids as percutaneous drug carriers. Qualitative in vivo investigations with antibody-loaded liposomes, Arzneim.-Forsch./Drug Res., 40:1363-1371 (1990). 77. C. Artmann, J. Roding, M. Ghyczy, and H. G. Pratzel, Liposomes from soya phospholipids as percutaneous drug carriers. Quantitative in vivo investigationswithradioactivelylabelledliposomes,Arzneim.-Forsch,lDrugRes., 40~1365-1368 (1990). 78. D. B. Yarosh, Liposome-encapsulated enzymes for DNA repair, in Liposome I. Maibach,eds.), Dermatics (0. Braun-Falco, H. C. Korting,andH. Springer, Berlin, 1992,258-269. on per79. C.Michel,T.Purmann, E. Mentrup, et al.,Effectofliposomes cutaneous penetration of lipophilic materials,Int. J. Pharm., 84:93-105 (1992). 80. C. Michel, N. Groth, T. Herrling, et al., Penetrationof spin-labeled retinoic acid from liposomal preparations into the skin ofSKHl hairless mice. Measurement by EPR tomography, Int.J. Pharm., 98:131-139 (1993). 81. H. E. BoddC,L.A. R.-M. Pechtold,M. T. A. Subnel, and F. H. N. De Haanpechtold subnel haan Monitoring in vivo skin hydration by liposomes using infrared spectroscopy in conjunction with tape stripping in Liposome Dermatics (0. Braun-FalcoH.C.Korting,andH.I.Maibach,eds.), Springer, Berlin, 1992, pp. 137-149. 82. L.M. Lieb, C. Ramachandran, K. Egbaria, and N. Weiner,Topicaldrug delivery enhancement with multilamellar liposomes into pilosebaceous units: I. In vitro evaluation using fluorescent techniques with the hamster ear model, J. Invest. Dermatol.,99:108-113 (1992). der per83. L.Kr6wczynskiand T. Stozek,LiposomenalsWirkstoffragerin kutanen Therapie, Pharmazie, 39:627-629 (1984). 8 4 . W. Raab,Liposomen-eineneueFormdermatologischerWirkstoffrager, Artzl. Kosmetol., 18:213-224 (1988). skin byliposome85 A.GesztesandM.Mezei,Topicalanesthesiaofthe encapsulated tetracaine, Anesth. Analg., 67:1079-1081 (1988). 86 R. A. Dudley, P. Edwards, R. P. Ekins, et al., Guidelines for immunoassay data processing,Clin. Chem., 31:1264-1271 (1985). 87. M. Jacobs, G. P. Martin, and C. Marriott, Effects of phosphatidylcholine in the topical bioavailability of corticosteroids assessed the human by skin blanching assay, J. Pharm. Pharmacol., 40:829-833 (1988). 88. W. Gehring, M.Ghyczy,M. Gloor, et al., Significance of empty liposomes I

I

346

89.

90, 91.

92. 93. 94. 95. 96.

97. 98. 99.

100. 101. 102. 103. 104.

van Hal et al. aloneandasdrugcarriersindermatology,Ameim.-Forsch./DrugRes., 40~1368-1371(1990). W. Gehring, M. Klein, and M. Gloor, Influence of various topical liposome preparations with and without active ingredients on the cutaneous blood flow,inLiposomeDermatics (0.Braun-Falco, H. C. Korting, and H. I. Maibach, eds.), Springer, Berlin, 1992, pp. 315-319. M. Foldvari, A. Gesztes, andM. Mezei, Dermal drug delivery by liposome encapsulation: Clinical and electron microscopic studies, J. Microencaps., 7:479-489 (1990). C. J. N. Oguefior,Topicaldosageform of M. Foldvari,B.Jarvis,and liposomal tetracaine: Effectof additives on'the in vitro release and in vivo efficacy, J. Control. Rel.,27:193-205 (1993). H. C. Korting, Increased activity and tolerability of a conventional glucocorticoid in a liposomal form, in Liposome Dermatics(0. BraunPalco, H. C. Korting, and H. I. Maibach, eds.), Springer, Berlin, 1992, pp. ,320-328. G. Lasch, R. Laub, and W. Wohlrab, How deep penetrate intact liposomes into human skin?J. Control. Rel., 18:55-58, 1991. M. G. Ganesan, N. D. Weiner, G. L. Flynn, and N. F. H. Ho, Influence of liposomaldrugentrapment on percutaneousabsorption,Int. J.Pharm., 20:139-154 (1984). Siciliano, 1985. Jessberger, 1988; R. Schubert, M. Joos, M.Deicher, et al., Destabilization ofegglecithin liposomes on the skin after topical application measured by perturbed TT angular correlation spectroscopy (PAC) with "'In, Biochim. Biophys. Acta, 1150~162-164 (1993). M. B. Fawzi, U. R. Lyer, and M:Mahjour, U.S. Patent 4,783,450,1988. W. Wohlrab, J. Lasch, R. Laub, et al., Distribution of liposome-encapsulated ingredients in human skin ex vivo, in Liposome Dermatics (0.Braun-Falco, H. C. Korting, and H. I. Maibach, eds.), Springer, 'Berlin, 1992, 215-225. M:Mahjour, B. Mauser,Z. Rashidbaigi, andM:B. Fawzi, Effect of egg yolk lecithins and commercial soybean lecithins on in vitro skin permeation of drugs, J. Control. Rel., 14:243-252 (1990). N. Weiner, Topical delivery of liposomally encapsulated ingreK. Egbaria and dients evaluated, in Liposome Dermatics (0.Braun-Falco, H. C. Korting, and H. I. Maibach, eds.), Springer, Berlin, 1992, pp. 172-181. J.RodingandC.Artmann,Thefateofliposomesinanimal skin, in Liposome Dermatics(0. Braun-Falco, H. C. Korting, H.I. Maibach, eds.), Springer, Berlin, 1992, pp. 110-117. on the M.Ghyczy,Chemicalcompositionofliposomesanditsinfluence humidity of normal skin, in Liposome Dermatics (0..Braun-Falco, H. C. Korting, and H. 1: Maibach, eds.), Springer, Berlin, 1992, pp. 308-314. M. Goodman and B. W. Barry, Action of penetration enhancerson human skin asassessedby thepermeation ofmodeldrugs5-fluorouraciland estradiol. I. Infinite dose technique, J. Invest. Dermatol., 91:323-327 (1988).

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ADDITIONAL READINGS

A. H. Armin, A. T. Florence, R. M. Handjani-Vila, et al., The effectof niosomes and polysorbate 80on the metabolism and excretion of methotrexatethe inmouse, J. Microencaps., 3:95-100 (1986). G. L. Flynn, S. H. Yalkowsky and T. J. Roseman, Mass transport phenomena and models: theoretical concepts,J. Pharm. Sci., 63:479-510 (1974). R. H. Guy and J. Hadgraft, Mathematical modelsof percutaneous absorption, in Percutaneous Absorption. Mechanisms, Methodology, Drug Delivery(R. L. BroMaibach, eds.), Marcel Dekker, New York, 1985, pp.5-15. naugh and H.I. D. J. Kerr, A. Rogerson, G . J. Morrison, et al., Antitumour activity and pharmacokinetics of niosome encapsulated adriamycin in monolayer, spheroid and xenograft, Br. J. Cancer, 58:432-436 (1988). L. Khand, A. Rogerson, G. W. Halbert, et al., The effect of cholesterol on the release of doxorubicin from non-ionic surfactant vesicles, J. Pharm. Pharmacol 39(Suppl.):41P (1987). M. Mezei, Multiphase liposomal drug delivery system, U.S. Patent 4,761,288,1988. L. K. Pershing, L. D. Lambert, and K. Knutson, Mechanism of ethanol-enhanced estradiol permeation across human skin in vivo, Pharm. Res., 7:170-175 (1990). A. Rogerson, J. Cummings, N. Willmott, and A. T. Florence. The distribution of doxorubicin in mice following administration in niosomes, J. Pharm. Pharmacol., 40~337-342 (1988). S. Stafford, A.J. Baillie, and A.T. Florence, Drug effects on the sizeof chemically J. Pharm. Pharmacol., 4O(Suppl.):26P (1988). defined non-ionic surfactant vesicles, U. Tauber, Drug metabolism in the skin: advantages and disadvantages, in Transdermal Drug Delivery (J. Hadgraft, and R. H. Guy, eds.), Marcel Dekker, New York, 1989, pp. 99-112.

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13 Microcapsules: Preparationby Interfacial Polymerization and Interfacial Complexation and Their Applications Tony L. Whateley University ofStrathclyde, Glasgow, Scotland

I. Introduction

11. Interfacial Polymerization A. Semipermeable, Aqueous Core, and Polyamide Microcapsules B. Nonbiomedical Aspects of Microcapsules with Rigid, Nonpermeable Shells and Liquid Cores

349 349

351 363

111. Interfacial Addition Polymerization A. Alkylcyanoacrylates

364 364

IV. Interfacial Complexation or Interfacial Polyelectrolyte Complexation A. Alginate/PolylysineMicrocapsules

366 366

1.

INTRODUCTION

The term interfuciulpolymerization is usedhere toinclude interfacial addition polymerization and interfacial condensation polymerization. However, interfacial complexation is a separate process and is generally applied to produce microcapsules with somewhat different properties and specialized applications from those produced from theinterfacial polymerization processes, and these topics will be considered separately. 349

350

Whateley

MICROSPHERES Monolithic Uniformdistribution of particlesor a molecular dispersion

.g:;:.:

0 .....:..-.'..

y:...

MICROCAPSULES Liquid contehts, nonpetmeable,rigid.. . membrane . . .

I

0

TYPE 2

.

Aqueous contents, semipetmeable membrane

,

I"\

'.,I

TYPE I

Solid core, protectiveor releasecontrolling coating

Fig. 1 'Qpes of microspheres and microcapsules. In general, all of the above processes produce liquid-cored (both aqueous and nonaqueous) microcapsules. Those microcapsules with semipermeable membranes and aqueous cores (type 1 in Fig. 1) are of special interest in the biomedical and pharmaceutical areas. The various types of microcapsules and microspheres are illustrated in Figure 1. The types of microcapsules produced by the processes to be considered in this chapter are types 1 and 2 (in general, only type 1is produced by interfacial complexation). Thus, we are considering microcapsules with liquid cores and either a igid, brittle, nonpermeable shell (type 2) or a semipermeable membraneshell (type 1): The preparative methodsfor both types have much in common. However, the applications are very different: i t is those with aqueous cores and semipermeable membranes that have most application in the pharmaceutical and biomedical fields. Interfacial polymerization 'and interfacial condensation polymerization are so similar that the topics will be considered together. Interfacial complexation (or interfacial polyelectrolyte complexation) tends to be mainly used to produce a particular type of microcapsule (biocompatible, aqueouscore with a semipermeable membrane) andits prepa. .. ration and applications will be considered separately. Interfacial addition polymerization has been of interest in the medical, biomedical, and Pharmaceutical fields because of the alkylcyanoacylates and their ability to polymerize readily. However, in general, the type of microcapsule (or probably more precisely, microsphere) produced solid, monolithic variety. This topic will also be treated separately. .'is of the An overview of drug delivery by various microencapsulated systems isgivenby Arshady [l]. Systems prepared by a wide range of micro,

,

Microcapsules Polymerization by

and Complexation

35 l

encapsulation methods are considered. Arshady [2] also reviewed the preparation of microspheresand microcapsulesby interfacial polycondensation techniques. Chang [3] reviewed recent advances. in artificial cells covering especially the biotechnology and medical applications. A number of books covering this area have been published; for example, those by Chang [3], Lim [4], Donbrow [ 5 ] , and Whateley [6]. 11.

INTERFACIALPOLYMERIZATION

A. Semipermeable, Aqueous Core, and Polyamide Microcapsules (Type 1 in Fig. 1)

This type of microcapsule was initiallydeveloped by Chang [7,8] in Canada in Japan have alsodone extensive work in the 1960s. Kondo and coworkers on these systems. Originally developed by Chang as “artificial cells,” this type of microcapsule has been used to encapsulate: Proteins Enzymes Antibodies Cells

,

.

The important features of the semipermeable encapsulating membrane are: ..

1. Not permeable to large molecules such as proteins, enzymes. 2. Permeable to small moleculessuch as enzyme substrates and products. A pore size of about 2 nm fulfillsthese objectives and can be achieved by the interfacial polymerization process. 3. Nonantigenic surface. 4. Although biodegradability is important in some applications, this .. has been difficult to achieve. ,

A range of reactive monomer-monomer combinations can be used to react at anoil-water interface, as detailed later; theinterface may be planar or at , . the perimeter of a small droplet, asshown in Fig.2. The basic feature of the method to produce microcapsules, which is the same in all’cases, is,the formation of a water-in-oil (W/O) emulsion, as illustrated in Fig. 3... Addition of the second monomer, B, to the preformed emulsion initiates the interfacial reaction. Details of preparative procedures are given later. . In this interfacial polymerization process, various combinations of monomers can be used to obtain a range of polymer membranes. Some

Whateley

352

Fig. 2 Microencapsulation by interfacial polymerization method.

Emulsion Water-in4

Fig. 3 Preparation of microcapsules by interfacial polymerization.

possibilities are illustrated in Figs. 4a-d. Figure 4a shows the polymerization between sabacoyl chloride and hexamethylene diamine (1,6-hexanediamine) to form the polyamide nylon 6,lO: This system was used in the early studies on microcapsule formation by the interfacial polymerization process. Such microcapsules, however, tend to be rather fragile and difficult to separate, wash, and handle. More robust microcapsules can be prepared using terephthalic (p-phthalic) acid dichloride with the diamine (Fig. 4b). Piperazine has frequently been used with terephthaloyl chloride to give a robust poly(terephthaloy1-piperazine) membrane. Substituted

Microcapsules Polymerization by Complexation and

353

diamines, for example, L-lysine (Fig. 4c)can be used in order to give the microcapsule membranea charged group. Other classes of polymeric membrane can be formed, for example, polyesters as shown in Fig. 4d. Polyurea and polyurethane membranescan also be produced by the appropriate choice of monomers (see, e.g., ref. g), although most work in the biomedicaYpharmaceutica1 area has been carried out with the polyfunctional halide plus polyamine system to give polyamide membranes, and thesesystems will be considered in more detail. The monograph by Morgan [lo] extensively reviewed the fundamentals of interfacial polymerization, and this review was extended by Millich and Carraher [ll].The complexity of the reaction has been shown by

Ii2N-(CH&”N

H2

1,6-hexane diamine

+ cl oc -(CH2)R-COCI

sebacoyl

chloride

polyamide HN-(CH,),-NH-~O-O-(CH,)B-OCO

Nylon6,lO

H ~ N - - ( C H ~ ) ~ - - N H ~ 1,6-hexane diamine

+ C1 W-@

-

CI

terephthaloylchloride

HN-(CH,),-NH-C

(b)

Fig. 4 Reaction for formation of (a) nylon 6,lO; (b) poly(terephtha1amide); (c) poly(terephthaloy1 L-lysine);(d) polyester.

354

Whateley

COOH

,I

L-lysine

H2N-(CH2)3XIi-NIi2 ,

(c)

.

+

' .

poly(terephthaloy1 L-lys'ine)

2,2-bis(4-hydroxyphenyl)

+ Cl OC-(CHz)a-COCI'

;&-sf (a

Fig. 4 Continued

e

.'

sebacoyl

chloride

-

c H

)

2

8

4

propane

Microcapsules Polymerization by

Complexation and

355

studies at simple unstirred planar interfaces, and some aspects of this will be considered here.

1. Factors Regarding the Interfacial Polymerization Reaction'

. .. .

a. Reaction rate. The interfacial formation of nylon 6 , l O is rapid, and a ,visible polymeric precipitate can be.seen almost immediately afterthe two components are brought together. The reaction rates.for polyamide,,formation have second-orderrate constants in the range10'-lo6 dm3mol" s-l [lo]. The reaction of aliphatic glycols with diacid chlorides is too slow to allow the fofmation of high molecular weight polymer, with hydrolysis of the acid chloride being the competingside reaction. In the interfacial polymerization reaction between piperazine and terephthaloyl. chloride; the oligomers react at a ,faster rate with the diacid chloride than piperazine-itself. Thus, atrelatively low concentrations, high molecular weight polymer (1O;OOO-40,000) can be formed.

b. Precipitation ofpolymer. The site of the reaction affects the precipitaof the polymer: me reaction forming nylon 6 ; l O has usually been assumed to occur onthe organic side of the interface. Precipitation of the polymer prevents diffusion of the monomers and thus stops thepolymerization reaction from.continuing to completion and limits the thickness of the membrane. In this respect, the choice of solvent is important: swollen polymers will allow diffusion of monomers to the reaction site.

. tion

c. Impurities. Typical &purities in interfacial reaAions of this type are monofunctional monomers which produce low molecular weight polymer as' a result of chain termination. The presence of ethanol as a stabilizer in chloroform appreciably reduces the molecular weight of poly(piperazine sebacamide) [lo, p. 901.

d. ' Diamine partition coeficient and interfacial transfer rates. The partitio*'coefficient.of the diamine.is easily measured atequilibrium and serires as a guide to exclude monomers which would.be unsuitab1e;forthis type of reaction. However, .transferrates-become .more important when polymer.ization' is in progress. ,Maintenance of adequate. levels of diamine at. the reaction s i t e k important, and thercfore.therate of transfer should exceed . . . 'the rate'ofremoval of diamine by polymerization. . e. Concentrationratioof reactants. The concentration ratio of the two reactants can be important when highmolecular weight polymer is desired. However, in microencapsulation applications this ratio has not been studied to any great extent; mainly because other. conditions of the reaction place restrictions on the concentrations which can be used. Thus,, in mi-

356

Whateley

croencapsulation, generally one reactant is used in excess. When oil-in water ( O N ) emulsions are used in the microencapsulation of pesticides, the water-soluble monomer can be used in excess, and therefore the microcapsule wall thickness is controlled by the complete diffusion of the oilsoluble monomer to form thewall. Diamine canbe used in excesswhen its partition coefficient is unfavorable for its transfer to the Organic solution, as is found whenlysine isused.

5

Stirring. Stirring is a critical variable and is found to affect the molecular weight of various types of polymers [lo]. When the interfacial reaction is carried out toproduce membranes aroundemulsion droplets, thestirring conditions can make thedifference between a usable product and auseless mass of polymeric material. The stirring produced by a magnetic follower in a beaker, as would be commonly available in a laboratory, can produce microcapsules down to about10-20 pm in diameter. More powerfulemulsification is generally required to produce smaller microcapsules. Kondo and coworkers (e.g., Wakamatsu et al. [12]) haveprepared small microcapsules (diameter about 1 pm) using a high-speed impeller in a special reaction vessel.

g . Hydrolysis of diacid chloride. Inactivation of diacid chlorides is perhaps one of the most important side reactions, but in many reactions, it causes no problem. The partition of the diacid chloride into the aqueous phase is the principal factor involved, and by restricting the choice of monomer, the problem can be eliminated. The kinetics of hydrolysis of terephthaloyl chloride have been studied in a two-phase system (n-heptane/ water) and found to befirst order with respect to the diacid chloride [13]. However, Bradbury et al., [l31 reported that hydrolysis was much slower than polyamidationwith piperazine as diamine. h. Transfer rates of salts. An important consideration is the elimination of HCl formed in the polycondensationreaction. This product is removed by the formation of the hydrochloride salt of the diamine which, being poorly soluble in the organic phase, diffuses to the aqueous phase where the HCl is neutralized by buffer included for this purpose. Hydrochloride salts of diamines are not able to react with diacidchloride and, therefore,if buffer is not included in the aqueousphase, the diamineitself will function as an acid acceptor. In most cases, the transfer of hydrochloride salts will be faster than the transfer, in the opposite direction, of diamine to the reaction site. i. Solvent:polarity,density,andtoxicity. Chloroform-cyclohexanemixtures in the ratios 1 : 4 or 1 : 3 have been widely used. Morgan [lo] dis-

Microcapsules Polymerization by Complexation and

357

cussed the polymer-solvent interactions of nylon 6,lO and has shown that thick films are expected from chloroform systems, whereas cyclohexane produces thin films. Good solvents tend to producehigh molecular weight polymer when comparedwith nonsolvents. The density of the organic solvent used in the production of the W/O emulsion in microencapsulation procedures will affect the stability of the emulsion and will determine whether the microcapsules sediment or float in the organic phase. This will affect the isolation technique used to obtain the newly formed microcapsules. In general, when microencapsulation for pharmaceutical applications has been studied, organic solvents have been chosenfrom solvents withlowtoxicity and toxic solvents havebeen avoided. j . Surfactants. The addition of surfactants in microencapsulation procedures is important for the formation of the emulsion. Surfactant also plays an important role in the transfer of the diamine to the organic phase. Kondo and coworkers have investigated the effect of Span 85 (sorbitan trioleate) on the partition coefficients of various diamines (e.g., Koishi et al. [14], Koishi et al. [15], and Shigeri et al. [16]) and bisphenols [12]. In general, increasing the concentration of surfactant results in an increase in the amount of diamine whichis transferred to the organic phase. For preparation ofpoly(1ysine terephthalamide) microcapsules, a surfactant concentration (sorbitan trioleate) of 20% v/v was successful. The choice of surfactant used in studies where aqueous solutions are encapsulated in semipermeable membranes has been limited mainly to sorbitan trioleate, which has a particularly low HLB (hydrophile-lyophilebalance) value (i.e., 1.8). The main requirement of any surfactant is that it does not react with the diacid chlorides, and it does not have impurities which would interfere with the polymerization reaction.

k. Temperature. The great advantage of interfacial condensation polymerization reactions of this type is that they can be carried out at room temperature. No advantage is found in heating the reaction and many preparations are improved by controlling the temperature rise. The reactions proceed well at 4°C. When enzymesand/or proteins are to bemicroencapsulated, the process can becarried out conveniently at 4"C, a temperature atwhich enzymes in solution have favorable stability. Ultrathinmembranes,formedatthe interface betweenaqueous polyethylenimine andhexane solutions of tolylene diisocyanate, were found to be extremelystrong [17]. Such filmsare able to beused in reverse osmosis where pressures of some 100 atm are encountered.

2. Preparation of PolyamideMembrane;Semipermeable, Aqueous-Core Microcdpsules by Interfacial Polymerization . A typical'method to produce nylon 6,lO microcapsules is as follows: 1; Nonaqueous phase: chloroform/cyclohaxane ( l : 4) containing 5% Span 85 (sorbitan trioleate). 2. Aqueous phase: sodium carbonatehicarbonate buffer, pH 9.8 containing,. for example, qnzyme to be microencapsulated plus protective/stabilizing protein (e.g., hemoglobin or bovine serum albumin) and 1,6-hexamethyiene:diamine. . 3. Above phases mixed in about a 1 : 10 volume ratio and stirred while cooling in an ice bath. Stimng rate and method to ,give required microcapsule size. 4. Further nonaqueousphase containing sebacoyLchloride added to . . stirred emulsion. Interfacial polymerization ,reaction allowed to 30 min. . . about for . . proceed 5. Polymerization reaction quenched by addition of excess nonaqueous phase, Microcapsules allowed to sediment and superpatant ,. decanted off. 6. Microcapsules resuspended in 50% %een 20 in saline,sedimented,, decanted, and this,process repeated to remove organic . . solvents. . 7. Microcapsules washed 2 times with saline. and resuspended in saline. . '

. .

'

,

Some particularaspects of the process: chloroform and cyclohexane do not readily denature proteins. The particolar ratio.'(l : 4).is chosen for 'two . ' reasons: . ' 1

I

.

a

1. The density (about 0.91) is'not far-removed from water so,that emulsion formation is not a problem, butstill allows the aqueous core microcapsules to sediment for separation purposes. 2. The partition coefficient of.the diamine between the aqueous and ., organiG,phases is important:.the 1 :.,4..rati,oallows diamine to diffuse into the organic phase, where the- interfacial polymeriiation reaction takesplace. AS the interfaciaimembrane, forms, the , . diffusionof the. diamine to the organic interface of the membrane is reduced,and the polymerization becomes self-limiting. I

,

,

Thus, about 200-nm thickness microcapsule membranes, which are strong enough for separation;washing, and applications, areformed with equiva. .. . . . lent pore sizes of about 0.2nm. : . ' Interfacial pdymerization .in a high-voltage electric field has been investigated as a means of obtaifiing more monodisperse collections.of "

4

'

Microcapsules Polymerization by

and Complexation

359

microcapsules [18]. Kondo and his group have studied this aspect 'of microcapsule formation (e.g., Muramatsu et al. [19]). The proteins human serum albumin (HSA), bovine fibrinogen, and ovalbumin were used [20]. to prepare' microcapsules by interfacial crosslinking using tere-phthaloyl chloride -and subsequent treatment with hydroxylamine to make the microcapsules capable of iron binding. 'Similar polyhydroxamic serum albumin microcapsules had been reported previously by Levy et al. [21].

-

3. Applications of Semipermeable, and Aqueous "Chang"-Type Microcapsules ,. '

This topic has been well covered'h'the book by Lim [4] and reviewed . . by Chang [3]. '

a. Artificialred blood cells. Thesehave been,used as asubstitute in blood transfusions [22]. Hemoglobin, crosslinked hemoglobin and,hemoglobin conjugated to polymers have been microencapsulated for this purtype of.microcapsule in obtaining pose. There is a major problem withathis long circulating times, The size mustbe in the rangeof,red blood cells (e.g., 4 0 pm). A more difficult problem is in achievingthe.elasticity.and fluidity of the membrane needed forpassage along small capillaries .(<
.

b. Microcapsules containing adsorbentsandimmunoadsorhents. Microencapsulated charcoal is biocompatible, that is, htis no adverse.effect on blood butstilhetains its ability to,absorbsignificant amounts of low molecular ,weighk material (see, e.g.,' 'refs. 23 and 24). Applications Of hemoperfusion with a device containing &out 70 g microencapsulated _.. . . charcoal include: . . . .

.

.

I.

'

...

.

. . .

Treatment ,of .acute,poisoning .( -Treatment of kidney,failure . . . .

. .

,

. . . .

8

. . . ......

PS

c;, Microcapsules coniaining h n igcells'.' :' topic is 'covered in Sedtion IV. The useof'interfacial polymerization in'this areahas been reviewedby Chang [3].

d. Microcapsules containirzg enzymes.(br.multienzymesystems): ,-.,

..

....

._ .-, .. .

.

...

0:'

.. .....

.... Urea removal in. kidney failure; using urease 'which converts' the urea .to.ammonia and carbon dioxide. Co-encapsulation of an-ammonia absorbentcaqremovethe.ammonia [3]. . . . __ ..

Hereditary enzymedeficiency; for example: Phenylketonuria: the 'enzyme phenylalanine ammonia lyase is-defi'.cient and is replaced by the microencapsulated.enzyme [3]..

Whateley

360

Histidenemia: the enzyme histidase has to be replaced-to avoid the build-up of histidine. Using a histidinemicmouse model, Wood et al. [25] worked on this problem. Histidase from a bacterial source was microencapsulated in poly(terephthaloy1piperazine) microcapsules and administered intraperitoneally. Although blood histidase levels were reduced 50%, there were problems with the toxicity of the microcapsule preparations. The multienzyme “artificial cell”-type system is covered in Lim [4] and Chang [3]. Enzyme stability: Arginase microencapsulation with a poly(phthaloy1piperazine) membrane was reported by Kondo [26]. He concluded that enzyme inactivation during the process of microencapsulation was due to contact of the arginase with the organic solvent and incorporation of the enzyme into themembrane. Wood and Whateley [27] investigated the loss of enzyme activity during the interfacial polymerization microencapsulation process. The low yields (about 40%) of activity found in polyamide microcapsules for bothchymotrypsin and histidase are typical of those reported in the literature. There was some diffusional restriction on the measured enzyme activity, but there was a considerable incorporation of model radiolabeled proteins into the microcapsule membrane during the microencapsulation procedure. Tables 1and 2 show the distribution of labeled albumin and fibrinogen, respectively, following the microencapsulation process. Table 3 shows that fibrinogen is adsorbed to a much greater extent than albumin on poly(piperazine-terephthalamide) microcapsules. Kidokoro et al. [28] looked at theinteraction of fibrinogen with microcapsules of poly(L-lysine&terephthalic acid). They found that fibrinogen increased the disintegration of the microcapsules by adsorbing via a hydrophobic interaction. Hoshino et al. [29] studied the thermostability of glucose oxidase within polyurea microcapsules. Thermostability markedly increased for the microencapsulated enzyme, but this was ascribed to the incorporation of Table 1. Incorporation of ‘ZSI-L,abelled Human Serum Albumin in Poly(piperazine terephthalamide) Microcapsules ‘%activity

System

phase Organic washingsAqueous Microcapsules (4.3)

(%) (3.8) 64.1

6.4,6.8,8.0 28.8 31.6,24.4,30.3 60.9,69.0,62.5

total

Mean with s.d. 7.1 (0.8) 100.0

361

Microcapsules by Polymerization and Complexation

Table 2. Incorporation of '=I-Labelled Fibrinogen in Poly(piperazine terephthalamide) Microcapsules System Organic phase 10.9 Aqueous washings Microcapsules

'%activity (%) 12.3,10.5

9.9, 15.2,9.5,13.3 65.4,75.8,57.6

total

Mean with s.d.

(1.2) 12.7 (2.9) 66.3 (9.1) 89.9

Table 3. Adsorption of '251-Labelled Proteinsonto Poly(piperazine terephthalamide) Microcapsules Contact time

Albumin %

5 min

3h 95 24 h

1.2 1.7 3.2

Fibrinogen % 92

(in 0.9% w/v NaCl, pH 7.4)

the enzyme into the microcapsule wallduring the interfacial polymerization process, as was also reported by Wood and Whateley[27]. Enzyme kinetics: The effect of microencapsulation on enzymekinetics has been investigated by many workers. Urease was microencapsulated at 83% yield with a semipermeable membrane[30]. Only 6% of the microencapsulated enzyme was incorporated into the nylon membrane, and the high activity of the encapsulated enzyme (92%) indicated minimal effects of mass-transfer limitation. Aisina[31]used Chang-type microcapsules with the inert filler of poly(ethy1enimine) to study trypsin and chymotrypsin kinetics. Not only were low molecular weight substrates used, but the effect on the autolysis of trypsin and the autoactivation of trypsinogen were also studied. The Michaelis constant, K,,,, was 0.49mM for thenative chymotrysin but 3.7 mM for the microencapsulated enzyme with acetyltyrosine ethyl ester (ATEE) as substrate. The catalytic constant, k,,, was reduced from 175 to 12 S" for the microencapsulated chymotrypsin. The maximum of the apparent pH-activity curve for chymotrypsin was found to beshifted 1pH unit to more alkaline values when the enzyme was microencapsulated in a polyamide membrane [27]. Fig. 5 shows this I effect. This phenomenon was explained in terms of the hydrogen ion concentration in the microenvironmentof the enzyme within the microcapsule being higher than in the bulk solution.

n

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Whateley

5

7

6

8

9

10

PH Fig. 5 pH-activitycurvesformicroencapsulateda-chymotrypsin chymotrypsin in solution (A)(1p M A a E , 50 p M NaCl at 25°C).

(0) and a-

. . e; Microcapsules containing antibodies; An interesting application was reported by Wallace ‘[32], who developed an assay method for thyroxine . and some steroids by microencapsulating,antibodies in microcapsules (25 ..pmdiameter with membrane poresize permitting entry of hormones of less than 4000 molecular weight) Routine assays were simplifiedby coen-capsulating magnetic iron dextran so that centrifugation could be super. seded by magnetic separation. Other reportsrelating to microcapsules containing antibodies are inciuded in the book by Lim [4] and the review by Chang [3]. . ’

5

’

D$ugdelivery.’ Although interfacial polymerization has notbeen widely used €or the microencapsulation of drugs, Luzzi ‘etai. [33], Florence and Jeikins [34]’,and McGinity and his group have expanded the method into this‘area [35,36].’Interestingly, some water-soluble polyamides for use as drug ca’mers have been synthesized by interfacial polymerization [37]. . . Amine derivatives of poly(ethy1ene glycol)were reacted with succinyl chloride at theinterface of a methylene chloride-water system. . . > ’. g . Otherapplications of microencapsulationby interfacial polymerization. A novel andinteresting application of semipermeable microcapsules was reviewed by 0’Neill[38l1 Magnetic ferric oxide.was.microencapsu1ated in a membrane consisting of a graft copolymer .of polyamide and poly(ethylenimine) (PEI). These were administered to animals together with

Microcapsules Polymerization by Complexation and

363

some known carcinogens, for example, nitrosoureas. The microcapsules were recoveredmagnetically and thespecies bound to thePE1 investigated in order tofollow metabolic changes andtypes of carcinogens occurring in the gut under various dietary situations. The microencapsulation of benzalkonium chloride based on the interfacial polycondensation of isocyanates has been reportedby Pense et al. [39-411. The use of thz HLB approach was shown to be appropriate for deciding the initial formulation [39,40]. An aqueous solution of benzalkonium chloride was poured into an oil phase (e.g., xylene or a blend of xylene and Miglyol) containing a polymeric surfactant (e.g., Hypermer A60). Polycondensationfollowed hydrolysisof the isocyanate and reaction between the formed amine and furtherisocyanate. The preparation and properties of polyurea microcapsules was reported by Yan et al. [42,43] usingcyanate and polyamine asthe monomers witha nonionic surfactant as the emulsifier. The same group [44] also studied polyamide microcapsules containing oily liquids. The preparation of oil-containing poly(terephtha1amide) microcapsules by interfacial polymerization has also been recently discussed by Alexandridou and Kiparissides [45]. 4. Studies of Microcapsules Prepared by Interfacial Polymerization Lee and Kondo [46] found at 20°C for KC1 a permeability of 10.2 X 10-7 cm S-' by following the diffusion though the membrane by an optical method. The same group [47] studied the effect of pH on thepermeability of poly(L-lysine-alt-terephthalic acid)microcapsules to electrolyte ions. A large increase in permeability in going from pH 4 to 6 (and above) was ascribed to an abruptincrease in the microcapsule size. Levy et al. [48,49] used Fourier transform infrared spectroscopy (FT" IR) to study the HSAmicrocapsules. By following the influence of reaction time, they were able to show after 5 min the progressive acylation of the hydroxy and carboxylate groups of HSA. Free amino groups were also determined by a back-titration method [48,50,51]. The interfacial polymerization of m-phenylenediamine andtrimesoyl chloride has been studied by Chai and Krantz [52] using the novel techniques of light reflection and pendant-drop tensiometry allowing rapid real-time assessment of the polyamide membrane formation.

B. Nonbiomedical Aspects of Microcapsules with Rigid, (Type 2 in Fig. 1) Nonpermeable Shells and Liquid Cores 1. The major application of microcapsules with rigid, nonpermeable and brittle membranes has been in the carbonless copy paper industry.

Whateley

364

2.

3.

4. 5.

111.

The underside of upper copies is coated with such inkcontaining microcapsules. These are ruptured by the pressure of writing, and the released ink interacts with the lower paper to leave the imprint. However, in general, such microcapsules are not prepared by interfacial polymerization. More specialized applications of this type of microcapsule allow preparation by the interfacial polymerization process, €or example, herbicide and pesticide microencapsulation for controlled release. As well as controlling the release, the active ingredient is also kept out of contact with the environment andhandlers. This topic is well covered in Wilkins [g]. Fragrances, for example, are microencapsulated for the “scratch ‘n sniff”-type application fixed to such articles as paper and “tee” shirts. Domestic gas samples are microencapsulatedto provide a simple and readily distributed means of indicating the smell of a gas leak. Pigments have been microencapsulated (e.g., see ref. 53). Duel1 and Pendergrass [54] have reported on the use of polyfunctional aziridines for interfacial polymerization for a variety of applications.

INTERFACIALADDITIONPOLYMERIZATION

Interfacial addition polymerization has features similar to interfacial polymerization, but the polymerisusuallycomposed of only one type of monomerand there is nootherproductformed in the reaction. This method, which does not generally require reactants in the aqueous droplets, should be particularly suitable for enzyme microencapsulation. However, in general, solid microspherestend to beproduced ratherthan aqueous-core-type microspheres. Alkylcyanoacrylates A.

There has been considerable interest in the use of the alkyl-cyanoacrylates to form microcapsules and nanoparticles for drug delivery and targeting by the groups of Couvreur andKreuter. Florence et al. [55]have also reported the preparation of biodegradable microcapsules from poly(alky1-2-cyanoacrylate) monomers. The anionic addition polymerization of analkyl cyanoacrylate is shown in Fig. 6. This process could be initiated, for example, by water in the aqueous phase of an water-in-oil emulsion with the cyanoacrylate dissolved in the oil phase.

Microcapsules by Polymerization and Complexation

365

CN I CH, = C -COOK

CN

I

1

CN CN I I HO-CH,- C - CH, - C - ClOOR I COOR

Fig. 6 Reaction for formation of poly(cyanoacry1ate).

Both the groups of Kreuter and Couvreur (e.g., seerefs. 56 and 57) have developedthese systems overanumber of years. Forexample, Kreuter et al. [58] studied the degradation of poly(butylcyanoacry1ate) and poly(hexylcyanoacry1ate) nanoparticles and their association with and toxicity toward isolated hepatocytes. Formaldehyde has been regarded as being the major problem in considering the toxicity of poly(cyanoacry1ate)(PCA) systems.However, the (low)toxicity toward hepatocytes (which did not take up nanoparticles to any significant extent) could not be attributed solely to formaldehyde during degradation. Leonard et al. [59] studied the degradation of the poly(alkylcyanoacry1ates) in detail. Troster and Kreuter [60] studied extensively the body distribution of radiolabeled poly(methyLmethacry1ate) nanoparticles with and without surface surfactants. Poloxamine 1508 was found to be the most effective in reducing liver uptake. The anticancer drug mitoxantrone was loaded into PCA nanoparticles and found to cause a significant volume reduction in the B16 tumor in rats [61]. Harmia et al.[62] and Harmia-Pulkkinen et al. [63] encapsulated pilocarpine in poly(butylcyanoacry1ate) nanoparticles for drug delivery to the eye. The effects of monomer and stabilizer concentrations on the properties of poly(buty1-2-cyanoacrylate) nanoparticles was investigated by -Nonso et al. [M].

Whateley

366

Poly(cyanoacry1ate) nanoparticles have been evaluated as drug carriers by other groups also; for example, Douglas et al. [65] and Puglisi et al. (661Detailed studies of the particle size distribution of poly(cyanoacry1ate) nanoparticles have been reported [67,68]. IV. INTERFACIAL COMPLEXATION OR INTERFACIAL POLYELECTROLYTE COMPLEXATION

A. AlginatelPolylysineMicrocapsules This process was introduced by Lim and Sun [69] and has been developed by their groups and Sefton et al. (e.g., see ref. 71). The topic has been reviewed recently by Sun et al. [70] and Chang [3]. The fact that calcium ions crosslink the alginate species (Fig. 7), initially in solution as the sodium salt, allows droplets of gel to be formed in the calcium chloride solution. The addition of an oppositely charged polyelectrolyte (in this case

poly(mannuronic) acid

poly(gu1uronic)

acid 0

poly(mannuronic) Coolt

- poly(gu1uronic) COOH

Fig. 7 Structures of alginate components.

acid

Microcapsules Polymerization by

and Complexation

367

the polylysine polycation) allows a layer of insoluble polyelectrolyte complex to form at theinterface. The calcium ions remaining in the interiorof the microcapsule are exchanged with an excess of sodium ions, rendering theinterior fluid again. Thus,an aqueous-cored, semipermeable microcapsule is formed under mild conditions (i.e., no extremes of pH, no chemical reactions). The polyelectrolyte complex membrane has proven to be very biocompatible, an aspect covered in more detaillater. l . Preparation of Microencapsulated Cells Cells are suspended in 0.8% sodium alginate (structure shownin Fig. 7) in saline: Droplets are extruded through a syringe into 1.5% calcium chloride solution. The crosslinking of the alginate macromolecules by the calcium ions causes the droplets to become gels (withthe cells entrapped in the gel sphere). The separatedgel microspheres are then resuspendedin a polylysine solution (0.02%, molecular weight, 35,000) for 5 min. An interfacial macromolecular complexation occurs at the surface of the gel microsphere, forming an insoluble (but semipermeable membrane). The gel microspheres are separated and resuspended in saline at pH 7.4; the calcium ions diffuse out of the gel interior of the microcapsule causing the contents to liquify. These microcapsules tend to be large (>l00 pm)perhaps macrocapsule is a better term? Mannuronic-rich alginate has been shown to link more strongly to the polylysineby Dupuy et al.[72], who used FT-IR to study polylysine/ alginate membranes containing various contents of mannuronic or guluronic acid residues (structures in Fig. 7). A novel method of preparing small alginate/polylysine microcapsules (in the range 5-15 pm) was reported by Kwoket al. [73]. Vaccine-containing microcapsules were preparedusing aTbrbotak air atomizer to spray sodium alginate solution into calcium chloride solution to form temporary microgel capsules which are subsequently crosslinked with polylysine. A different polyelectrolyte complexation reaction was used by Stange et al. [74] to encapsulate liver microsomes. The polyanion cellulose sulfate (sulfate substitution ratio 0.4-0.5)wasusedwith the polycation poly(dimethyldiallylammonium chloride) (molecular weight 40,000)The resulting membrane was semipermeable with an exclusion limit varying from 25,000 to 150,000 molecular weight depending on the preparation conditions. Increased enzymic activity and increased wall strength were claimed for this procedure. The microencapsulation and in vitro performance of various mammalian cells in polyacrylate membranes has been reviewed by the Sefton group at Toronto [71]. The cell suspension is coextruded with the polymer

368

Whateley

solution through a concentric needle assembly and the concept of this novel way for the controlled delivery of therapeutic agents is discussed. Drug encapsulation in alginate microspheres has been reported by by an emulsiWan et al. [75]. Calcium alginate microspheres were produced fication process and hardenedwith isopropyl alcohol or acetone. However, drug loading was found to below.

2. Applications of AlginatelPolylysine Microcapsules Because of the mild conditions in preparation and the biocompatible nature of the formed microcapsules, the vast majority of applications of microcapsules prepared by the interfacial complexation method are in the pharmaceutical, biomedical, and medical fields, and only these applications will be considered. These applications have been reviewed by Sun et al. [70] and Chang [3]. The transplantation of mammalian cells encapsulated in a protective, biocompatible, and semipermeable membrane has greatclinical potential for a range of diseases requiring enzyme- or cell-replacement therapy. The interfacial complexation process has been used to encapsulate islets of Langerhans, the cells which secrete insulin, for the treatment of diabetes. Limited success has been achieved [69,76,77]. Hepatocytes have also been microencapsulated by this process for the treatment of liver failure [78]. Another majorapplication of microencapsulated cells has beenin the production of monoclonal antibodies. The hybridoma cells, producting the monoclonal antibodies, are entrappedin the microcapsules: nutrients, oxygen, and so forth can enter through the semipermeable membrane. The monoclonal antibodies secreted by the encapsulated hybridoma cells remain trapped inside the microcapsules and accumulateto a greater concentration thanin the absenceof encapsulation. To recover the antibodies, the microcapsules are separated from the culture medium and ruptured to release the antibodies. In theabsence of encapsulation, the antibodies must be isolated from a large volume of culture medium.

3. Permeability of AlginatelPolylysine Microcapsules CoromiliandChang [79] haveused a heterogeneousmixture of dextrans with molecular weights 10,000-500,000 to study the membrane permeability of alginatelpolylysine microcapsules. Hybrid artificial organs must permit the diffusion of smaller molecules, includingpeptides and proteins, but must exclude leukocytes and immunoglobulins( M W >150,000) Fluorescein isothiocyanate-labeled dextrans were used by Vandenbossche et al. [80] to determine the molecular weight cut-off of similar microcapsules. The molecular weight cutoff of alginatelpolylysinemicrocapsules has

Microcapsules Polymerization by

and Complexation

369

been shown to be dependent on preparative conditions [81]. Microcapsules prepared at 40°C showed a lower MW cutoff when the concentration of polylysine wasincreased from O.l%(w/v) The ultrastructural characterization of alginatellysine microcapsules was reported by Shimi et al. [82] Poly(1ysine) ofmolecular weight 22,000 was found to be optimum in forming robust microcapsules whichwere relatively impermeable to high molecular weight species such as immunoglobulins. 4. Biocompatibility of AlginatelPolylysine Microcapsules The topic of biocompatibility is a huge area in its own right: A few recent references are discussed here concerning the biocompatibility of alginate/polylysine microcapsules. The biocompatibility of alginate/polylysine microcapsules was enhanced by Sawhney et al. [83] by photopolymerizing a PEG-based hydrogel onto thesurface of alginate/polylysinemicrocapsules. A less inflammatory response with no fibrotic response was reported. The effect of composition on biocompatibility was examined by Clayton et al. [84], who found that in the peritoneum, high mannuronic acid alginate capsules provoked the weakest response. Islets of Langerhans inmicrocapsules of alginate/polylysine have been proposed as a artificial pancreas (e.g., see ref. 85), who studied the capacity of such microcapsules to activate macrophages in order tounderstand the foreign body reaction withfibrosiswhich has been observed around implanted microcapsules. The host reaction against alginate/polylysine microcapsules has been studied with both emptymicrocapsules [86] and with microcapsules containing living cells [87]. Histological evaluation showed a significantly higher number of cells stickingto microcapsules containing cells as comparedwith empty microcapsules. REFERENCES 1. R. Arshady, Biodegradable microcapsular drug delivery systems: Manufacturing methodology, release control and targeting prospects, J. Bioactive Compatible Polymers, 5:315-342 (1990). 2. R. Arshady,Preparation of microspheresandmicrocapsulesbyinterfacial polycondensation techniques, J. Microencaps., 6:13-28 (1989). 3. T. M. S. Chang, Recent advances in artificial cells based on microencapsulation, in Microcapsules and Nanoparticles in Medicine andPharmacy (M. Donbrow, ed.), CRC Press, Boca Raton, FL, 1992, pp. 323-339. 4. F. Lim, (ed.), Biomedical Applications of Microencapsulation, CRC Press, Boca Raton, FL, 1984.

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5. M. Donbrow, (ed.), Microcapsules and Nanoparticles in Medicine And Phar-

macy, CRC Press, Boca Raton, FL, 1992. 6. T. L. Whateley (ed.), Microencapsulation of Drugs, Harwood Academic Publishers, Reading, UK, 1992. 7. T. M. S. Chang, Semipermeable microcapsules, Science, 146524-525 (1964). 8. T. M. S. Chang, F. C. Macintosh, andS. G . Mason, Semipermeable aqueous microcapsules, Can.J. Physiol. Pharmacol., 44:115-128 (1966). 9. R. M. Wilkins, (ed.), Controlled Delivery of Crop-Protection Agents, Taylor and Francis, London, 1990. 10. P.W. Morgan,CondensationPolymers:InterfacialandSolutionMethods, NY, PolymerReviewsSeries N0.10, IntersciencePublishers,JohnWiley, (1965). 11. F. Millich and C. E. Carraher, Jr., (eds.), Interfacial Synthesis, Volume 1: Fundamentals, Marcel Dekker,N Y , USA (1977). 12. Y. Wakamatsu, M. Koishi and T. Kondo, Microcapsules 17: Effect of chemical structureof acid chlorides and bisphenolson the formation of polyphenyl ester microcapsules, Chem. Pharm. Bull. 22,1319-1325 (1974). A. N.Hambly,Kineticsof interfacial 13. J. H.Bradbury, P.J.Crawfordand polycondensation reaction, Trans. Far. Soc. 64, 1337-1347 (1968). 14. M.Koishi, N. FukuharaandT.Kondo,Microcapsules2:Preparationof polyphthalamide microcapsules, Chem. Pharm. Bull. 17,804-809 (1969). 15. M. Koishi, N. Fukuhara and T. Kondo, Studies on microcapsules 4: Preparationandsomeproperties ofsulphonatedpolyphthalamidemicrocapsules, Can. J. Chem. 47,3447-3451 (1969). 16. Y. Shigeri, M. Koishi, T. Kondo, M. Shiba andS. Tomioka, Microcapsules 6: Effect of variationsin polymerization conditionson microcapsule size, KolloidZ Z.Polym., 249,1051-1055. Strength of interfacial polymerization films, 17. K. J. Mysels and W. Wrasidlo, Langmuir, 7:3052-3053 (1991). Etuk, andK.R.Murray,Formationofnylon6,lO 18. L.R.Weatherley,B. capsules by interfacial polymerization in a high-voltage electric-field, Powder Technol., 65227-233 (1992). T. Kondo,Preparation ofpolyamidemi19. N. Muramatsu,K.Shiga,and crocapsuleshavingnarrowsizedistributions,J.Microencaps.11:171-178 (1994). Levy, Polyhydroxamicmicrocapsules 20. D. Hettler, M.C.Andry,andM.C. Mipreparedfromproteins--Anoveltypeofchelatingmicrocapsules,J. croencaps., 11:213-224 (1994). 21. M.C. Levy, D. Hettler, M.C.Andry,and P. Rambourg, Polyhydroxamic serum albumin microcapsules: Preparation and chelating properties, Int. J. Pharm., 69:Rl-R4 (1991). New 22. T. M. S. Chang and R. Geyer (eds.), Blood Substitutes, Marcel Dekker, York, 1989. 23. T. M. S. Chang, Blood compatible coatingsof synthetic immunoadsorbents, Trans. Am. Soc. Intern. Artif. Organs, 26:546 (1980).

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S. Chang, P. Barre,and S. Kuruvilla, Long termreducedtime haemoperfusion-haemodialysiscompared to standarddialysis,Trans.Am.

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Soc. Intern. Artif. Organs, 31572 (1985). 25. D. A. Wood, T. L.Whateley,andA. T. Florence, Microencapsulation of histidaseforenzymereplacementtherapy,J.Pharm.Pharmacol.,31:79P (1979). 26. T. Kondo, Enzyme inactivation in microencapsulation, in Microencapsulation (J. R. Nixon, ed.), Marcel Dekker, New York, 1976, pp. 67-75. T. L. Whateley,Astudyofenzymeand protein micro27. D.A.Woodand encapsulation-somefactorsaffecting the low apparentenzymicactivity yields, J. Pharm. Pharmacol.,3432-557 (1982). 28. M. Kidokoro, H. Ohshima, and T. Kondo, Interaction of poly(L-lysine-aftterephthalic acid) microcapsules with fibrinogen, J. Microencaps. , 8:63-70 (1991). on the thermostabilityof 29. K. Hoshino, N. Muramatsu, and T. Kondo, A study microencapsulated glucose oxidase, J. Microencaps. ,6:205-211 (1989). 30. M. Monshipouri and R.J. Neufeld, Activity and distribution of urease followingmicroencapsulationwithinpolyamidemembranes,EnzymeMicrobial Techn., 31:309-313 (1991). 31. R. B. Aisina, Effectof microencapsulated enzymes, in Microencapsulation of Drugs (T. L. Whateley, ed.), Harwood Academic Press, Reading, UK, 1992, pp. 215-231. 32. A. M. Wallace, Novel immunoassays for clinically important hormones based on microencapsulatedantibodies,inMicroencapsulationofDrugs(T.L. Whateley, ed.), Harwood Academic Press, Reading, UK, 1992, pp. 241-255. 33. L. A. Luzzi, M. A. Zoglio, and H. V. Moulding, Preparation and evaluation J. Pharm. Sci., 59:338of prolonged release properties of nylon microcapsules 341 (1970). 34. A. T. Florenceand A. W. Jenkins,Invitroassessmentofmicroencapsulated drug systems as sustained release parenteral dosage forms, in Microencapsulation,(J. R. Nixon,ed.),MarcelDekker,NewYork,1976,pp. 39-55. 35. G. W. Cuff and J. W. McGinity, Expanded versatility of microcapsules prepared by interfacial polymerisation,J. Microencaps. 1:343-347 (1984). 36. G. W. Cuff, A. B. Combs, and J. W. McGinity, Effect of formulation factors on the matrix pH of nylon microcapsules, J. Microencaps., 1:27-32 (1984). 37. U. Chiba, E. W. Neuse, J. C . Swarts, and G. J. Lamprecht, Water-soluble polyamides as potential-drug carriers. 7. Synthesis of polymers containing intrachain-type or extrachain-type amine ligands by interfacial polymerization, Angewandte Makromolekulare Chemie, 214:137-152 (1994). 38. I. O’Neill. Reactive microcapsules for detectionof carcinogen sources in the gut, J. Microencaps., 10:283-308 (1993). 39. A. M. Pense, C . Vauthier-Holtzscherer, and J.-P. Benoit, Microencapsulation of benzalkonoium chloride, in Microencapsulation of Drugs (T. L. Whateley, ed.), Harwood Academic Press, Reading, UK, 1-5.

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40. A. M. Pense, C. Vauthier, F. Puisieux, and J.-P. Benoit, Microencapsulation of benzalkonium chloride, Int. J. Pharm.,81:lll-117 (1992). 41. A. M. Pense, C. Vauthier, and J.-P. Benoit, Study of the interfacial polycondensationofisocyanateinthepreparation of benzalkoniumchloride loaded microcapsules, Colloid Polymer Sci., 272:211-219 (1994). Li, Size distribution and zeta-potential of micro42. N. Yan, M. Zhang and P. capsules. J. Membrane Science, 72, 163-169 (1992). of polyurea mi43. N. Yan, P. Ni, and M. Zhang, Preparation and properties crocapsules with non-ionic surfactant as emulsifier. J. Microencaps., 10:375383 (1993). on polyamide microcapsules containing 44. N. Yan, M. Zhang, and P. Ni, Study oily liquids, J. Microencaps., 11:365-372 (1994). 45. S. AlexandridouandC.Kiparissides,Productionofoilcontainingpoly(terephthalamide) microcapsulesby interfacial polymerisation: Effectof process variableson the microcapsule sue distribution,J. Microencaps., 11:603614 (1994). 46. K. B. Lee, and T. Kondo,Permeabilitycoefficientofmicrocapsulemembrane, J. Microencaps., 199-356 (1984). on pH of permeabil47. E. Miyauchi, Y.Togawa, K. Makino, et al., Dependence poly(L-lysine-alt-terephthalic acid)microitytowardselectrolyteionsof capsule membranes, J. Microencaps., 9:329-333,1992. 48. M. C. Levy, S. Lefebvre, M. Rahmouni, et al., Fourier-transform infrared spectroscopic studies of human serum-albumin microcapsules prepared by interfacialcross-linkingwith terephthaloylchloride-influence of polycondensation pHon spectra and relation with microcapsule morphology and size, J. Pharm. Sci., 80578-585 (1991). 49. M. C. Levy, S. Lefebvre, M. C.Andry, et al., Fourier-transform infrared spectroscopic studies of cross-linked human serum-albumin microcapsules. 2. Influence of reaction-time on spectra and correlation with microcapsule morphology and size, J. Pharm. Sci., 83:419-422 (1994). 50. F. Edwards-Levy, M. C. Andry, and M. C. Levy, Determination of free amino group content of serum-albumin microcapsules using trinitrobenzenesulfonic acid-Effect of variations in polycondensation pH, Int. J. Pharm., 96:85-90 (1993). 51. F. Edwards-Levy, M. C. Andry, and M.C. Levy, Determination of free amino group content ofserum-albuminmicrocapsules.2.Effectof variationsin reaction-timeandinterephthaloylchlorideconcentration,Int.J.Pharm., 103:253-257 (1994). 52. G . Y. Chai and W. B. Krantz, Formation and characterization of polyamide membranes via interfacial polymerization, J. Membr. Sci., 93:175-192 (1994). 53. H. S. Tan, T. H. Ng, and H.K. Mahabadi, Interfacial polymerization encapsulation of a viscous pigment mix-emulsification conditions and particle-size distribution, J. Microencaps., 8525-536 (1991). 54. B.L.Duel1 and D. B. Pendergrass, Microencapsulation by interfacial polymerization ofpolyfunctionalAziridines,AbstractPapersAm.Chem.Soc., 208(Pt. 2):336-P0ly (1994).

Microcapsules Polymerization by and 55.

56. 57. 58.

59. 60.

61.

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67. 68. 69. 70. 71.

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A. T. Florence, T. L. Whateley, and D. A. Wood, Potentially biodegradable poly(alky1-2-cyanoacrylate) membranes, Pharm. J. microcapsules with Pharmacol., 31:422-424 (1979). P. Couvreur, Poly(cyanoacry1ates) as colloidal drug camers, CRC Crit. Rev. Ther. DrugCamer Sys., 51-20 (1988). P. Couvreur and C. Vauthier, Poly(alkylcyanoacry1ate) nanoparticles as drug carriers: Present state and perspectives, J. Controll. Rel., 17:187-198 (1991). J. Kreuter,C. G. Wilson,J. R. Fry, et al., Toxicityandassociationof poly(cyanoacry1ate) nanoparticles with hepatocytes, J. Microencaps. 1:253257 (1984). of F. Leonard, R.K. Kulkami, G . Brandes, et al., Synthesis and Degradation Poly(alky1 cyanoacrylates) J. Appl. PolymerSci., 10:259-272 (1966). S. D. Troster and J. Kreuter, Influenceof the surface properties of low contact angle surfactants on the body distributionof C-l4 poly(methy1 methacylate) nanoparticles, J. Microencaps. 9,19-28 (1992). I. Fichtner,Influenceofpoly(buty1P. Beck,J.Kreuter,R.Reszka,and on the efficacy and toxicity of the cyanoacrylate) nanoparticles and liposomes anticancerdrugmitoxantroneinmurinetumourmodels,J.Microencaps., 1O:lOl-114 (1993). T. Harmia, P. Speiser, and J. Kreuter, A solid colloidal drug delivery system for the eye: Encapsulation of pilocarpine in nanoparticles, J. Microencaps., 3~3-12(1986). T. Harmia-Pulkkinen, A. lbomi, and E. Kristoffersson, Manufactureof poly(alkylcyanoacrylate) nanoparticles with pilocarpine andbytimolol micelle polymerisation: Factors influencing particle formation, J. Microencaps., 6237-93 (1989). M. J. Alonso, A. Sanchez, D. Torres, et al., Joint effects of monomer and stabiliser concentrations on physico-chemical characteristics of poly(butyl-2cyanoacrylate) nanoparticles, J. Microencaps., 7517-526 (1990). S. J. Douglas, S. S. Davis, and L. Illum. Nanoparticles in drug delivery, CRC Crit. Rev. Ther. Drug Carrier Syst., 3,233-261 (1987). et al.,Evaluationofpoly(alky1G. Puglisi, G . Giammona,M.Fresta, cyanoacrylate) nanoparticles as a potential drug camer: Preparation, morphologicalcharacterisationandloadingcapacity,J.Microencaps.,10:353-366 (1993). K. Shankland and T. L. Whateley, Particle size distribution of polycyanaoacrylate nanoparticles, J. Pharm. Pharmacol., 41:133P (1989). K. Shanklandand T. L.Whateley,Determinationofrefractiveindicesof polycyanoacrylate particles, J. Coll. Interface Sci., 154:160-165 (1992). F. LimandA.M. Sun, Microencapsulated islets as bioartificial endocrine pancreas, Science, 210:908 (1980). A. M. Sun, I. Vacek, andI. Tai, Microencapsulation of living cells and tissues, in Microcapsules and Nanoparticles in Medicine and Pharmacy (M. Donbrow, ed.), 1992, pp. 315-322, CRC Press, Boca Raton, Florida. H.Uludag,L.Kharlip,and M. V. Sefton,Proteindelivery bymicroencapsulated cells, Adv. Drug Del. Rev., 10:115-130 (1993).

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B.Dupuy,A.Arien,andA. P. MinnotFT-IRofmembranesmadewith alginate/polylysine complexes-variations with the mannuronic or guluronic 22,71content of the polysaccharides, Artif. Blood Subst. Immobil. Biotech. 82 (1994).

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K. K. Kwok, M.J. Groves, and D.J. Burgess, Production of5-15 micrometre diameter alginate-polylysine microcapsules by an air-atomization technique, Pharm. Res., 8:341-344 (1991). 74. J. Stange, E. Brugmann, D. Falfenhurst,et al., A new technique for encapsulation of liver microsomes, in Microencapsulation of Drugs(T. L. Whateley, ed.), Harwood Academic Press, Reading, UK,1992, pp. 257-262. 75. L. S. C. Wan, P. W. S. Heng, and L.W. Chan, Drug encapsulation in alginate microspheres by emulsification,J. Microencaps., 9:309-316 (1992). et al., An artificial endocrine 76. A.M. Sun, W. Parisius,H.G.Macmorine, pancreascontainingculturedislets of Langerhans,Artif.Organs, 4:275 (1980). 77.

M. Y. Fan, Z. P. Lum, X. W. Fu, et al., Reversal of diabetes in BB rats by transplantationof encapsulated pancreatic islets, Diabetes,39519 (1990). 78. Z.Cai, Z.Shi, M. Sherman, andA.M. Sun, Development and evaluation of asystemofmicroencapsulationofprimaryrathepatocytes,Hepatology, 10:855 (1989). 79. V. Coromili andT. M. S. Chang, Polydisperse dextran as a diffusing test solute

to study the membrane permeability of alginate polylysine microcapsules, Biomat. Artif. Cells Immobil. Biotech.21,427-444 (1993). 80. G . M. R. Vandenbossche, P. Vanoostveldt and J. P. Remon, A fluoroescence method for the determination of the molecular weight cut-off of alginate polylysine microcapsules, J. Pharmac. Pharmacal. 43,275-277 (1991). 81. G. M. R. Vandenbossche, P. Vanoostveldt, J. Demeester and J. P. Remon, The molecular weight cut-off of microcapsules is determined by the reaction 42, 381-386 betweenalginateandpolylysine,Biotechnol.andBioeng. (1993). 82. S. M. Shimi, E. L. Newman, D. Hopwood, and A. Cushieri, Semi-permeable microcapsulesforcellculture:Ultra-structuralcharacterisation, J. Microencaps., 8:307-316 (1991). 83. A. S. Sawhney, C. P. Pathak,and J. A.Hubbell,Interfacialphotopoly-

merization of poly(ethy1ene glycol)-based hydrogels upon alginate poly(L lysine) microcapsules for enhanced biocompatibility, Biomaterials, 14:10081016 (1993).

H. A. Clayton, N. J. M. London, P. S. Colloby, et al., The effect of capsule J. Micomposition on the biocompatibility of alginate/poly(l-lysine) capsules, croencaps., 8:221-233 (1991). 85. M. E. Pueyo, S. Darquy, F. Capron and G. Reach. In-vitro activation of humanmacrophagesbyalginatepolylysinemicrocapsules, J. Biomat.Sci. Palp. Ed., 5,197-203 (1993). 86. G. M. R. Vandenbossche, M. E. Bracke, C. A. Cuvelier, H. E. Bortier, M. M. Mareel, andJ. P. Remon, Host reaction against empty alginate polylysine 84.

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14 Lipospheres for Controlled Delivery of Substances Abraham J. Domb* and Lev Bergelson

TheHebrew Universityof Jerusalem, Jerusalem, Israel Shimon Amselem Phurmos Ltd., Weizmann IndustrialPark, Rehovot, Israel

Introduction I.

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111. Physical Characterization of Lipospheres A. Morphology B. Structure C. Particle Size D. Viscosity and Drug Loading E. DrugDistribution F. InVitroDrugRelease

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IV. Applications of Lipospheres A. Parenteral Delivery of Local Anesthetics B. Parenteral Delivery of Antibiotics C. Parenteral Delivery of Vaccines and Adjuvants D. Topical Delivery of Insect Repellent E. Nanolipospheres for Cell Targeting of Anticancer Drugs

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*Affiliated with the David Bloom Center for Pharmacy, Jerusalem, Israel.

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INTRODUCTION

Many dispersion systems are currently in use as carriers of biologically active compounds. Dispersion systems used for pharmaceutical and cosmetic formulations can be categorized as either suspensions, emulsions, and dispersions. Suspensions are defined as solid particles ranging in size from a few nanometers up to hundreds of micrometers: dispersed in a liquid medium using suspending agents. Solid particles include microspheres, microcapsules, and nanospheres [l]. Emulsions can be defined as dispersions of one liquid in another, stabilized by an interfacial film of emulsifiers such as surfactants and phospholipids. Despite their long history, emulsions are used less often today than many other dosage forms owing to their inherent instability. Emulsion formulations include water-in-oil and oil-in-water dispersions, multiple water-in-oil-in-water emulsions, and microemulsions [2]. Lipid dispersions such as liposomes are defined as phospholipidvesicles, and they are obtained by dispersing phospholipids in aqueous medium. Liposomes may be unilamellar or multilamellar, depending on the number of lamellae or bilayers, separated each from the other by a water domain. Multilayer liposomes consist of a highly ordered assembly of concentric phospholipid membranes or bilayers entrapping internal aqueous compartments [3]. The rigidity and permeability of phospholipid bilayers can be adjusted by including other water-insoluble components such as sterols and amphiphilesin addition to thephospholipid matrix. Lipospheres represent a new type of fat-based encapsulation system developed forparenteral and topical drug delivery of bioactive compounds [4-lo]. Lipospheres consist of water-dispersible solid microparticles of particle size between 0.2 to 100 pm in diameter composed of a solid hydrophobic fatcore (triglycerides) stabilized by one monolayerof phospholipid molecules embedded in their surface. The internal core contains the bioactive compound dissolved or dispersed in the solid-fat matrix. The liposphere system has beenused for the controlled delivery of various types of drugs, including anti-inflammatory compounds, local anesthetics, antibiotics, and anticancer agents. They have alsobeen used successfullyas carriers of vaccines and adjuvants [7-lo]. Similar systems based on solid fats and phospholipids have beendescribed recently [U-131. Lipospheres have several advantages over other delivery systems, including emulsions, liposomes, and microspheres, forexample, better physical stability, low costof ingredients, ease of preparation and scale-up, high dispersability in an aqueous medium, high entrapment of hydrophobic

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drugs, controlled particle size, and extended release of entrapped drug after a single injection from a few hours to several days. This chapter describes the preparation and physicochemical properties of lipospheres and their use for the parenteral delivery of various drugs and vaccines and the topical administration of active agents. 11.

PREPARATION OF LIPOSPHERES

In contrast to certain oil emulsions, the liposphere approach utilizes naturally occumng biodegradable lipid constituents. The internal hydrophobic core of lipospheres is composed of fats, mainly solid triglycerides, whereas the surface activity of liposphere particles is provided by the surrounding lecithin layer composed of phospholipid molecules. The neutral fats used in the preparation of the hydrophobic core of the liposphere formulations described here include tricaprin, trilaurin, and tristearin, stearic acid, ethyl stearate, and hydrogenatedvegetable oil. The polymers used in the preparation of polymeric biodegradablelipospheres were low molecular weight poly(1actic acid) and poly(capro1actone). The phospholipids used to form the surroundinglayer of lipospheres were pure-egg phosphatidylcholine, soybean phosphatidylcholine, dimyristoyl phosphatidylglycerol, and phosphatidylethanolamine. Food-grade lecithin (96% acetone insoluble) was used inthe preparationof lipospheres for topioal and veterinary applications. Liposphere formulations are prepared by a solvent or melt process. In themelt method, theactive agent is dissolved or dispersed in the melted solid camer; that is, tristearin or polycaprolactone and a hot buffer solution is added at once along with the phospholipid powder. The hot mixture is homogenized for about 2-5 min using a homogenizer or ultrasound probe, after which a uniform emulsionis obtained. The milky formulation is then rapidly cooled down to about 20°C by immersing the formulation flask inan acetone-dry ice bath while homogenizationis continued to yield a uniform dispersion of lipospheres. Alternatively, lipospheres might be prepared by a solvent technique. In this case, the active agent, the solid camer, and phospholipid are dissolved in an organic solvent such as acetone, ethyl acetate, ethanol, or dichloromethane. The solvent is then evaporated and.tberesulting solid is mixed with warm buffer solution and mixing is continued until a homogeneous dispersion of lipospheres is obtained. In a typical preparation, dexamethasone(200mg), tristearin (400mg), and propylparaben (5 mg) are added to a 50-mL round-bottom glass flask. The flask isheated at75°C to melt the tristearin-dexamethasone mixture and

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hot 0.1 M phosphate buffer solution pH 7.4 (75"C, 9.3 g) isadded along with egg phosphatidylcholine (200 mg). The mixture is homogenized for 2 min until a uniform milky formulation is obtained. The hot formulation is rapidly cooled to below 20°Cby immersing the flask in adry ice-acetone bath with continued mixing to yield a white, thin dispersion.If needed, the pHof the formulation is adjusted to 7.4 with a 1N HC1 solution. The formulation may contain antioxidantssuch as tocopherol and preservatives such as parabens. Submicron-size lipospheres are prepared by passing (four times) the liposphere formulation by extrusion through a submicron series of filters at a temperature 5°C above the melting point of the liposphere core composition. Particlesize maybe reduced to about200 nm. Polymeric biodegradable lipospheres can also be prepared by a solvent or melt process. The difference between polymeric lipospheres and the standard liposphere formulations is the composition of the internal core of the particles. Standard lipospheresas those previously described consist of a solid hydrophobic fat core composed of neutral fats like tristearin, whereas in the polymeric lipospheres, biodegradable polymers such as polylactide or polycaprolactone substituted the triglycerides. Both typesof lipospheres are thought to be stabilized by one layer of phospholipid mole111). cules embedded in their surface (see Section Sterile liposphere formulations are prepared by sterile filtration of the dispersion in the hot stage during preparation through a filter at 0.2 pm a temperature5°C above themelting point of the liposphere corecomposition. Heat sterilization using a standard autoclave cycle decomposed the formulation. y-Sterilization of liposphere formulationsdid not affect their physical properties. Lipospheres formulations of 1 : 4 : 2 and 2 : 4 : 2 bupivacaine-tristearin-phospholipid(w/w % ratio) were irradiated with a samples were analyzed forparticlesize, dose of 2.33 Mrad,andthe bupivacaine content, in vitro release characteristics, and in vivo activity. The irradiated formulations had a similar particle size, bupivacaine content, release rate, and anesthetic effectiveness in the rat paw analgesia model to bupivacaine HCl solution (Marcaine). However, a more careful analysis of the formulation ingredients should be performed, since phospholipids may degrade during irradiation [14]. 111.

PHYSICAL CHARACTERIZATION OF LIPOSPHERES

A. Morphology Fig. 1 shows a light microscopic picture of a typical preparation of lipospheres composed of tristearin and lecithin consisting of particles having a uniform spherical shape. Microscopic examination of a typical liposphere

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Fig. 1 Light microscope pictureof a typical preparationof lipospheres composed of tristearin and lecithin with average particlesize of 10 pm.

Fig. 2 Transmission electron microscopy (TEM) of a single nanoliposphere (200 nm) .

formulation using transmission electron microscopy (TEM) showed spherical particles (Fig. 2). B. Structure

The phospholipid content on the surfaceof lipospheres was determined by 31Pnuclearmagneticresonance (NMR) before andaftermanogenase

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(Mn”) or proseodimium(Prt3) ion complexation [l51 and by trinitrobenzenesulfonic acid (TNBS) labeling using liposphere formulations containing phosphatidylethanolamine (PE) [16]. In both methods, the agents (Mn+2, Prt3, or TNBS) interact specifically with the exposed phosphateor amino groups of the phospholipid. Determination of the surface phospholipid using the TNBS methodshowed that 70-90% of the phospholipid polar heads are on the surface of the liposphere particles prepared from triglyceride-phospholipid at a 1 : 0.5 to 1 : 0.25 w/w ratio. Increasing the phospholipid content decreases the percentageof surface phospholipid polar heads, which indicates the formation of other phospholipid structures such as liposomesin the formulation. Similar results were obtained by 31P NMR analysis using Mn+’and Prf3ions for interaction with the phosphate polar heads [15]. This is illustrated in Fig. 3, which shows”P NMR spectra of drug-free liposphere vesicles obtained in the absence (A) and presence (B) of Mn+’. The decrease in the peak areaafter the addition of 5 mM Mn+’ represent therelative amount of phospholipid in the outer monolayer avail-

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Fig. 3 31PNMR spectra of lipospheres composed of tricaprin-phospholipid in a weight ratio of 2 : 1 in the absence (A) and presence (B) of Mnt2. 0 ppm corresponds with the resonance position of diacylphosphatidylcholine undergoing isotropic motion.

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able for interaction with Mn+2. For lipospheres composed of tricaprinphospholipid at a weight ratio of 2 : 1 and 5 : 1, 75 and 90% of the phospholipid polar heads wereaccessible to theparamagnetic ions, respectively. For comparison, liposomes of the same composition and size but without tricaprin had only 40% of the phospholipid polar heads on the surface [151. These datasuggest that the proposed structure of a liposphere is that of spherical particle with a monolayer of phospholipid molecules surrounding theinternal solid fatcore,wherethehydrophobic chains of the phospholipids are embedded in the internal triglyceride core containing the active agent, asillustrated in Fig. 4.

C. Particle Size Analysis of the particle size distribution of lipospheres was performed using a LS 100 Coulter Counter Particle Size Analyzer (Coulter Corp., Hialeah, FL). This instrument can measure particles from 0.4 to 800 pm by particle size-dependent light diffraction patterns. The particle size of submicron lipospheres was estimated from the transmission electron microscope pictures and using a particle size analyzer for 0.01- to 3-pm particle size diameter range. Blank and drug-loaded lipospheres prepared by the melt methodwithout farther treatment had a unimodal shape with an average particle size

Fig. 4 Schematic illustrationof proposed structureof a liposphere particle.

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between 5 and 15 pm with less than 2% of particles greater than 100 pm. This particle size formulation is useful for subcutaneous or intramuscular injection or fortopical applications. In an effort to reduce the particle size, lipospheres of 16.9 pm average diameter preparedby the melt method were passed through a laboratory Microfluidizer (Microfluidics, MA, USA). Thereby the average size was reduced to 10.8 pm after8 passes and to 9.4 pm after 40 cycles. Similar results were obtained when using a homogenizer. Lipospheres containing the malaria R32NS1 antigen and differingin their fat composition were prepared by the melt procedure [&lo]. Three groups of neutralfats were used: (1) solid fats such as tristearin and stearic acid with melting points in the rangeof 6570°C; (2) semisolid fats such as tricaprin and ethylstearate with melting points in the range of 30-35°C; and (3) liquid fats such as olive oil and corn oil. ?kro populations of liposphere particles usually coexisted, one in the size range of 1-10 pm in diameter (population A), and a second population with a diameter between10 and 100 pm (population B). No correlation was found between fat physical state (solid, semisolid, liquid) and particle size distribution of lipospheres formed. Lipospheresmade of tristearin were the most homogeneous formulations, with 100%of the particleshaving an average diameterof about 7 2 3 Pm 181. Biodegradable polymeric lipospheres made of polylactide and lecithin showed a very broad particle size distribution from 2 to 100 pm in diameter, with a mean average size of 30.6 & 25.9 pm and median (% of particles >50pm) equal to 21.6 [lo]. Polycaprolactone lipospheres showed a similar range of particle size distribution, but with a 1.5-fold mean average size (45.6 f 29.5 pm) and a doublemedian value (43.6) compared with polylactide lipospheres. Inclusion of lipid A in the composition of the polymeric lipospheres reduced their mean average particlesize by a factor of 0.25 regardless of the polymer type [lo]. All the liposphere formulations with a polymeric core prepared remained stable. during the3-month period of the study, and no phase separation or appearanceof aggregates were observed. D. Viscosity and Drug Loading

For drugs such as oxytetracycline [17], itraconazole, and dexamethasone, up to20% drug loading into lipospheres was obtained and the formulations were fluid enough to beinjected.Forotheragentslikebupivacaine, lidocaine, and chloramphenicol, drug loading above 10% produced aviscous lotion. Theviscosity of the liposphere formulation is dependent on the drug properties, the ionic strength andofpH the continuous aqueous solu-

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tion, and the ratio and amount of phospholipids and triglycerides. An increase in the contentof the insoluble ingredients (drug and lipids) and in the salt concentration of the aqueousmedium increases the viscosity of the formulation. For topical applications, lotion and paste consistencies are desired, which are achieved by adding NaCl to thewater phase and increasing the fatand phospholipid content of the liposphere lipid phase [M].

E. DrugDistribution The drugdistribution in a liposphere formulation was determined by isolating the particles by centrifugation and determiningthe drug contentin the isolated cake after extraction of the free drug with, for example, acidic buffer solution for basic drugs. The amount of unencapsulated free drug can be estimated by microscopic analysis(the free drug appearsusually as crystals, whereas lipospheres appear as round shapes). A study was conducted to de". Ane bupivacaine distribution in the 2% bupivacaine formulation [19]. It was found that the nonencapsulated ,drug was observed as needle-shaped crystals dispersed among the liposphere particles. These lipospheres anddrug crystals were isolated by centrifugation, thebupivacaine crystals were extracted from the mixturewith acidic buffer solution, and the drug-loaded lipospheres were analyzed for bupivacaine contept after dissolution-extraction of the lipospheres in Triton solution. The drug loading was in the rangeof 70-85 weight% in the core and about20% of the drug was extracted by the acidic solution appearingasnonincorporated drug. About 4% of the drug was soluble in the aqueous solution, which is the solubility of bupivacaine in the buffer solution. In an attempt to determine the form of the unincorporated drug, bupivacaine and phospholipid weredispersed in aqueous mediumin the absence of the tristearin component. A uniform and stable submicrondispersion was obtained. Microscopic examination of this fat-free preparation showed that the dispersed drug microparticles are nonspherical but in the form of long needles. It should be noted thatbupivacaine free base is not dispersible in buffer solution without asurfactant such as phospholipids. Thus, it is apparent that the unincorporatedbupivacaine in the tristearin liposphere formulation is in a form of dispersible microparticles composed of the solid drug and phospholipids. Taxol, verapamil, and piridoxamine wereincorporated in submicronsize lipospheres up to 90% encapsulation yield. F. InVitro Drug Release

In vitro release studies were conductedusing a large-pore dialysis tubing of 300,000 molecular weight cut-off (MWCO) to minimize the effect of the

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tubing on the release rate from the formulation (a regular pore size tubing of 12,000 MWCO did affectthe drug release rate). Thecontrol solutions of drugs such as bupivacaine (Marcaine-0.75%in solution for injection) were released through the large-pore tubing within a few hours. Drug releases from 2 and 5% bupivacaine-loaded liposphere formulations are shown in Fig. 5. Both formulations released the drug for 48 h following a first-order kinetics (R2=0.97).These release profiles are expected for formulations that contain 4% of the free drugsoluble in the aqueousvehicle which are released immediately throughthe dialysis tubing. The release of etoposide, a water-insoluble anticancer agent, from 2% loaded lipospheres placedin a dialysis tubing (300,000 MWCO) is shown in Fig. 6 . Over 90% of the drug was constantly released during a period of 80 h. Lipospheres containing I4C-diazepamwere preparedaccording to the melt technique. The lipospheres showed a very uniform particle size distribution with an average particle size of 8 pm. The lipospheres were evaluated in vitro by placing 0.5 mL of the formulation in a dialysis tubing (300,000 MWCO). The dialysis tubing was placed in a 0.1" phosphate

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Fig. 6 Release of etoposidefromalipospheredispersion.Totalamount etoposide in the test specimen was 6.2 mg.

of

buffer, and the cumulative release of diazepam was determined by measuring the radioactivity in the release medium. The release medium was changed periodically to provide sink conditions. Sustained release of diazepam was obtained over a period of 3 days. The in vitro release was also determined by mixing a sample of the formulation in excessbuffer solution, specimens of the mixture were taken every few hours, and the drug released into the solution was determined after removalof the lipospheres by ultracentrifugation. The release rate by this method was faster than from dialysistubing. IV. APPLICATIONS OF LIPOSPHERES This section describes the use of lipospheres for parenteral administration of long-acting local anesthetics and oxytetracycline and fortopical application of N,N-diethyl-m-toluamide(DEET) insect repellent.

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Parenteral Delivery of Local Anesthetics

Local anesthetics are preferred over general anesthetics because of the serious complications that can occur during general anesthesia. However, even local anesthetics, which are usually injected as an aqueous solution, are eventually absorbed from the site of application into the circulation. Frequent administration of local anesthetics may result in the development of systemic toxicity. A long-acting formulation that provides extended regional blockade may be useful for pain management following surgery or for chronic pain relief. Lipospheres wereused for the delivery of the common local anesthetics bupivacaine and lidocaine for the purpose of extending their effectiveness to a few daysafter a single injection. Liposphere formulations containing local anesthetics were prepared by the melt and solvent methods as described above. Sterile formulations wereprepared bydissolving bupivacaine (100 g) and egg phospholipid (100 g)in ethanol followed by sterile filtration through a 0.22-pm filter.Tristearin (200 g) was dissolved in hot ethanol andfiltered into the same flask and the ethanolwas evaporated to dryness. To the remaining semisolid, 5 L of hot (65°C) sterile 0.1" phosphate buffer solution containing 0.05% methyl paraben and 0.1% propyl paraben as preservatives was added and the mixture was homogenized for 5 min at high speed. The uniform milky preparation was rapidly cooled down to below 20°C by immersing the flask in a dry ice-acetone bath while homogenization continued. The formulation was filled into 10mL vials and stored underaseptic conditions at 4°C until used. A variety of models have been used for the evaluation of peripheral analgesic agents. However, only three have been extensively used to evaluate pharmaceutical compositions. There are the Randall-Selitto test, the abdominal construction writhing response to intraperitonealinjection of an irritant, and thepain response of mice after formalin injection [20,21]. The local effects as a function of time of liposphere formulations were tested using the Randall-Selitto experimental animal model. To induce hyperalgesia, animals were first anesthetized and then a yeast or carrageenan solution was injected through the foot pad nearest the first digit. The foot withdrawal score was then measured at the indicated times after the injection, on a scale from 0-25 (0 = could not stand any pressure; 25 = could stand extensive pressure) using the RandallSelitto instrument. Liposphere formulations to be tested were coadministered with the yeast solution. Control animals received an identical volume of water. For animals being tested for more than 24 h, the liposphere formulation was injected at time 0 and yeast was injected 24 h prior to

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liposphere injection, since yeast-induced hyperalgesia is often not measurable at times greater than 24 h postadministration. The hyperalgesic response was developed within the first hour after injection and maintained for 48 or 72 h. The effectiveness of several bupivacaine formulations was investigated. Liposphere formulations containing tristearin as the fat and phosphatidylcholine from eitheregg (PCE) or soybeans (PCS) and loaded with 1 or 2% bupivacaine produced long-lasting analgesia for at least 48 h and for almost 72 h [22]. The blank liposphere formulation did not produce any analgesia, and the Marcainecontrol solution (0.75% in saline) produced a strong analgesic effect which lasted for less than 3 h. A rat sciatic nerve preparation was developed to study prolonged local anesthetic blockade from bupivacaine lipospheres [23,24]. Blockade andor section of the rat sciatic nerve is a time-honored system to study regional anesthesia. The nerve is large and easily seen, and the effects of motor and sympathetic blockade in the foot can be detected. Sensory blockadecan also bemeasuredprovidedthere is no spillover in the saphenousnerve distribution. After anesthesia was induced, bilateral posterolateral incisions were made in the upper thighs, and the sciatic nerves were visualized. Sham vehicle wasinjected around the nerve onone side, and vehicle containing 5 and 10% bupivacaine (0.5-mL dose) was injected on the other side. The facia wasthen closed over the deepcompartment to restrict partially egress of drug formulation. Motor blockade was scored on a 4-point scale based on visual observation: 1, normal appearance; 2, impaired ability to splay toes when elevated by the tail;3, toes and foot remained plantar flexedwith no splaying ability; and 4, loss of dorsiflexion, flexion of toes, and impairmentof gait. Both 5 and 10% bupivacaine liposphere formulations showed significant levels of motor blockade throughday 3, and in some cases, day 4(Fig. 7). Motor function in all animals returned to normalby day 6. Sympathetic blockade was determined indirectly by foot pad temperature measurements. The foot receives sympathetic innervation largely from the sciatic nerve, although there may be a contribution from the saphenous nerve as well. Skin temperature measurementsof the blockadeside were also monitored as an indication of vascular tone. During the period when motor block was apparent (Fig. 7), the blockaded side was essentially always warmerthan the unblockaded side (Fig. 8). Temperature differences seemed to dissipate within 1 day after motor block resolved. Since sensory blockade measurementsin this experiment are complicated by several factors, it was important to find a method of detecting sensory blockade that is relatively independent of motor responses. Vocalizations in response to defined transcutaneous electrical stimulation of points on the feet were

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TIME (days after injection) Fig. 7 Motor blockade of 5% (n = 6) and 10% (n = 3) liposphere formulations injected near the sciatic nerve of rabbits. The motor rating were:4, clubbed foot; 3, partial clubbing (does not splay); 2, obvious difference in splaying; and 1, normal.

used as a criterion for the measurementof sensory blockade. The threshold to hind paw electric shock and hind paw pad temperature measures of sympathetic block where both increased for 3-4 days. No impairments were observed on the contralateral control side. One week after liposphere administration, sciatic nerves were removed and histologically evaluated. No evidence of nerve damage andvery little inflammation of foreign body response wereobserved. . A study was conducted to evaluate the efficacy of 2% bupivacaine liposphere formulation to produce analgesia inthe ratformalin model [24]. The model was designed to assess the antinociceptive ability against chronic pain caused by a test compound. An additional foot flick thermal method was also performed utilizing the sameanimals. The formalin study

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Time (days)

Fig. 8 Vocalizationthreshold to hindpawelectricalstimulationafterbilateral sciatic nerve administration. compared the effects of administration of 2% loaded bupivacaine lipospheres with blank lipospheres, standard bupivacaine solution (Marcaine, 0.5% i i t h 1 : 200,000 epinephrine), and physiological saline on antinociception in the rat. Rats were pretreated at various times with test or control formulations by infiltration injection into the right popliteal fossa and theninjected with 5% formalin into thedorsal surface of the right hind paw. Nociception wasthen measuredin the form of paw flinches ina 5-min period during both the acute and tonic phases. Both 2% bupivacaine liposphere and Marcaine had an onset of action times within 10 minutes. However, the liposphere formulation was able to maintain significant antinociception for at least 9 h in both acute and tonic phases as compared with less than 3 h for Marcaine. Similar results were obtained in the foot flick thermal stimulus model. Sensory blockade was measured by the time

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required for each rat to withdraw its hind paw from a 56°C plate. Latency to withdraw eachhind paw from the hot plate was recorded by alternating the hot plate within 15 S, the trialwas paws. If no withdrawal occurred from terminated. Neither the Marcaine nor liposphere formulation caused a change in pawvolume, nordid they significantly influencethe development of formalin-induced edema. The long-acting effect of the bupivacaine liposphere formulation as compared with Marcaine solution was further confirmed bythe rattail flick model [25]. The study compared administration of the 2% bupivacaine liposphere, blank liposphere, standard Marcaine, andsaline. After administration of the formulations, tail flick latencies weredetermined.The liposphere formulation exceeded the anesthetic duration of Marcaine by 12-fold. Treatment with blank lipospheres was not significantly different from treatmentwith the saline control. A pilot pharmacokinetic study was performed in rabbits to compare the Cmax andTmax values obtainedafter intramuscular injection of Marcaine HC1 solution and a liposphere formulation (Fig. 9). A total amount equivalent to of 20 mg bupivacaine was injected to rabbits (n = 3) and blood was collected for 72 h. Bupivacaine bloodconcentrations were determined by high pressure liquid chromatography (HPLC) following a United States Pharmacopeia (USP) method. Lidocainewas used as internal standard with a linear calibration curve for bupivacaine between 50 and 1000 ng/mL.Bupivacaine blood levels for bupivacaine solution (Marcaine) and lipospheres after a single injection are given in Fig. 9. The maximal blood concentrations were 681 f 246 and 200 & 55 ng/mL for bupivacaine in solution and in lipospheres, respectively. The toxicity of bupivacaine lipospheres in rats was evaluated. Since the liposphere formulation consists of natural inert components, phospholipids, and triglycerides, they are expected to be biocompatible in vivo [26]. The incidence of microscopic observations after intramuscular injection of liposphere formulations was studied (Table 1). Blank lipospheres, 1% and 5% bupivacaine in lipospheres, 5% dextrose solution, and bupivacaine solution (0.1 mL) were injected in rats followed by histological examination of the site of injection at days 3, 7, and 14. The degree of inflammation, necrosis, and fibrosis was scaledfrom 0 to 4, where 0 means absent and 4 means marked. The degree of inflammation, necrosis, and fibrosiswas similar for all formulations. At day 3, some irritation and inflammation was observed, which was reduced after 14 days. In a second study, the local toxicity of a 2% bupivacaine formulation after daily injections for 2 weeks in dogs was estimated. Minimal local irritation was observed in these studies as determined by histological examination.

=

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400

300

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48

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72

Time (hours)

Fig. 9 Bupivacainebloodlevelsafterintramuscularinjection of 2% loaded lipospheres as compared with bupivacaine HCl solution.Drug levels in blood were determined by HPLC. B. Parenteral Delivery of Antibiotics

Several antibiotics, including ofloxacin, norfloxacin, chloramphenicol palmitate, and oxytetracycline, and antigungal agents, such as nystatin and amphotericin B, have beenincorporated into lipospheres in highencapsulation yield. The use of lipospheres for antibiotic delivery was demonstrated by the development of liposphere oxytetracycline formulations for veterinary use [17]. Parenteral oxytetracycline (OTC) therapy in farm animals requires daily administration of the drug over several days, usually 3-5 days, in order to provide prolongedtherapeutic blood levels. Serum OTC concentrations of potential clinical and therapeutic values in the treatmentof OTC-sensitive organisms are estimated in the rangeof 0.15-1.5 mg/mL. The minimum inhibitory concentration (MIC) in milligramsper milliliter for certain patho-

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Table 1. Incidence of Microscopic Observations After Livosvhere Administration

Formulation Blank lipospheres(0%) inflammation necrosis fibrosis Bupivacaine lipospheres(1%; inflammation necrosis fibrosis Bupivacaine lipospheres(5%) inflammation necrosis fibrosis Dextrose solution inflammation necrosis fibrosis Bupivacaine HC1 solution (0.75%) inflammation necrosis fibrosis

1Day

3 Days

14 Days

112 222 000

221 010 110

212 000 221

111 222 000

212 313 101

211 OOO 110

112 333 000

222 212 111

211 000 200

111 212 OOO

211 011 111

101 000 101

111 223 OOO

222 211 111

OOO

111

111

Rats (groups of three) were injected subcutaneously with either formulation and the injection site was histogically examined. The results for eachare ratgiven in the table.Liposphereformulationswerecomposed of bupivacaine-tristearin-phospholipid at a1:2:1 weight ratio. Dextrose used was 5% a solution, and bupivacaine HCI solution was commercial Marcaine solution(0.75% bupivacaine). The degree of inflammation, necrosis, and fibrosis was scored from 0 to 4, where 0 means absent and4 means marked. gens of farm animals are 0.15 for Pasteurella multocida, 0.3 for Staphylococcus and Pasteurella anatipestifer, 0.4 for Haemophyllis paragallinarum, 0.8 for Mycoplasma gallisepticum, and 1.5 for Escherichia coli. Blood levels of above 0.5 mg/mL are required for treatment of most bacterial infections. Several long-acting oxytetracycline formulations have been reported [27-291. These formulationshave been tested in various farm animals and showed adequate blood levels for 72 h after asingle injection at adose of 20 mg/kg* Oxytetracycline was encapsulated in good yields in solid triglyceride liposphere formulations. The microdispersion containing up to 15 wt% oxytetracycline was injectable through a 20-G needle and was stable for at least 1year when stored under refrigerated conditions.OTC was released

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in vitro from dialysis tubing for 5 days, whereas OTC was released from solution through the tubing in less than 6 h. Blood levels in turkey birds or rabbits were maintainedfor up to4-5 days for the8% loaded formulation. An increase in OTC concentration required a decrease in the content of triglyceride and phospholipid in the formulation resulting in a decrease in the durationof drug release. The formulation degraded in tissue, but remnants of the formulations remained for periods longer than 4 weeks. A study was conducted to determine theeffect of liposphere composition on its physical properties, drug release rate, and injectability through a 22-G needle. The parameters investigated were triglyceride composition, phospholipid type, aqueous phase composition, drug loading, and OTCtriglyceride : phopholipid ratio. Three triglycerides were used, tristearin (mp 75"C), trilaurin (mp 42"C), and tricaprin (mp 33°C). Three types of lecithin were utilized, highly pure (99%)egg and soybeanphospholipid and foot-grade lecithin (96% acetone-insoluble material). The continuous medium was either 0.1 M phosphate buffer or 5% dextrose solution. The formulations based on tricaprin, regardless of the type of phospholipid or aqueous medium composition, loaded with up to 15% OTC were stable and injectable and provided a prolonged drugrelease rate in vitro. Formulations containing less than 5% phospholipid or more than 15%OTC were less stable and often caused cloggingof the needle, and therefore they were discarded [171. Release studies were conducted in dialysis tubing with a molecular weight cut-off of 300,000 (Spectrum, CA)or without tubing. One milliliter of liposphere formulation was introduced into the prewashed dialysis tubing and placed in a jar containing 800 mL of 0.1 M phosphate buffer pH 4.5. Alternatively, 1mL dispersion was added directly to thebuffer solution and placed on anorbital shaker at 37°C. Samples were takenat discrete times, centrifuged, and analyzed by ultraviolet (UV) spectrophotometry at 365 nm to determine the drug release rate from the formulations. A commercial OTC solution (Dabecycline, 10% OTC concentration) used as control was placed in the dialysis tubing to determine if the tubing was limitingthe rate of release. The release was slower from thedialysis tubing; however, OTC was released from the formulation for about 3-5 days from both releasing systems. Two animal studies were conductedin order toevaluate the controlledrelease effect of the liposphere formulations by following the OTC blood levels and theelimination of the administered dose fromthe injection site. In the first study, four OTC-loaded liposphere formulations differing in their compositions were compared with an OTC solution (10% OTC in acidic solution) used as reference and a blank liposphere formulation as control (Table 2). The formulations were injected intramuscularly to groups of six

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OTC blood levelswere determined. The injection sites turkey birds and the were observed forresiduals 7,11, and 28 days postinjection. The average OTC blood levels for the six formulations are given in Fig. 10. All four liposphere formulationsshow OTC levels above MIC for at least 3 days. Formulation E, composed of trilaurin (80 mg/mL) and phospholipid in a ratio of 2 : 1, showed similar OTC blood levels to the formulations based on tristearin and the more concentrated (120 mg/mL) trilaurin-based formulation. The formulation composed of trilaurin-phospholipid in a 1 : 1 ratio was less effective. These results indicate that a lower phospholipid to triglyceride ratio improves the duration of drug release and that higher drug loading (formulation C) does not affect the formulation effectiveness. Although tristearin showed good results, it is not preferred, because itis less susceptible for elimination from the injection site, as discussed below.

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The residual amountsevaluatedat7 and 11 days were maximal (about 90% of the original dose) for tristearin-based formulation (A), about 50% for the trilaurin-based formulations (B, C, and E), and about 20%.for theblank formulation. The deposits contained less than 10% OTC of the original dose. After 28 days, only formulation A had significant amounts of deposits at the injection site, which was mostly tristearin. No OTC was detected in any of the deposits retrieved from theanimals after 28 days. These results indicate that formulations based on trilaurin, asemisolid fat at body temperature (mp = 44°C) are more rapidly eliminated from the injection site[17]. Based on these results, second a animal study was conducted in order to identify an optimal formulation which wouldprovide maximal OTC blood levels after a single injection. The composition of formulation E (Table 2) was tested in comparison with a mixture of formulation E and DabecyclineOTC solution in a 4 : 3 volume ratio. Dabecycline-OTC solution was usedas reference. Dabecycline was added to formulation E in order toincrease the initial blood levels, which were too low for the first 5 h after injection.The three formulations were administered intramuscularly to turkey birds (10 birds foreach formulation). The OTCsolution (Dabecycline 100) resulted in very high serum levels at 3and 6 h, 19.9 and 3.5 mg/mL, respectively, and drug levels of 2.1, 0.4, and <0.1 mg/mL were found at 24, 48, and 72 h, respectively (Fig. 11). The mixture of Dabecycline and the liposphere formulation gave intermediate blood levels during the first day and higher levels 11days than lipospheresalone for 96 h. Evaluation of the injection sites after showed localizedresidues which were estimated as 40-50% and 30-40% of the injected dose for the liposphere and the Dabecycline liposphere formulations, respectively. The residue was mostlythe triglyceride and phospholipid of OTC. No residuals were found in the injection and a substantial amount site of OTC solution (Dabecycline). All animals in these studies were healthy and gained weight as the nontreated animals with no pathological signs. In all injection sites there were no signs of damage, swelling, or inflammation. No injection sites showed any necrosis or encapsulation even when precipitates were observed. C. Parenteral Deliveryof Vaccines and Aduvants

Severalreports describing the improvement of theimmuneresponse achieved by the association of antigens with lipidcamers such as liposomes [30] or microparticles like polymeric biodegradable microcapsules [31,32] have been published. The ability of these delivery systems to enhance

Domb et al.

400 8-

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0

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Dabecycline

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40

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Time (hours)

Fig. 11 Oxytetracyclinebloodconcentrations of turkeybirdsafteradministration of OTC solution, OTC lipospheres,and a mixture of OTC solutionand liposphere formulations. immunogenicity was related to the physicochemical characteristics of the particles. Lipospheres represent a new type of fat-based encapsulation that have been also used successfully as carriers of vaccines and adjuvants [8101. The physicochemical properties and immunogenic activity of different liposphere vaccine formulations containing a recombinant malaria antigen, R32NS1, derived from the circumsporozoite protein of Plasmodium falciparum as the model antigen have been described. The manufacture of lipospheres was accomplished by gently melting the neutral fat in the presence of phospholipid and dispersing the mixture in an aqueoussolution of the antigen by vigorous shaking, which results on cooling in the formation of a phospholipid-stabilized solid hydrophobic fat core containing the antigen [9].

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The effect of the type of fat usedin the preparation of lipospheres on their immune response to encapsulated antigen was tested. Mice were immunized twice at weeks 0 and 4 with lipospheres containing R32NS1 malaria antigen. Although for all liposphere formulations, the first immunization at week 0 caused a very small immune response, however, after the booster injection, a very marked increased of mean immunoglobulin (IgG) antibody levels was observed for most of the liposphere vaccine formulation tested, and the immune response obtained remained very at highlevels of IgG antibodytiters even after the 12-week period of the experiment [8]. The most immunogenic liposphere formulation was the one made of ethylstearate, whereas lipospheres made of stearic acid showed thelowest IgG ELISA titers. The complete order of immunogenic activity (based on fat composition) of the liposphere formulations tested was ethylstearate > olive oil > tristearin > tricaprin > corn oil stearic acid [8]. No correlation between liposphere particle sue or fat chemicalcharacteristics and immunogenicity was found. It is worth noting that the IgG antibody ELISA titers obtained on immunizing rabbits with liposphere R32NS1 were superior to those obtained following similar immunizations with the freeantigen absorbed to alum, which showed no antibodyactivity at the sameantigen concentrations. It was previously shown that this antigen was also poorly immunogenic in humans when injected alone as an aqueous solution or when adsorbed onalum [33]. Incorporation of a negatively charged phospholipid, dimyristoyl phosphatidylglycerol (DMPG) in the liposphere lipid phase, caused asignificant increase in the antibody response to the encapsulated R32NS1 antigen [9]. Enhancement of immunogenicity by inclusion of charged lipids have also beenobserved with certain antigens inliposomes.Negatively charged liposomes produced a better immune response to diphtheria toxoid than positively charged liposomes [34]. However, when liposomes were prepared with other antigens, positively charged liposomes workedequally as well as those bearing a negative charge [34]. Further studies are needed to determine whethernegative charges in lipospheres have general abilities to enhance immunogenicity or whether, as with liposomes, charge effects are dependent onindividual antigen composition. An interesting correlation was observed betweenthe liposphere fat to phospholipid (FPL) molar ratio, particle size, and immunogenicity. Low F/PL ratios (50.75) were found to induce the formation of lipospheres of small particle size (70% less than 10 pm in diameter), and this apparently resulted in increased antibody titers [9].Among the ratios tested, a maximal level of IgG antibody production was obtained at a F/PL ratioof 0.75, whereas at larger ratios, decreased antibody production was observed. Although the reason for this phenomenon is unknown, apossible explana-

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tion may be theoccurrence of better antigen orientation and epitope exposures in the small lipospheres because of higher surface curvature. W Opopulations of particles usually coexist in vaccine-loaded liposphere formulations, one in the size range of 1-10 pm in diameter (population A) and a second population with a diameter between 10 and 80 pm (population B). As mentioned previously, the particle size distribution of lipospheres depends on the fat phospholipid to (FPL) molar ratio, and the immune response to liposphere-encapsulated R32NS1 was also dependent on the F P L ratio. The average size of the particles increases with increasing F/PL molarratio:Under conditions where theF P L ratio is high (22.5), the large particle population is predominant (approximately 80% of the particles had an averagesize of 73 pm) [9]. In order to examine the influence of different routes of administration of lipospheres on their immunogenicity, rabbits wereimmunized orally or parenterally (by subcutaneous, intraperitoneal, intramuscular, and intravenous routes) with lipospheres made of tristearin and lecithin (1 : 1 molar ratio) and containing the malaria antigen [S]. The immune response obtained was followed for a period of 12 weeks postimmunization. No antibody activity wasfound after oral immunizationin any of the individual rabbits immunized with the liposphere R32NS1 vaccine formulation. However, rabbit immunization by all parenteral routes tested resulted in enhanced immunogenicity with increased antibody IgG levels over the entire postimmunization period. The individual rabbit immune response shows that. immunization by subcutaneous injection was the most effective vaccination route among all parenteral routes of administration tested [S]. Incorporation of lipid A, the terminal portion of gram-negative bacterial lipopolysaccharide, in lipospheres increased significantly the immune response to R32NS1 malaria antigen resulting in double IgG levels compared with R32NS1 lipospheres lacking the lipid A. The adjuvanteffect of lipid A incorporated in lipospheres was observed evenafter 1600-fold dilution of the rabbit sera [g]. The adjuvant effect of different doses of lipid A in lipospheres was also examined by immunizing rabbits with lipospheres containing R32NS1 and prepared at different final concentrations of lipid A. A gradual increase .in IgG antibody titer with increasing lipid A dose was observed. The strongest antibody activitywas obtained with lipospheres containing 150 pg of lipid Nrabbit. Athigher lipid A dose(200 kg/ rabbit), a decrease in ELISA units was observed [9]. The preparation and use of polymeric biodegradablelipospheres as a studied. The potential vehicle for thecontrolled release of vaccines was also immunogenicity of polymericlipospheres composed of polylactide (PLD) or polycaprolactone (PCL) and containing the recombinant R32NS1 malaria

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antigen was tested in rabbits after intramuscular injection of the formulations [101. High levels of specificIgG antibodies were observedin the seraof the immunized rabbits up to 12 weeks after primary immunizationusing a solid phase ELISA assay. PCL lipospheres containing the malaria antigen were ableto induce sustained antibody activity after one single injection in the absence of immunomodulators.PCL lipospheres showedsuperior immunogenicity comparedwith PLD lipospheres, with the difference being attributed to thedifferent biodegradation rates of the polymers. D. Topical Delivery of Insect Repellent A common mean for repelling insects consists of applying the compound N,N-diethyl-m-toluamide(DEET) to theskin. The commercially available liquid DEET formulations contain between 15 and 100%DEET, and they are not recommended foruse on children. The toxicity of DEET has been extensively reported and related to its high skin absorption after topical administration [36-381. Previous studies have demonstratedthat as much as 50% of a topical dose of DEET is systematically absorbed [37,38]. In an effort to develop a new topical formulation for DEET thatpossesses reduced skin absorption as well as an increase in the durationof repellency, we have encapsulated DEET intolipospheres and studied its skin-absorption dynamics and durationof action [39,40].We hypothesized that theencapsulation of DEET will reduce its contact surface area with the skin and reduce its evaporation rate from theskin surface, resulting in reduced dermal uptake and extended repellent activity. The liposphere microdispersions containing DEET incorporated in solid triglyceride particles were prepared by the melt method using a m mon naturalingredients in one step without the use of solvents. The formulation was preserved by parabens, propylparaben in the oil phase, and methylparaben in the aqueous phase. The average particle size was in the range of 15 pm, and microscopic examination showed spherical particles. The formulations were stable for at least 1 year when stored at 4°C and 25°C in a closed glass container, and DEET content, particle size, and viscosity remained almost constant. The residual efficacyof liposphere formulations was evaluated on volunteers by applying the formulations to the skin and exposing the subjects to mosquitoes [40]. The time of 100% repellency (zero biting) was the index for determining the effectiveness of a formulation. The formulations were applied on four locations on the armof volunteers ata 2.5 mg/cm2on a total areaof 12-cm2skin surface. Mosquitoes were placed in a screen-bottomed (%mesh netting, 10-cm2exposure area) cylindrical cup containing 50 5- to 15-day old female mosquitoesdisplaying host-

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seeking behavior with access to the skin through the netting. The forearm was placed on the mosquito netting for 10 min every 30 min and the number of biting mosquitoes (evident by a blood meal) were recorded. Prior to any efficacy experiment, the mosquitoes weretested on untreated skin to confirm their host-seeking behavior. W Omosquito species were tested: Aedes aegypti and Anopheles stephemi, both aggressive biters. The results of this experiment is given in Table3.The formulations were repellent for a minimum of 2.5,3.5, and 6.3 h for the6.5, 10, and 20% DEETcontaining lipospheres, respectively. The DEET-freeformulation (control) and the untreated groupsdid not show any activity against Aedes aegypti, and the 10% DEET solution in alcohol was repellent for about 1.5 h. The blood concentrations after intravenous (IV) dosing and topical administration of 10% DEET in ethanol (group A) and 10% DEET in lipospheres (group B) were measured(Fig. 12). The AUC after dosing was 24,494 DPM/mL for group A (10% alcohol solution) and 8444 DPM/mL for groupB (liposphere formulation). Therefore, theabsolute bioavailabilTable 3. Repellency Effectivenessof DEET Liposphere Formulationsa

No.of biting Aedes aegypti and Anopheles stephemi % DEET in formulation

Time after application Untreatedb Controlb (min) 2

15 30

60

(4)

90 120 150 180 4 210 240 270 300 380

(5) 5 (4) 3(5) 3(5) 5 (5) 5 (3) 3 (5) 5 (3) 3 (4) 4 (6) 5 (6)

(4)

4 (0) 5 (0) 5 (3) 5 (3) 5 (5) 5 (5) 4 5 (3) 3 (4) 4 (3) 4 (5) 5 (5)

(2)

6.20 5

10

0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 1(1) 1(1) 1(2) 2 3 (2)

0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 0 (2) 0 (2) 1(2)

Referencec 0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 0 (0) 0 (0)

0 (0) 0 (0) 0 (0) 0 (0) 1(1) 0 (1) 1(2) 1(1) 2 (1)

a Cups with mosquitoes were placed for10 min every 30 min on the skin of volunteers treatedwith formulation(2.5 mg/cm2) and the number of bites were recorded. The results are averageof three independent tests (cup placements), the results in parenthesis areof Anopheles stephensi. Untreated group were without any application and control group was treated w DEET-free liposphere formulation. c Reference group received a10 wt% solution of deet in alcohol.

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c.

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I 0

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8

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Fig. 12 Bloodconcentrations(DPWmL)aftertopicaladministration of 10% DEET in ethanol or liposphere formulation administeredtopically (1 mL,20 pCi mL) to rabbits (surface area 64 cm’). Results representthe average of eight rabbits. ity of DEET from a 10% ethanol solution was 45%, whereas the bioavailability from DEET lipospheres was only 16%, a threefold reduction in the amount of DEET absorbed. About 74% of the IV administered dose was collected in the urine, and 39 and 19% of the topically administered doses were collected for the alcoholic and liposphere formulations, respectively. Assuming that the error in urine collection issimilar in all experiments, the difference in radioactivity contained in the urine after topical administration of the liposphere dosage form is about 50% of that of the alcoholic dose, which corresponds to theblood bioavailability calculations. The total amount of DEET recovered fromskin (washingof residual dose andextraction from skin) was similar for both formulations, indicating that both formulations were similarly exposed to the skin, and thus the results are comparable. The nonrecovered DEET after topical administration is probably due to evaporation of DEET from the nonocclusive patches to the environment. The lower DEET recovery of the liposphere formulation can be interpreted as more DEETis released to the environment resulting in more repellent activity [39].

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F. Nanolipospheres for Cell Targeting of Anticancer Drugs

The potential use of lipospheres loaded with paclitakel (Taxol) to overcome tumor cells’ acquired resistance to the drug was investigated [41]. It was assumed that, if absorbed by the cells, the encapsulation of Taxol willprevent the rapid expulsion of the drug outside the cell, and a sufficient cytotoxic level of drug concentration will be maintainedin cell plasma. %o liposphere formulations, one based on tricaprin and the other on polycaprolactone, were comparedwith liposomes of the same compositionbut without the core component(tricaprin or polycaprolactone). The formulations were preparedby the solvent method and extruded through a series of submicron filters to yield nanoparticles of 200-nmsize. The rateand amount of uptake of particles by cells wasdetermined using lipospheres or liposomes containing phosphatidylethanolamine fluorescein. The rate of uptake was followed by fluorescence microscopic visualizationof the amountof fluorescence accumulatedin cells or by measuring the amount of fluorescence in each cell usingthe FACS system. The formulations were incubatedwith the wild-type F98 glioma cellline or with the F98 cell lines resistant to 1 X 7.5 X and 8 X mMTaxol. The results indicate that (1) it takesabout 24 h of incubation of particles with cells to reach saturation of particle uptake, (2) cells accumulate higher concentrations of liposomes than lipospheres, and (3) cells which are more resistant to Taxol accumulate higher Taxol concentrations on incubation with Taxol liposomes and Taxol lipospheres than on incubation with the free drug. The cytotoxicity of liposphere- or liposome-encapsulated Taxol on the percentage of cell survivalof two celllines (wild type and F-98/1 X resistant cells) after 6 h of treatment with drug and further incubation of cells up to72 h was studied. F98 cells were incubated with a range of drug concentrations and different preparations for 6 h, then washed with fresh medium, and incubated for additional 66 h. The number of cells in each plate was counted daily. Thedata indicate that Taxol encapsulated in liposomes or lipospheres had a higher cytotoxic effect than freeTaxol. The results also demonstrate that although there is no significant difference betweenthe cytotoxic effect of free Taxol or Taxol encapsulated in liposomes on the wild-type cells,there is significantdifference in the effects on resistant cell lines. Taxolencapsulated in liposomes is about 30 and 50% more cytotoxic than free Taxol in cells resistant to and 10” m M Taxol, respectively. These preliminary results indicate that both lipospheres and liposomes were effective to overcome drug resistance, with the liposomes being moreeffective. The blood circulation time of nanolipospheres was compared with that of liposomes [42]. Lipospheres andliposomes of particle size between

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100 and 200 nm containing radioactive cholesteryl hexadecyl ether were injected to mice and the tissue distribution was determined by following the radioactivity content in blood, liver, and spleen at 1,3,8,and 24 h. The cholesteryl hexadecyl ether in organs was extracted with chloroform and the radioactivity in the extract was determined. The liposphere formulation was excreted faster thanliposomes and was concentrated in the liver rather than in the spleen for liposomes. To increase the retention time of lipospheres in the bloodcirculation, lipospheres were coatedwith polyethylene glycol (PEG) of 5000 molecular weight, which has been shown to increase blood circulation [43]. Liposphere formulation made from phospholipid containing 10% w/w phosphatidylethanolaminewere reacted with aldehyde-terminatedmethoxy-PEG to form PEG-coated lipospheres. The imine-bound PEG was reduced to the amine linkage, which is more stable. Preliminary in vitro experiments indicate that PEG-coatedlipospheres are stable in serum [42].

V. SUMMARY Lipospheres are solid, water-insoluble microparticles composed of a solid hydrophobic core having a layer of a phospholipid embedded on the surface of the core. The hydrophobic core is made of solid triglycerides, fatty acid esters, or bioerodible polymers containing the active agent. Liposphere formulations were effective in delivering various drugs and biological agents, including local anesthetics, antibiotics, vaccines, and anticancer agents with a prolonged activity of up to 4-5 days. The results presented in this study demonstrate that enhanced immunogenic efficacy can be achievedbyusing liposphere-based antigen formulations indicating the potential usefulness of lipospheres in the formulation of humanand veterinary vaccines. The liposphere approach employs a fat-lipid environment to achieve several goals: to serve as a carrier to protect the antigen, to serve as “depot,”and to provide a surface interphase necessary for adjuvant activity. It is reasonable to presume that the immunogenic and adjuvant activity of lipospheres may be due toa combination of factors. These factors may include a focused and enhanced delivery of the antigen to an antigenpresenting cell (macrophage) andprotection of the antigen from metabolic destruction at other sites in the body that do noparticipate in the immune response. The liposphere-delivery system as a fat-based adjuvant formulation may provide both the surface interphase necessary for solubilization and proper orientation of the adjuvant-active material and aspotential carriers for vaccines, which may allowbetter position for processing and presentation of the incorporated antigens resulting in enhanced immunogenicity.

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The feasibility of polymeric biodegradable lipospheres as carriersfor the controlled release of a recombinant malaria antigen was also demonstrated. The microencapsulated peptide induced efficient and potent primary immune responses. Polymeric lipospheres containing R32NS1 malaria antigen were able to induce very high levels of antibody activity after one single injection in the absence of immunomodulators. REFERENCES 1. M. Donbrow (ed.), Microcapsules and Nanoparticles in Medicine and Phar-

macy, CRC Press, Boca Raton, FL, 1992. 2. P. Becher (ed.), Encyclopediaof Emulsion Technology, Marcel Dekker, New York, 1985. 3. G.Gregoriadis(ed.),LiposomeTechnology, 2nd ed., Vol. 1, CRC Press, Boca Raton, FL, 1993, pp. 527-616. 4. A. J. Domb, Lipospheres for Controlled Delivery of Substances. U.S. Patent 5,188,837, Feb. 1993. 5. A. J. Domb and M. Maniar. Lipospheres forthe Delivery of Local Anesthetics. U.S. Patent, Allowed 1993. 6. A. J. Domb, Lipospheres for the Delivery of Insect Repellent. U.S. Patent, Allowed 1993. 7. A. J. Domb, Lipospheres for the Delivery of Vaccines, U.S. Patent Application, 1990, allowed August 1992. 8. S. Amselem, C. Alving, and A. Domb, Lipospheres for the deliveryof vac(S. Cohen andH. Berncines, in Microparticulate Systems for Drug Delivery stein, eds.), Marcel Dekker,New York, 1996 (in press). as a vaccinecamer 9. S. Amselem, A. J. Domb, and C. R. Alving, Lipospheres system: Effects of size, charge, and phospholipid composition, Vaccine Res., 1~383-395 (1992). 10. S. Amselem, C. R. Alving, and A. J. Domb, Polymeric biodegradable lipospheres as vaccine delivery systems, Polym. Adv. Technol., 3:351-357 (1992). 11. S. Amselem, A. Yogev, E. Zawoznik, and D. Friedman, Emulsomes, a novel drug delivery technology, Proceed. Int. Symp. Control. Rel. Bioact. Mater., 21 ~668-669 (1994). 12. S. Amselem, A. Yogev, E. Zawoznik, and D. Friedman, Emulsomes, a New of LipidAssembly,inNonmedicalApplicationsofLiposomes (Y. BarenholzandD.D.Lasic,eds.),CRCPress,BocaRaton, FL, 1995(in press). W. Mehnert, Incorporation of 13. R. H. Muller, C. Schwarz, A. zur Muhlen, lipophilic drugs and drug release profiles of solid lipidnanoparticles, Proceed. Int. Symp. Control. Rel. Bioact. Mater., 21:146-147 (1994). 14. S. lanzini, L. Guidony, P. L. Indovina, et al., y-Irradiation effect on phosphatidyl choline multilayer liposomes: Calorimetric, NMR and spectrofluorimetric studies, Rad. Res., 98:154-166 (1984).

me

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15. D. B. Fenske, Structural and motional propertiesof vesicles as revealed by nuclear magnetic resonance, Chem. Phys. Lipids, 64:143-162 (1993). 16. Y. Barenholz andS. Amselem, Quality control assays in the development and clinical use of liposome-based formulations, in Liposome Technology, 2nd ed., Vol. 1 (G. Gregoriadis, ed.), CRC Press, Boca Raton, FL, 1993, pp. 527-616. oxytetracycline-lipospheresformulations, 17. A. J. Domb, Long acting unjectable Int. J. Pharm., 124:271-278 (1995). 18. D. Hanibbal, M. Rock, and J. Knowles, DEET Bioavailability from lipospheres, Proceed. Intern. Symp. Control. Rel. Bioact. Mater., 19 (1992). pp. 446-447 19. M. Maniar, R. Burch, and A. Domb, In Vitro and In Vivo Evaluation of a Sustained Release Local Anesthetic Formulation,M P S Meeting, Washington, D.C., November 1991. 20. D. Dubuisson and S. G. Dannis, The formalin test: A quantitative study of the analgesic effectsof morphine, mepetidine, and brain stem stimulation in rats and cats, Pain, 4:161-174 (1977). 21. S. Haunskaar, 0.B. Fasmer, and K. Hole, Formalin test in mice, a useful techniqueforevaluatingmildanalgesics, J. Neurosci.Methods14:69-76 (1985). 22. A. J. Domb,Liposphereparenteraldeliverysystem,Proceed.Int.Symp. Control. Rel. Bioact. Mater.,20:121 (1993). 23. D. Masters and C. Berde, Drug delivery to peripheral nerves, in Polymer SiteSpecific Pharmacotherapy (A. J. Domb, ed.), Wiley, Chichester, UK, 1994, pp. 443-455. 24. D. Masters and A. J. Domb, Perinerve timed-release of local anesthetic and prolonged neural block from injectable liposphere formulations (submitted). 25. E. V. Hersh M. Maniar, M. Green, and S. A. Cooper, Anesthetic activity of the lipospheres bupivacaine delivery system in the rat, Anest. Prog., 39:197200 (1993). 26. M. J. Palham, Liposome phospholipid. Toxicological and environmental ad(0. Brown,H.C.Korting,and H. I. vantages,inLiposomeDermatics Maibach, eds.), Springer-Verlag, Berlin, 1992, pp. 57-68. 27. M.F.Landoniand J. 0. Errecalde, Tissue concentrations of a long-acting oxytetracycline formulation after intramuscular administration in cattle, Rev. Sci. Technol., 11:909-915 (1992). 28. M. Oukessou, V. Uccelli-Thomas, and P. L. Toutain, Pharmacokinetics and J. local toleranceof a long-acting oxytetracycline formulation in camels, Am. Vet. Res., 53:1658-1662 (1992). 29. D. A. Adawa, A. Z. Hassan, S. U. Abdullah, et al., Clinical trial of longacting oxytetracycline and peroxicam inthe treatment of canine ehrlichiosis, Vet. Q. 14:118-120 (1992). 30. C. R. Alving, Liposomes as carriers of vaccines, in Liposomes: From Biophysics to Therapeutics (M.J. Ostro, ed.), Marcel Dekker, New York, 1987, pp. 195-218.

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31. J. H. Eldridge, J.K. Staas, J. A. Meulbroek, et al., Biodegradable microspheres as a vaccine delivery system, Mol. Immunol., 28:287 (1991). 32. F. Stieneker, J. Kreuter,andJ.Lower,Highantibodytetersinmicewith polymethylmethacrylate nanoparticlesas adjuvant for HIV vaccines, AIDS, 5:431-435 (1991). 33. L. S. Rickman, D. M. Gordon, R. Wistar, Jr., et al., Use of adjuvant containing mycobacterial cell-wall skeleton, monophosphoryl lipidA, and squalene in malaria circumsporozoite protein vaccine, Lancet, 337:998-1001 (1991). 34. A. C. Allison and G. Gregoriadis, Liposomes as immunological adjuvants, Nature 252252 (1974). 35. T. D., Heath, D. C. Edwards, and B. E. Ryman, The adjuvant properties of liposomes, Biochem. Soc. Trans., 4:129 (1976). 36. J. R. Clem,D. F. Havemann,andM.A.Raebel.Insectrepellent(N,Ndiethyl-m-toluamide)cardiovasculartoxicityinanadult,Ann.Pharmacother., 27:289-293 (1993). 37. J. W. Lipscomb, J. E. Kramer, and J. B. Leikin. Seizure following brief exposure to the insect repellent N,N-diethyl-m-toluamide, Ann. Emerg. Med., 21~315-317 (1992). 38. H. L. Snodgrass, D. C. Nelson, and M. H. Weeks, Dermal penetration and potential for placental transfer of the insect repellent N,N-diethyl-m-toluamide, Am. Indust. Hyg. J., 43:747-753 (1982). 39. A. J. Domb,A.Marlinsky,M.Maniar,andL.Teomim,Insectrepellent formulationsofN,N-diethyl-m-touamide(DEET)inlipospheresystem,J. Am. Mosq. Control ASSOC., (1995). P. Patent, 40. A. J. Domb, Lipospheres for the delivery of insect repellent. U. 5,221,535 (1993). 41. A. Gur, Taxol Incorporated in Nanoliposphere Formulations Against Taxol ResistantCells,M.Sc.Thesis,TheHebrewUniversity,Jerusalem,Israel, 1994. 42. L. Lichtman-Teomim, Injectable systems for the delivery of insoluble anticanceragents,M.Sc.Thesis,TheHebrewUniversity,Jerusalem,Israel, 1994. 43. M.C. Woodle,M. S. Newman, andF. J.Martin, Liposome leakage and blood circulation: Comparison of absorbed block copolymers with covalent attachment of PEG, Intern.J. Pharm., 88:327-334 (1992).

15 Pharmaceutical Emulsions, Double Emulsions, and Microemulsions Nissim Garti and Abraham Aserin Casali Institute of Applied Chemisty, School ofApplied Science and Technology, The Hebrew University of Jerusalem, Jerusalem, Israel

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XII. Double Emulsions A. Definitions and Formation B. Stability and Transport Mechanisms C. New Approaches to Improve Stability D. Naturally Occurring Macromolecular Amphiphiles

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1. DEFINITIONS AND INTRODUCTION

An emulsion is prepared by the dispersion of one immiscible liquid in another, and then it is stabilized by a third component, the emulsifier [l]. The art and science which describe and characterize these systems, in practice and in theory, are known as emulsion technology. The need to bring two immiscible liquids into a prolonged intimate contact with a high surface area has long been a task of emulsion technology. Emulsions.have been widely used in many areas of application-in industry [2], agriculture [3-51, food [6-121, pharmaceuticals [13-151, and cosmetics [16-181. Their industrial uses have an extremely wide range of possible applications such as explosives [19]; petroleum [20], concrete [21,22], rubber, leather,textile [23,24], metal [25], and paper[26] products; and mineral flotation [27]. Pharmaceutical and medical applications are, however, the most studied and are used mainly for drug administration. Emulsions in which the water is the internal dispersed phase are termed water-in-oil emulsions (W/O), whereas emulsions in whichthe oil is the dispersed phase and thewater is the continuous phase are known as O/ W emulsions [28-331. More complex systems, in which one emulsion is further dispersed into another continuous phase, are called double emul-

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multiple (oil) drop internal aqueous drops surfactant films

external aqueous phase

Fig. 1. Schematic illustration of a double-emulsion droplet. sions, multiple emulsions, or emulsified emulsions [34-421. Two main types of double emulsions wererecognized and studied-the W/O/W and the O/ W/O double emulsions (Fig. 1). The droplet-size distribution of emulsion droplets (sometimes also termed macroemulsion) is 0.1-50 pm. A more common averagedroplets size is0.5-5.0 pm. The inner droplet size distribution of the W/O emulsion in the double emulsionis usually smaller than 0.5 pm, whereas theouter, external double emulsionis quite large and can exceed 10 pm (Fig. 2). Amphiphilic compounds,used to stabilize emulsions (the emulsifier), possess two distinct groups in the samemolecule, which differ greatly in their solubility relationships. Such compounds were termed “amphipathics”(or amphiphiles) by Hartley [43,44] to denote the presence of a lipophilic group having an affinity for the solvent and a second group having an affinity to water. The amphiphile has, using more picturesque terms, an hydrophilic “head” anda hydrophobic (lipophilic) “tail.” Amphiphiles tendto concentrate as a monolayer at the water interface and reduce the surface tension (therefore also termed surfactants or surface-active agents). Any excess surfactant that is not accommodated at the surface migrates to thebulk and forms aggregates known as micelles (or reverse micelles, if the bulk is organic solvent or oil) [28,30,31] (Fig. 3). The micelles can accommodate organic molecules intheir hydrophobic core or may be associated only with the amphiphile within the interface layer. The increased solubility of a compound associated with the formation of micelles or inverted micells has been termed by McBain [45] “solubilization.” The solubilized compounds, when partially hydrophilic, may partition between the continuous phase,the interior of the micelle, and the interfacial region. Thus, the interfacial region

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Drop Diameter (pm) Fig. 2. Size distribution of the internal aqueous (W/O emulsion) and multiple drops (W/O/W emulsion) of a multiple emulsion at the preparation stage.

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Fig. 3. Modes of surfactant action on liquid-air interfaceon solid-liquid interface and in the liquid bulk. 474

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may not consist entirely of the primary amphiphilic compound. Molecules that exhibit a substantial presence within the interfacial layers are sometimes called cosolvents or cosurfactants. These molecules are typically mediumchain alcohols. By changing the proportions of the surfactant to water and/or oil (constructing a complete phase diamgram), a micellar solution can progressively be converted into various other structures [46-501 (Fig. 4). The microstructures of the intermediate states are notfully understood. Depending on a large number of factors, including the structure of the amphiphile, its relative concentration, the temperature, and the presence orabsence of certain additives, several fascinating structures could be defined and characterized in the solution (Fig. 5). Some are isotropic solutions, amorphous in structure and mostly spherical in shape, called microemulsions, which are defined as S, (or L,) or S2 (or h)(depending on the continuous phase being water or organic solvent) [51-531. Some are isotropic solutions with a fluctuating structure (the continuous phase being oil and water alternatives) called bicontinuous structures or type I11 structures, and some are isotropic but more rigid in their structure and more viscous, appearing as gel-like, and these are called G and M, or M, phases or liquid crystalline phases. The intermediate liquid-crystalline phases may appear in a large variety of mesophases but usually in a given sequence. The most widely occurring liquid crystalline structure hasa lamellar structure [51-55] (Figs. 5 and 6 ) . The less-structured isotropic solutions, microemulsions, which are in essence “mixtures of aqueous solutions (sometimes with electrolytes), hy-

0

Fig. 4. Hypothetical phase regions of microemulsion system composed of oil (0),water (W), andsurfactant (S), showing a, inverted micelles; b, W/O microemulsions; c, cylinders; d, lamellae; e, O N microemulsion;f, normal micelles; and g, macroemulsion phases.

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Alcohol ”-.

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SPHERICAL MICELLES

Fig. 5. In a combination of water, a surfactant, and a long-chain alcohol, the areas for solutions of normal and inverse micelles are separatedby a micellar liquid crystal, drocarbons (‘oils’) and amphiphilic compounds,” have beenthe subject of extensive research (Fig. 7). The name microemulsions was introduced by Hoar and Schulman [56] to describe transparent or translucent systems obtained by titration of an ordinary emulsion havinga milky appearance to clarify it by the addition of medium-chain alcohol. There is muchdebate as to the term microemulsion beingused to describe such systems. Some prefer to call sucha system a swollenmicellar solution or solubilized micellar solution. However, microemulsions greatly differ from macroemulsions in the following characteristics: droplet sizes (the size ofmicroemulsions is less than 0.1 pm whereas the size of emulsions is 0.1 pm); appearance (transparent or translucent for microemulsions and milky for emulsions); thermodynamic stability (emulsions are unstable); and kinetic and time-dependent behavior (emulsion droplet size will grow with time and centrifugation); and modeof separation (microemulsionsare independent in the orderof mixing). Fig. 6. (a) M, mesophase. Rodlike micelles of indefinite length and arranged in hexagonal array. (b)G phase lamellar structures; sometimes called the Neat phase. of cubicpacking of (ModifiedfromRef.[56a])(c)Schematicrepresentation spheres. The spheres are generated by interaction of amphiphilic molecules and water.

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Fig. 7. Schematic structure of water-in-oiland oil-in-water microemulsions.

Although many investigators feel that one must distinguish between swollen micellar solutions and microemulsions in terms of the characteristics of the “reservoir” (the continuity of the oil) of the solubilized matter, there are no good instrumental methods to distinguish these differences. Thermodynamically speaking, microemulsions are in essence composed of bulk phases of water and oil separated by an interfacial region rich in surfactant, whereas for micellar systems, this is not clearly distinguished and most probably does not exist. However, with the lack of proper instrumentation, it is difficult to use different terms and, therefore, most scientists use the terms attheir convenience. II. INTERFACIALPHENOMENA

The most fundamental thermodynamic property of any interface is known as the interfacial tension [31]. The imbalanced perpendicular forces at the liquid-air and liquid-liquid interfaces were considered as the main reasons for two immiscible liquids to separate. The surface cannot be in mechanical equilibrium if there is a net force normal to the interface. The net forces at the interface were called interfacial tensions between liquids or liquid-air [30,57-591. However, many investigators concluded that it is not simple to interpret interfacial tension purely in terms of some net forces on molecules perpendicular to the surfaces, and therefore detailedstatistical mechanical models have been considered to try to describe the interfacial region [60,61]. Other approaches consider the density and the pressure profiles of the interfaces in order toexplain the complexity of the forces at the interface. Such extensive studies resulted in the discovery of various other interfacial forces, which play a significant role in explaining

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the factors affecting the disruption of liquid interfaces and other possible recombinations. The basic macroscopic model of an interface and its properties still show two bulk phaseshaving uniformthermodynamic properties separated by an interfacial region of some thickness and over which the thermodynamic variables may possibly vary. Experimental techniques and theories (distribution functions [62,63], partition functions [59-631, summation of interaction forces in the whole interfacial region [58]) have been developed to quantify the interfacial forces (interfacial tensions). The existing methods fall roughly into two categories [64-661: those in which the properties of the meniscus is measured atequilibrium (e.g., pendant drop shape[58], sessile drop shape [67], and Wilhelmy plate methods [68]) and those measuring properties at nonequilibrium (e.g., the drop volume [69] and du Nouy ring methods [70], which are faster than any of the other methods). One has to bear in mind that often it is difficult to interpret results with regard to rupture (formation) recombination of liquids (coalescence) in terms of interfacial tensions alone and more complex rheological interfacial measurements may be required. Interfacial shear viscosity measurements (the resistance to shear stress of monolayered moleculesin the plane of the surface) and dilatational (compressional) measurements [58] (the interfacial potential [71,72] which results from spreading a changed monolayer) must be measured and considered as a major contributing factor to the formation and/or coalescence of immiscible liquids. The art and science of bringing two mutually insoluble (or only slightly soluble) liquid phases together in the form of droplets (Figs. 8 and 9) is known as emulsion technology[1,31-331. An emulsion is thermodynamically unstable and the tendencyof the two liquids to disrupt and separate is a natural and favorable process. The success of emulsion technology lies in “keeping” the system in a metastable state by opposing the mutual interfacial flow of liquids of one droplet to the otheronce thedroplets approach each other asa result of Brownian motionin thesystem. In practice, the emulsion technologists deal with. two separateprocesses: the formation of the emulsion and thedestruction (rupture) of the droplets and the consequent separation of the droplets into two immiscible liquids. Many technologists will call it the constant battle to keep the emulsion stable. 111.

EMULSION FORMATION

The formation of an emulsion is a “fast” process and takesplace in milliseconds. It is achieved by applying mechanical energy [l].First, the interface between the two phasesis deformed, forminga liquid film between the two liquids, which later deforms to such an extent that droplets form. These

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Oil

Water

Fig. 8. Schematicpresentation of normalemulsiondropletsstabilizedby the adsorption of an emulsifier at its surface in comparison with an oil-in-water microemulsion (emphasizing the size, curvature, and packing differences). droplets are mostly far too large and they are subsequently broken-up or disrupted into smaller ones. Hence, the disruption of droplets is a critical step in the emulsification (Fig. 10). The deformation of the droplets is opposed by the Laplace pressure. The pressure at the concave side of a curved interface with interfacial tension, y, is higher than thatat theconvex side by an amount of AP = (2y/r) (where r is the radius of a spherical droplet). Any deformation leads to an increase in AP. The interface be-

Emulsions and Microemulsions 42 1

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tween the liquid deformation requires additionalenergy known as the viscous forces (velocity gradients) excreted by surrounding liquids, and a viscous stress has to be overcome (same magnitude as Laplace pressure). The pressure gradients(very smallportion of the totalenergy required, (35 kJ m-3) or the velocity gradients needed (approximately 3 mJ * mb3, which dissipates into heat) are mostly supplied by agitation and other external shearforces. However, certain surfactant molecules can be of significant help in reducing these gradient pressures. The molecules that are capable of adsorbing onto the droplets' surfaces and alter their properties and consequently reduce the interfacial tensions and hence stabilize the emulsion are the emulsifiers (Fig. 11). The emulsifier lowers y (e.g., from 40 to 5 mN m-') and thereby the Laplace pressure. The emulsifiers also help with the formation of film of a continuous phase between the droplets (which is a prerequisite for separation of the droplets). The adsorption and surface layer formation is mostly spread by the intense agitationof energy, and .it depends greatly on the natureand concentration of the surfactant. Adding a suitable surfactantmay reduce considerably the agitation energy (factor

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of 10) needed to obtain a certain droplet, since the surfactant reduces the gradient viscosity pressure. One must realize that much of the surfactant is depleted from the interface during the emulsification process owing to the large increase in the interfacial surface area. The formation of emulsion immediately causes the droplets to seek ways to recombine and coalesce again. Luckily the coagulation process is comparatively “slow” and inpractical systems it takes minutes or months, with the net process resulting in the formation of metastable dispersed liquid droplets in the continuous phase termed emulsion droplets. An alternative way of making metastable emulsion with a large number of droplets is to start from a very tiny nucleus and then allow them to grow to the required size-the condensation method, which is relatively easy with solid particles dispersed in liquid, but is difficult for liquid-inliquid dispersions. The surfactant’s role in the emulsion formation is critical, since it largely determines which phase is going to be the continuous one: It was experimentally established by Bancroft’s rule that the continuous phase will be the one in which the surfactant is soluble. The explanation is presumably that droplets of the phase in which the surfactant is soluble are very unstable to coalesce or that a film devoid of surfactant cannot be made. IV. EMULSIFICATIONTECHNIQUES

Many devices have been designed to produce emulsions. The principles of some of those emulsifying machines are based on construction of vessels with strong flow (with baffles) on which the liquid can impinge (laminar and/or turbulent pipe flow); injection of one liquid into the other; rotorstator machines for vigorous stirring; colloid mills; ball and roller mills; high-pressure homogenizers; ultrasonic techniques; aerosol injection into liquids; and electrical and condensation devices [1,73]. It is beyond the scope of this chapter to elaborate on these various methods (Fig. 12). Emulsions in which water is the internal dispersed phase are termed water-in-oil emulsions (W/O), whereas emulsions in which oil is the dispersed phase and the water is the continuous phase are known as soil-inwater ( O N ) emulsions. More complex systems in which one emulsifier is further dispersed into anothercontinuous phase are called double or multiple emulsions (W/O/W or O/W/O emulsions). The way the substances are added is important. Usually the surfactant, which should be most soluble in the continuous phase, is indeed dissolved in that phase, butit is possible to dissolve it in the dispersed phase, which can result in the formation of smaller droplets. The in situ formation of an emulsifier is also possible (the so-called nascent-soap method). Slow addition of the dispersed phase into

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Fig. 12. Principle of some emulsifying machines. The slit width in the homogenizer valve is very much exaggerated. Arrows indicate the directions of flow and rotation.

the continuous one during emulsification is often advantageous: It may enable addition of surfactant to the dispersed phase. If both phases are mixed in bulk, the one containing the surfactant usually becomes the continuous phase. Temperature plays a significant role in the emulsion formation. Often, it is required to first dissolve the phases or the emulsifier (if solid). Frequently, the solubility of the emulsifier and its surface tension reaction strongly depend (ethoxylated surfactants) on the temperature, and the hydrophilic water-soluble emulsifier becomes much more lipophilic and more water insoluble as the temperatureis raised. Therefore, it is essential to consider the emulsification temperature prior to any emulsification action. One must also remember that the emulsification process produces energy, the dissipation of the viscosity (velocity) gradient energy, which may affect many variables. The emulsification process can be evaluated in terms of emulsion capacity (the maximum amount of dispersed phases that can be emulsified under specified conditions). The emulsifier stability (the amount of phase separation that takes place with time under specified conditions) and the droplets size distribution (which seems to be the most practical way to study emulsification). Much has been written on the critical “events” that take place at the different stages of emulsion formation. Excellent discussions can be found in the Walstra’s chapter on “Formation of Emulsions’’ in the Encyclopedia of Emulsion Technology [l].The conclusions that should be stressed are (1) any emulsification process goes first through a film formation between the liquids which must exist for a little while; and (2) the film tends to drain rapidly under the influence of gravity. Interfacial forces tend to straighten the interface. The surfactant stabilizes the film. It allows an interfacial tension to exist, causing a considerable resistance to

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drainage of the film. As a result, it implies that slow stirring at the beginning of the emulsification (lower drainage) and high surfactant concentration are recommended. At the second stage, the plane (film) interface is disrupted into droplets. The plane disruption is accelerated by turbulence (only when y is verylow), capillary ripples (surface waves whichcan cause shredding of the droplets), and othereffects. The damping of low-amplitude capillary waves has been thoroughly studied [74]. It turns out that both the viscosity of the liquids and the presence of a surfactant enhance damping. Butif surfactant concentration is locallydifferent, as may occur in practice, the amplitudeof the wave may actually increase at the stepof lower y. It has been reasoned thatthis may be a cause of droplet formationduring emulsification. In some cases, an interface is unstable, that is, local perturbation tends to increase, and a Rayleigh-Taylor instability [75,76] occurs mostly when the interface is accelerated perpendicularly and directed from the lighter to the heavier phase (Fig. 13). Kelvin-Helmholtz instability [77] takes place inmany emulsions and arises when two phases move with different velocities parallel to the interfaces.

2

Fig. 13. Rayleigh-Taylor instability of an interface; the denser phase is below; the interface is accelerated in the downward direction. At the left side, pure Rayleigh-Taylor instabilityis depicted; at the right side, Kelvin-Helmholtz instability develops at the flanks of the waves. (From refs. 75 and 76.)

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EMULSION STABILITY

Emulsions consisting of microscopic droplets of 0.1-50 pm and dispersed in a continuousphase are thermodynamically unstable and tendto coalesce so that thereis a net reduction in the interfacial area. Hence, coalescence is a thermodynamic spontaneous process. Nevertheless, there are numerous examples of spontaneous, naturally occurring emulsions, which are stable. Milk is a good example. Stable emulsions exist in some areas of application; for example, in food,; (mayonnaise), pharmaceutics (intravenous fat emulsions), cosmetics (some skin creams), and agriculture (some pesticide emulsions). Much has been done in the last decades to understand the factors affectingsuch unusual stabilities. In order to better understand the concepts of stabilityhtability, it is essential to study the stepsand mechanisms that lead to phase separation of the liquids. Three to four main “events” can take place either in parallel orin sequence in each emulsion: creaming, flocculation, coalescence, and in some cases also Ostwald ripening (Fig. 14). All the processes lead eventually to an “emulsion breaking” and phase separation. It is essential to distinguish between the processes and to learn to control them to some extent. Creaming is the rise of dispersed droplets under the action of gravity, with the droplets remaining separated when they touch. Creaming takes place in any dispersion if the phases are not exactly equal in density and if the dispersion medium is truly fluid. It is well known that forsmall spherical undistorted dropletscreaming will depend strongly on the radius of the droplets, the difference in the density of the two phases, and the viscosity of the continuous phase. The rate of creaming is governed by Stoke’s equation:

is the creaming rate, pc-pdis the density difference between the two phases, 7 is the viscosity of the continuous phase,and r is the radiusof the droplets. Stoke’s law assumes that thefollowing conditions are valid [77]: (1) a spherical shape, (2) no droplet deformation, (3) a single particle (dilute conditions), (4) Newtonian fluid of the continuous phase with continuous properties, and (5) no diffusion. Emulsion droplets under rest are spherical, but they can be deformed in the shear field caused by the creaming. With large droplets or droplets stabilized by solid interfacial particles, distortion of the dropletscan occur owing to changes in pressure onthe different sides of the droplets (topvs bottom) which willlead to an increase in the surface area and resultin variation from Stoke’s equation. The deformation effect

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Flocculation

Creaming

Fig. 14. Schematic representation of the instability processes occurring in emulsions. takes place (as can be easily calculated) only for very large droplets (>l00 pm) andtherefore it is not a severe drawback. However, Stoke's law has an additional limitation, since it is good only for noninteracting droplets, which means it is good only for diluted systems. For more concentratedsystems, the term mass rate of creaming has been adopted (i.e., the motion of the center o f the mass of the dispersed phase) (Greenwald) [ 7 8 ] . Additional complexation is related to the fact that the emulsion droplets vary in size (polydispersity), which also affects the creaming rate, and finally parallel phenomena such as flocculation and coalescence should not be neglected when creaming rates are measured or evaluated. Creaming can be hastened by applying (for charged emulsions) an external electrostatic field across the end of the vessel, centrifugation, or dilution. Creaming rates can be slowed down by decreasing the emulsion droplet sizes, equalizing the densities of the two phases; and increasing the

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viscosity of the continuous phase (by adding viscosity builders with high specific gravitysuch as polysaccharides). Flocculation is the act of stickingtogether of droplets and the formation of three-dimensional clusters. In otherwards, flocculation is a process in which an aggregation of droplets takes place as a result of collisions in combination with adhesive interdroplets forces. It is clear that since in both creaming and flocculation, there is no flow of liquid from one dispersed droplet to another, these two phenomena are reversible. Flocculation is a complex phenomenon caused by Brownian movement, gravity, and shear motions of the droplets and isverydifficult for quantitative evaluation. The free energy change of flocculation, for weakly interacting droplets AG,,, can be positive or negative depending on the droplets’ concentration, and thus one can assume a critical flocculation concentration belowwhich the emulsionis thermodynamically stable with respect to flocculation but above which reversible flocculation to an equilibrium extent occurs. For strongly interacting droplets, this critical droplet concentration is so low as to be practically insignificant, and the flocculation may be classified as being irreversible. The flocculation process is then sometimes referred to as coagulation. In thermodynamic flocculated unstable systems, flocculation may be effectively prevented if a large enough free energybarrier exists between the flocculated droplets. The free energy barrier is derived from the interfacial forces such as long-range Van der Waals forces and electrostatic forces. The net force of the two sets of interactions will determine the behavior of the emulsion droplets to flocculate and further coalesce or to stay stable in the kinetic sense (Fig. 15). A quantitative treatment of the three different fundamental mechanisms which mayinduce flocculation (Brownian, shear-induced, and, gravity induced) was recently done by Bergenstahl [77]. The Brownian flocculation was first introduced by Smoluchowski [79]and described as the reaction (interaction) between twoparticles colliding and producinga new aggregate. The flocculation rates depend onthe number of particles present in the dispersion (concentration) and on the interaction time. Bergenstahl [77] mentioned twoadditional factors that affect the Brownian flocculation: the surface forces (surface potential or surface charges) and the hydrodynamic interactions. Fig. 16illustrates the two effects on theemulsion stability to flocculate. The two factorsare verysignificant whenBrownian flocculation takes place and must be strongly addressed if such flocculation has to beminimized. Bergenstahl [77] also found that the shear-induced flocculation (also known as the orthorhombic flocculation) as described by Smoluchowski [81] has similar limitations to those of the Brownian flocculation; that is, surface forces were not included, hydrodynamic interactions were notcon-

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h/n m

Fig. 15. Calculated examples of repulsive(GR), attractive(GA), and total interacas a function of surface separation distance, h, of two identical tion free energy(GT) (DLVO); (b) steric spheres: (a) electrostatic repulsion and van der Waals attraction repulsion and van der Waals attractions.

10

2

Fig. 16. The stability to flocculation factor including viscous interactions versus the surface potential and the salinity (surface charge) of the emulsion [77,80].

Emulsions and Microemulsions

43 1

sidered, and deflocculation was neglected. Gravity-induced flocculation, which is due to settling of particles, was first estimated by Saffman and -mer [82] assuming that every gravity-induced collision leads to attachment of the particles. Bergenstahl added to the basic Stokes’ law for settling of particles, the hydrodynamic interactions, and the surface interactions (as described by Melik and Fogler [83,84] and was able to predict stability areas forgravity-induced sedimentation as, a function of the droplet size, surface potential, and density difference between the particles. The mathematical expressions are very complicated but allow better evaluation of the possible mechanisms leading to flocculation, better predictions of the factors affecting the emulsions, and good evaluation of the stability to fluctuation that any emulsion can have. Coalescence is joining of small droplets in an emulsion to form large droplets or the process inwhichtwo droplets form one larger droplet leading ultimately to the formation of two separate liquid layers (Fig. 17).

Flocculation

l

0 Fig. 17. The primary destabilization of an emulsion is the flocculation that leaves aggregates of droplets(A). Subsequent coalescence leaves an emulsion with a wide distribution of droplet size (B).

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Table 1. Factors InfluencingTime for Coalescence of Emulsionsa Parameter

Effects

Droplet size

Exponential increase increasing with radius Exponential decreaseincreaswith 23 ing radius Interfacial tension 25increasing withIncrease interfacial tension Decreaseincreasing with interfacial tension.

References 23,24

26

‘Measured as time for separation rate of droplets’ size increase or as droplets’ lifetime

The coalescence process, examined from the point of view ofthe individual droplets in an emulsion, contains four basic steps: (1)The approach of two droplets toward each other achieve to an adhesive contact. (2) the drainage of the film between droplets in contact. The rate of the drainage process determines how rapid the critical thickness of the film is reached. (3) The rupture of the film. This is a stochastic process, the probability of which is determined by the thickness of the film. (4) The merging of the droplets. If the viscosity is low the process is rapid. However, emulsions with high viscosity will merge very slowly and may even be disrupted during the process. Table 1 summarizes the factors influencing time (rate) of coalescence. It can be seen that the droplets size and the interfacial tensions are key factors in the coalescence times. The coalescence process is very difficult to explain in simple mechanical events and the literature is still quite confusing. The main treatments arebased on thekinetics of the individual droplets, as was first suggested byVan den Tempe1 [S51 and later was elaborated by others.The main steps include creaming, consolidation (flocculation) (Fig. 18), film drainage, and film rupture. Bergenstahl [77], in his recent close examination and critical review of the coalescence process, examined a set of principal effects that can influence the coalescence process (Table 2). The large number of parameters affecting the coalescence emphasize the complexity of the process that takes place in the course of the collision of two droplets. VI. THE SURFACTANT AND THE HYDROPHILE-LIPOPHILE CONCEPT

Surfactants play a significant role in any of the two destabilization phenomena (flocculation and coalescence) by adsorbing onto the oil-water interface and contributing to the repulsive forces and the London dispersion

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Table 2. Principal Effects of Different Variableson the Various Process Steps Involved inthe Phase Separation Consolidation Drainage Rupture Creaming Variable

viscosity Droplet viscosityphase Continuous size Droplet Volume fractionof disphase persed difference Density tension Surface viscosity Surface elasticity Surface Depth of attractive minima interaction Repulsive thicknesslayerAdsorbed Solubilityof emulsifier in phasedispersed the HLB (hydrophilicflipophilic of balance emulsifier) the

0

+

0 0

0

+

-

+

+ +

+ +

+

-

-

0 0 0 0 0 0

0 0 0

-

-

+

+ + + 0 + +

0

0

-

0

0

-

+

+

+

+ 0

+ 0

0

0 0

(+) denotes that the variable has a positive effect; (-) denotes a negative effect; (0) the variable hasno effect on the phenomenon (process).

forces. Mixed surfactants are particularly more efficient than single surfactants with respect to the coalescence rate. The enhanced stability is attributed to the formation of intermolecular complexes at the oil-water interface, forming a densely packed layer and reducing the interfacial tension to very low values of about 0.1 mN * m-'. In addition, the interfacial complexes are capable of increasing the interfacial viscosity (although each component giveslowviscosity). Some investigators [86] claim that the synergism is derived from the enhanced adsorption kinetics of the surfactant when such two surfactant mixtures are used. The coherent, strong interfacial film should act as a barrier preventing coalescence by virtue of its rheological properties; for example, its high dilutional viscoelectricity. As a result, most technologists prefer using mixtures of two surfactants in most emulsion applications. The selection of surfactants is still quite difficult and requires much experience and experimental work. Griffin [87,88] has introduced the empirical concept of hydrophile-lipophile balance (HLB) for selecting a surfactant or a blend of surfactants. Surfactants with low HLB will form W/O emulsions and those with high HLB will form O/W emulsions (Table 3). Several methods have been proposed to determine the HLB of some nonionic surfactants (the HLB values are meaningful only for nonionic

W/O

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Table 3. Summary of HLB Ranges and Their Application

HLB range

Application

3-6 7-9

8-18 13-15 15-18

agent Wetting OIW emulsifier Detergent Solubilizer

surfactants and practically meaningless for any charged surfactants) and tables with the HLBvalues are available in any textbook. Moore andBell [89] suggestedthe H L numbers (hydrophile-lipophile ratio) as an improvedcriteria for selection of surfactants. Greenwald [90] suggested a water titration procedure for surfactant in organic solution, but this method is not widely used. Davies [91,92] has developed a method by which he calculated the HLB values of surfactants directly from their chemical formula, using empirically determined group numbers: HLB = 7

+ 2 hydrophilic group number - 2 lipophilic group number

The values of the group numbersare given ina simple HLB table(Table 4). The match betweenthe Davies method for evaluating HLB and the empirical work is quite good. The experimental determination of HLB of a surfactant is also not an easy task. Greenwald [90] developed a titration procedure andRacz and Orban [93] used a calorimetric method (based on the heat of hydration of ethoxylated surfactants) (Fig. 19). Becher and Birkmeier [94] suggested an HLB determination by gas-liquid-chromatography (GLC). In any event, there was a need to find the relationship between the surfactant’s HLB and the coalescence (stability) of emulsion. As a result, different oils were assigned to different required HLBs in order to obtain maximum stability (Table 5 ) [88,95]. The formulator is asked to match the required HLB of a given “oil”or mixture of organic solvents to the HLB of the surfactant (or mixtureof surfactants) by varying the natureof the surfactants and their relative proportions in the blend. Davies [91,92] has also demonstrated the effect of the partition of the surfactant between the oil and water phases in the emulsion coalescence and related it in an empirical equation to the HLB surfactant. He showed that the HLB group numberis proportional to theenergy barrier to coalescence set up by the water which is firmly bound to the hydroxyl or ester group on thesurface-active agent molecule. He also explained the correlation between HLB and thecloud point of nonionic surfactants.

p

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Table 4. HLB GroupNumbers Groups Hydrophilic -S04Na+

-COO-H+ -COO-Na+ N (tertiary amine) Ester (sorbitan ring) Ester (free) -COOH -0-

CH (sorbitan ring) Lipophilic -CH"CH, -CH-,-CHDerived -CHZ-CH2-0-CHZ-CHz-CHZ-O-

38.7 21.2 19.1 9.4 6.8 2.4 2.1 1.3 0.5 0.475

0.33 -0.15

Source: From ref. 91.

One must realize that the HLB concept is based on a simplified picture of the coalescence and the formationprocess and does not take into account the important role of liquid-film thinning between droplets and the importance of shear and dilational properties of the adsorbed surfactant layer around the droplets. Therefore, it is not surprising that in practice there aremany examples of emulsifications that do not obey the HLBrules and concepts. Many investigators have cast doubt on the validity of the concept [73,75,76,96,97]. In spite of the criticism, the HLB index can be very usefuland practical consideration in the process of making emulsions. In practice, in formulating a given emulsion, one may take any pairof emulsifying agents, which fallat opposite ends of the HLBscale, for example, ' b e e n 80 (sorbitan monooleate, with 20 ethylene oxide units, HLB=5), and use themin various proportions to cover a wide range of HLBs. Emulsions are made in a given way for all combinations with fewpercentages of the emulsifying blend. The stability of the emulsions is then evaluated at each HLB number, and from the rate of droplet coalescence, the most effective HLB value, that is, the HLBvalue providing optimum stability, can be found. Various other surfactant pairs are compared at these HLBnumbers until the most effective pair is found [2].

18 -

/ /

16 1L

Wlth ethylene oxlde e+

, 1

/.

,

-

1210 m

-

86-

oxide

1-

Water number

Fig. 19. Correlation of HLB in the water number. (From ref. 90.) Table 5. HLB Values for Various Oil Phases Application Cream, all-purpose Cream, antiperspirant Cream, cold Cream, stearic acid Creams and lotions Lotions Oil, perfume Oil, mineral Oil, vegetable Oil, vitamin Ointment bases absorption washable Ointment, emollient Polishes

type

Emulsion

HLB range

OIW OIW OIW OIW

6-8 14-17 7-15 6-15 4-6 6-18 9-16 9-12 7-12 5-10

W10 OIW OIW OIW OIW OIW

W10 OIW OIW OIW

2-4 10-12 8-14 8-12

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VII. PHASEINVERSION

In many cases, when an emulsion is made, a sudden event takes place which implies the inversion of the internal phase of the emulsion into the continuous external phase; for example, O/W emulsion inverts into W/O or vice versa. This phenomenonis related to the HLBvalue of the surfactant used, or theinternal phase volume. It has been shown that ata given emulsifierconcentration, the viscosity of the emulsion increases as theinternal phase volume(G) increases. At there is a sharp decrease in the viscosity of the emula certain critical GC, sion, which corresponds to the inversion of the emulsion. The @c depends on the emulsifier concentration, and inversion is expected to take place at higher GCas theemulsifier concentration increases [75,98]. In theearly days of emulsion technology, the inversion was attributed toa packing factorof the droplets. Thus, it was assumed that at a phase volume of db0.74, of the highemulsion droplets cannotaccommodatetogetherbecause density packing, and any attempt to increase the phase volume will cause an inversion or breaking of the emulsion. New theories and recent examples show that an inversion can also take place at low volume phases [99,100] and doesnot occur atvery highlypacked systems (99% by volume) [99]. It seems thatthe inversion isrelated more to the natureof the emulsifier than to theinternal phase volume, andthe emulsion tends to invert if the emulsifier used does not have the proper hydrophilic-lypophilic balances and tendsto migrate from the interface to thecontinuous phaseat a given concentration or temperature. Shinoda [101-1041 has shown that W/O emulsions prepared with nonionic emulsifiers tend to invert if the temperature is raised (since nonionic surfactants become less water soluble in water asthe temperatureincreases and leave the interface). In a series of papers Shinoda and coworkers [96,97,100-1101 have introduced a new concept for emulsifierperformance-the HLB-PIT relationship-and have shown that the phaseinversion temperature (PIT) of nonionic emulsifiers is influenced the by surfactant HLB number (Figs. 20 and 21) [96,97].The following conclusionswere drawn:(1) the size of emulsion droplets depends on the temperature and the HLB of emulsifiers; (2) the droplets are less stable toward coalescence close to the PIT; (3) relatively stable O/W emulsions are obtained when the PITof the system is some 2065°C higher than thestorage temperature; (4) a stable emulsionis obtained by rapid cooling after formation at thePIT; and( 5 ) the optimum stability of an emulsion is relativelyinsensitive to changes of HLB value or PITof the emulsifier, but instability is very sensitive to the PITof the system. Later, Shinoda et al. [l031 studied the effect of the molar mass and the molar mass distribution of the hydrophilic chain lengths of alkyl or

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439

-

14

-

L

QI

n

E,

z

m

2

120' 0'6

IO

3

d4 20

40 60 80 100 Phase Inversion Temperature ('C)

120

Fig. 20. Correlation between the HLB numbers of nonionic surfactants and the phase inversion temperatures (PIT,HLB temperature) incyclohexane-wateremulsions stabilized with the surfactants(3 wt% system). 1. Tween 40 9. i-R8C6H40(CH2CH20)8,5H 2. i - ~ C 6 H 4 0 ( C H 2 C H , ~ , 7 , 7 H 10. i-R,2C6H40(CHzCH~)g~7H 3. Tween60 11. i-R,,0(CH,CH20)6,,H 4. i-RgC6H40(CH,CH20)14H 12. i-R,,C,H,O(CH,CH~),,H 5. i-R,,C6H40(CH2CH, O)& 13. i-R9C6H40(CH2CH20)7,4H 6. RL?.0(CH2CH20)10.8H 14. R,20(CH2CH,0)4,2H 7. i-R8C6H40(CH~CH2O),,,H 15. i-R&H4O(CH2CH,0)6H 8. i-R9C6H40(CH,CH20),,,H 16. i-R9C6H4C(CH,CH20)6.2H

alkyl aryl polyoxyethylene ethers and emulsifiers (having the same PIT) on the stability of O/W and W/O emulsions. They found that stability against coalescence increases markedly as the molar mass of the lipophilic and hydrophilic groups increases. Moreover, the emulsions showed maximum stability when the distribution of the hydrophilic groups was fairly broad. It was also found that in those cases where the distribution of the hydrophilic chains is broad, the cloud point is lower and the PIT is higher than in the corresponding case for narrow-size distributions. Thus, the PIT and HLB numbers are directly related parameters provided the distribution of the hydrophilic chains of the emulsifiers is similar. The HLB number of an

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440 16

14

m

1

12

IO

20

100 Phase Inversion Temperature (‘C)

60

140

Fig. 21. Correlation between HLB numbers andPITS in various oil-water (1:l) emulsions stabilizedwith nonionic surfactants(1.5% system). 1. i-%C6H40(CH2CH,03,,,H 4 i-R9C6H,O(CHZCH,O),,,H 2. i-&C6H,0(CH,CH,0),,H 5. i-%C6H,0(CH,CH,0)6,2H 3. i-R9C6H4O(CH,CH,O),,H 6 . i-R9C6H,0(CH,CH,0),,3H

emulsifier of broad distribution is lower than that of the pure emulsifier having the same PIT. A plot of HLB value and PIT forcyclohexane-water emulsions (1 : 1by volume), stabilized by a number of nonionic surfactants [96],is shown in Fig. 22. This figure is useful for the estimation of the change of the HLB value and the optimum hydrophilic chain length from the change of the PIT atvarious temperatures. VIM. STERICSTABILIZATION

Emulsions can be stabilized also by naturally occurring macromolecular substances such as gums and proteins. Such stabilization takes place quite often in food and pharmaceutical emulsions. In the past, stabilization by amphiphilic macromolecules was attributed to the formation of “films” atthe interface. The adsorption of

44 7

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Fig. 22. Relationshipbetween HLB and PIT forcyclohexane-wateremulsions stabilized with nonionic surfactant(5%).

macromolecules at theliquid-liquid interface is characteristic, for example, protein like casein, bovine serum albumin (BSA), and soy proteins, and strongly depends onthe molar mass distribution of the adsorbedmolecule. The molecules form an adsorbedlayer comprising loops and tails that are anchored in the continuous phase. Thus, when two oil droplets approach each other, through aBrownian encounter, a repulsive force is generated owing to the presence of the adsorbed layers of the amphiphilic polymer. Such stabilization is known as steric stabilization. The steric interaction is effective when droplets approach a distance of b<2d, resulting from the interference of the two adsorbed layers (Fig. 23a). Theories have been

Id

lb)

Fig. 23. Tho limiting(constantadsorption)modelsfor steric stabilization: (a) interpenetration of adsorbed layers without compression; (b) compression without interpenetration.

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Gatfi and Aserin

designed to explain the contribution of the adsorbed macromolecules to the stability of the emulsion. The two main contributions to the stability/ instability of macromolecular-stabilized emulsions are related to droplets approaching a distance where compression occurs (without interpenetration) (Fig. 23b). 1. Volume restriction: the volume restriction term G,

or G, is derived from the reduction in the entropy of polymeric chains (configurational entropy) arising from the reduction in the total volume available to each chain if an interaction takes place (Fig. 24). 2. Mixing term: the mixing term G, comes. from a build-up in the polymeric amphiphile segment concentration in the interaction zone between thedroplets which leads to anincrease in the local osmotic pressure and the free energy (Fig. 24). The G, is alwaysrepulsive in its nature and depends on the nature of the polymers. However, the G, can havea repulsive or attractive effect on the total interaction depending strongly on the nature of the solvent (the continuous phase must be defined as AG, term better than8 solvent for thesurfactant in order for the to be repulsive), nature of the polymer (copolymeris by far better), and themolecular mass of the polymer. The roleof G, and G, in determining the stability-flocculation behavior of dispersions and emulsionsis divided into three sections: (1) high molar

Fig. 24. Total interaction free energy between two flat plates having adsorbed tails or loops: for polymers with the following characteristics:A = lO-’OJ; (Y = 1.2; W =2 x gcm-’; MW = 6000; (8)= 52 nm;area per chain = 50 nm’.

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x (better

443

0.5

lhon Q-solvent)

(W

Fig. 25. (a) The interaction free energy-separation curves for two droplets stabilized by high molar mass polymer. (b) Theinteraction free energy-separation curves for two droplets stabilized bylow molar mass polymer. mass, terminally anchored polymer chains; (2) low molar mass, terminally anchored chains; (3) multipoint anchored chains (e.g;, adsorbed homopolymer having a loop and train-type configuration at the interface). It is, therefore, very difficult in practice to evaluate quantitatively those terms and to predict whether the adsorbed polymer will cause steric stabilization of flocculation (Figs. 24 and 25). In order for dispersion a or emulsion, stabilized by adsorbed polymer, to remain deflocculated, the following criteria must be fulfilled: a. There should be full coverage (by the adsorbed polymer) of the particle or droplet interface. If this is not the case,bridging flocculation may then arise; that is, polymer molecules may become simultaneously adsorbed on the interfaces of both droplets or particles. Alternatively, “bare patches” on

444

Garti and Aserin the two surfaces may come into direct contact, leading to coagulation (or even coalescence). b.The polymermust be firmly anchored attheinterface. Homopolymers are not usually the most effective structures in this respect.For emulsions, AB block copolymers are particularly suitable, where A is soluble in one liquid phase and B is soluble in the other. If the polymer is not strongly adsorbed at the interface, desorption may occur during a droplet collision. c. The polymer layer must be thick enough so that G,, is negligibly small. A fuller discussion of the relationship between stability and G,, is given later in this section. d. The stabilizing moiety (e.g., that part of an AI3 block copolymer which is soluble in the external phase) must be in a good solvent environment. If p 0 . 5 , then Gmhbecomes negative; that is, the steric interaction becomes a net attraction. It is shown schematically in Fig.25 that a significant value of G,, is attained when x>0.5; that is, just beyond conditions for thestabilizing polymer.

In this way, Napper [l111 was able to demonstrate aclose correlation between the critical flocculation temperature (CFT) of a dispersion and the corresponding e temperature of the stabilizing polymer, and also between the critical flocculation volume fraction (CFV) of a dispersion and the volume fractionof added nonsolvent required to give a 8 solvent mixture for the stabilizing polymer. With regard to the CFT,some dispersions flocculate on cooling and others onheating. Generally speaking (but not always), the former occurs when a nonaqueoussolvent is the external phase,whereas the latteroccurs when water is the external phase. This behavior mirrors the temperature phase diagrams for thepolymer plus solvent system in question. There has not been a great deal of work carried out to date on the application of these ideas of emulsion systems. Recently, we [l141 have demonstrated that excellent steric stabilization can be obtained in double emulsions in whichthe outeroil droplets are very large (20 pm) and with specially designed block copolymers (with siliconic backbone) that strongly anchor to the oil phase. Similar results have been obtained with BSA, a naturaloccurring amphiphilic protein.

IX. MECHANICALSTABILIZATION Emulsions can be stabilized also with solid particles which are not amphiphilic in their nature (inorganic compounds) provided that the solids

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Fig. 26. Schematic illustration of solid particles adsorbed at the oil-water interface to provide mechanical stabilization.

adsorb onto theinterface (Fig. 26). Such stabilization is knownas mechanical stabilization,’’ since the solid particles provide a mechanical barrier between the droplets to prevent them from colliding and coalescing. The mechanical stabilization has only limited applications and has not been widely studied. The most impressing attempt was done by Oza and Frank [112,113] to stabilized double emulsions in the presence of a mechanical barrier obtained fromstrong adsorption of colloidal microcrystalline cellulose (CMCC). The double emulsion isa reservoir for a drug thatis strongly entrapped in the inner water phase and does not leach “on the shelf” (storage conditions). The emulsion is stable, and the entrapped drug is released slowly through the mechanical barrier. X.

EMULSIONCHARACTERIZATION

Emulsion are mostly characterized by the size distribution of the droplets and other physical properties such as dielectric properties, optical properties, thermal behavior, rheological properties, and other microscopic and macroscopic observations. The size distribution of the droplets is a very important parameter when characterizing any emulsion. Creaming, flocculation, and coalescence can be evaluated and influenced by the droplet size distribution. Therefore, it is essential to seek adequate methods to determine quantitatively the droplets’ distribution of an emulsion as a function of temperature, time, pH, and other factors that are responsible for the changes in droplet distribution. Practically speaking, only a limited number of drop-

lets are ever examined, but the resulting measurements are treated as if they constitute a continuous distribution of sues. Usually, when dealing with multivalued variables, statistical methodswill be utilized to limit the number of variables. Emulsion droplet datacan be obtained in terms of the number of droplets of a specific diameter, D, by means of optical or electron microscopy [115-1161; as diameter versus volume, V, by electrical resistance counting [117], and by diameter, D, versus mass, M, by x-ray coupled with sedimentation [118]. As a result, terms such as number of droplets, diameter (length), area, volume, and mass can be estimated or evaluated. Table 6 demonstrates the parameters that can be calculated. The datacan be presented in of droplets in range, diameter table form (droplets diameter range, number interval, and percentage on a numberof boxes of the range, the less then maximum diameter, and so forth), in histograms and in cumulative plots (of number, surface area,and volume (Fig. 27).

Table 6. Mean Diameter Definitions for Emulsion Droplets ~

Symbol ession8 literature)in

mean

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446

~

_

_

_

Mathematical

Descriptive name found (alternate

Number-length mean (arithmetic Number-surface (surface mean, diameterof average Number-volume mean (volume mean, diameter of average volume) mean Length-surface (linear mean, length diameter mean) mean Length-volume (volume-diameter mean) Surface-volume mean (surface mean, Sauter) Volume-(or weight-) moment mean (volume-(or weight-) mean, weight average particle size, De Brouckere) Could be expressed in terms numbers (AN).

. D,

(W) 122

(?%F) D,

U3

D, D, D*, D,

or D,

of number percentages (AP) as well as in actual

Emulsions and Microemulsions

12

14

18

18

447

20

22

24 28 26

Fig. 27. Histogram of dropletwithinrange droplet diameter range (pm).

30

32

34

36

38

40

42

44

(% by number) as a function of

Out of all the techniques mentioned above, the most useful information is the droplets’ size distribution behavior with time (indicating the rates of coalescence and flocculation); with temperature, electrolytes, and the presence or absence of various emulsifiers (as single blends and/or in combination with macromolecules). Much can be learned on the emulsion stability from its rheological properties. Emulsions exhibit a non-Newtonian behavior which is emphasized by its variation in the viscosities as a function of its flow properties (shear rate orshear stress). A creep compliance as a function of small shear stress over a period of time gives important information relevant to emulsion stability. For more detailed information on the rheological properties of emulsion, the reader is referred tothe Encyclopedia of EmulsionTechnology, Vol. 1 [l]. A sharp decrease in viscositymeasurements (at high or medium shear rates) of highly concentrated emulsions is clearly associated with the increase in mean drop size. Several such examples exist both with W/O and O N emulsions [119-1211.

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XI. EMULSIONAPPLICATIONS A.GeneralConsiderations

Emulsions are widely used ina range of industrial and foodapplications as well as agricultural, medical, pharmaceutical, and cosmetics. This chapterwill obviouslyconcentrate only on themedical andpharmaceutical applications. The use of emulsions in pharmaceuticals was widely spread in the early 1960s and 1970s, but its importance has been diminishing in the past years, mostly because more attractive ways of intake and dosage forms have become available. The main reason for this is the increasing need for formulations that canprovide sustained, prolonged, slow, and controlled release of active substances. Furthermore, there has been a rising interest in many solid dosages of active matter such as, tablets, capsules, microspheres, microcapsules, nanoparticulated matter, and powders entrapped onpolymeric matrices. Emulsions, microemulsions, and double emulsionsalso have to compete with various types of vehicles suchas liposomes. They also suffer from a lack of thermodynamic stability and, therefore, their shelf lives are restricted and requirespecial storage conditions. Microemulsions, although thermodynamicallystable, suffer from low-entrapment capacity and require large amounts of emulsifiers, which is not necessarily permitted for drug application. Any change in the emulsion’s or microemulsion’s stability (temperature, storage, decomposition of components) will affect the drugrelease, which willimmediately affect the stability. However, since emulsions are very inexpensive and easy to prepare, they will always be attractive to formulators for certain applications. The progress which has occurred in recent years in the area of double emulsions (better stability, smaller droplet size, high capacity, and better controlled release of active matter) has been attracting renewed attention, and new formulations have beensuggested and tried. The dropletsize distribution of the emulsion is yet another significant problem. Most emulsions cannot be used for parenteral applications because of blockage (emboli) problems. Microemulsionsare small enough in size but are made of emulsifiers that do not seem to have health permits. Their applications will be considered according to their delivery methods: by injection (parenteral), by mouth (oral), and by the skin and eyes (topical). It is beyond the scope of this chapter to discuss all the possible applications mentioned in the literature orin patents, and itwould be difficult to list all the constraints related to the use of emulsifiers and oils (health and physical stability, as well as chemical or biological stability). An attempt is

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made to bring some interesting examples of novel systems for controlled release, and discussions are brought only if the example helps in understanding or solving basic and/or critical problems related to the emulsion technology. B. Parenteral Emulsions

Parenteral emulsions are used as vehicles for carrying lipid-soluble materials, to control therelease of drugs, and, if possible, also to targetit. Most of the emulsions mentioned for parenteral use are of O/W or W/O/W type. Intravenous emulsions have very strict requirements regarding droplet size. Large droplets (over 5 pm) cause blockage (emboli) in the body. Toxicity effects have been reported in fat emulsions when meansize rises above 1.5 pm (Table 7) [122].Physicochemical parameters such as the viscosity of the external phase, phase volume ratios, and the partition coefficient of the drug are key factors influencing the release of a drug [123,124]. Variables such as droplet size distribution, surface characteristics, and surface charge are relevant factors in the relationship between physicochemical properties and physiological responses. It has been demonstrated [125,126] that fine particles “clear” from the blood slower than coarser particle-size emulsions; charged droplets clear quicker than neutral droplets and emulsionsstabilized by low molecular emulsifiers are cleared than those stabilized by high molecular weight emulsifiers [127-1291. Emulsions stabilized using phosphatides (phosphatidyl choline) were cleared quickly, whereas those stabilized with thenonionicpoloxamer (Pluronic F108) were cleared slowly. A mixed emulsifier system of phosphatide and Pluronic had intermediate characteristics (Fig. 28) [129]. The addition of drugs to the oil phase had a modifying effect on the clearance. The results support the concept that the emulsifying agent is of major importance for the removal of the emulsion from the bloodstream. Geyer Table 7. Effect of Particle Size on the Toxicity of Intravenous Lipid Emulsions Administered Intravenouslyto Rats Mean

Maximum

(pm) size (pm) System size 0.8 Fine

Medium Coarse

0.35 0.79 1.65

- 112 (103-122) 5.0

% Greater than 1pm 0 22 84

LDS0 (95% confidence limits) (mJJb)

(104-124)114

820% fat emulsionof soybean oil stabilizedby egg lecithin.

60 (53.5-66.5)

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2 t 0

b 100

200 lime (min)

300

Fig. 28. Effect of emulsification on the elimination of lipid emulsion. A, Pluronic F108:0, egg lecithin; 0 ,mixture of Pluronic F108 and egg lecithin.

[l271 has studied the effect of the emulsifier molecular weight on its clearance and found that clearance of fat emulsionsstabilized by the poloxamers with high molecular weight surfactants were cleared more slowly than from the blood than those emulsified with lowmolecular weight poloxamers. In spite of all those limitations, the administration of essential nutrients via the parenteral route is common practice. Glucose, amino acids, electrolytes, and vitamins are between the water-soluble matters intravenously injected. Fats areimportant sources of energy andmany efforts have been invested in introducing fat-emulsion for parenteralintake. Some commercially available lipid emulsions are listed in Table8. The oils were almost exclusively of vegetable origin, although synthetic glycerides, including simulated human fats and acetoglycerides, have been studied. Soybean oil and safflower oil [l301 appear to give rise to a low incidence of toxic reactions when purifiedand are resistant to oxidative changes (rancidity). Natural and synthetic compounds have both been considered as possible emulsifying agents. The natural agents were various types of lecithin (phospholipids) and the synthetics were nonionic polyoxyethylene-polyoxypropylenederivatives. A wide range of nonionic materials have been investigated as potential emulsifying agents for intravenous fats: polyethylene glycol stearate,

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Table 8. Some Commercially Available Lipid Emulsions Trade phase name Oil

(%)

Emulsifier (%)

Intralipid Soybean 10 orEgg lecithin 20 1.2 (Kabi-Vitrum) Lipofundin S Soybean 10 or 20 Soybean lecithin (Braun) 0.75 or 1.2 Lipofundin Cottonseed 10 Soybean lecithin 0.75 (Braun) lecithin Egg Liposyn Safflower 10 1.2 (Abbott) 20) (and Travemulsion Soybean 10 or 20 Egg lecithin 1.2 (Travenol)

Other components (%) Glycerol 2.5 Xylitol 5.0 Sorbitol 5.0 Glycerol 2.5 Glycerol 2.5

diacetyltartarateester of monoglyceride (DATEM), partially esterified polyglycerol, polyoxyethylene monostearate (Myrj), polyethylene glycol (Carbowax), nonylphenyl ethers (Tergitol), and polyoxyethylene sorbitan monoesters (Tweens). However, all of these materials gave rise to toxic reactions of one form or another. Only one rangeof nonionics was found to be free from toxic effects: the poloxamers (polyoxyethylene-polyoxypropylene derivatives). These have been widelyused for fat emulsions either ontheir own or as coemulsifiers withlecithin [131,132]. Although Black and Popovich [l331have attemptedto give guidelines and recommendations. Burnham and others [l341 have reported that a stable fat emulsion (Intralipid) intravenous-based feeding mixture have been preparedto simplify nutritional support for patients with gastrointestinal disease. Mixing of Intralipid with other nutrients before administration allows a constant infusion through peripheral veins. The physical properties of fat particles in Intralipid formulations were studied before and aftermixing withthree different combinations of amino acids, dextrose, and electrolyte. The effect of storage on lipid particle size was also examined. The mean particle size of the dropletsincreased slightly over a 48-h storage period at 4"C, but no large droplets were observed. Measurement of the electrophoretic mobility of the droplets indicated that the mixture of a fat emulsionwith an aminoacid, dextrose, and electrolyte did not produce any permanent change in the stabilizingfilm of phospholipid. Total concentrations of divalent cations are greater than 2.5 mmol/L, such as calcium and magnesium, couldlead to aggregation of fat droplets and the separation of the oil as a cream layer. These systems had poor short-term stability, and consequently Burnham et al. [l341 recorn-

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mended that feeding mixtures containing amounts of Ca2' and M e above this limit should beused with care. In similar fashion, Hardy et al. [l351 have shown that mixtures of amino acid, electrolytes, trace elements, glucose, and 10% soybean oil lipid emulsions werestable when prepared aseptically in3-L plastic containers and stored at refrigerated temperature for 72 h. The pH-buffering property of the amino acid solution was believed to be an important contributor to the stability of the mixture [136]. The lipid emulsion (mixed with amino acids and glucose for parenteral administration of dogs) was eliminated andmetabolized to triglyceride, free fatty acids, andphospholipids in a fashion similar to that observed during separateadministration of the same nutrients. Clinical data supportthese findings [137]. The clinical use of fat emulsions in parenteral nutrition has been described in detail in various textbooks and reviews [130,138-1411, and a variety of products is available, including those containing amino acids together with fat emulsions [130,142,143]. The possibilities of administering complete intravenous nutrition with fat as one of the sources of energy has been demonstrated in many investigations. Emulsified vegetable oils were used as test systems to explore the characteristics of the reticuloendothelial system. The role of a,-macroglobulin in the phagocytic process (in rats) using a lipid emulsion has been investigated by Molnar et al. [144]. They found thata,-macroglobulin was an important component, but thatthis material was not related to the a,macroglobulin in humans. They pointed out that plasma contains at least 60 proteins, and this does not include enzymes, clotting factors, complementing components, and hormone carriers. Ashworth and others [l451 examined the uptake of small lipid droplets by hepatic and Kupffercells. The uptakeof small particles was dependent on thesize and potencyof the hepatic sinusoids. In addition, the size, composition, structure, and charge on the particles was considered to be relevant. A critical particle size of 100 nm was proposed. Tonaki et al. [l461 used a gelatin test emulsion to investigate plasma components that promote phagocytosis (opsonins). Particles coated with human serum albumin werealso employed. Different uptakes in liver and spleen regions as well as small uptake in the lung were reported. They indicated thatthere was some specific interaction between reticuloendothelial cells and particles which was dependent on the nature of the particle surface. Intravenous O/W emulsion systems are also employed as diagnostic agents in the form of injectable radiopaques. Kunz et al. [l471 described in detail the preparation,sterilization, and stability of 50% oil-in-water emul-

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sions of iodized oil, iophendylate injection, and ethiodized oil (Ethiodol). Three emulsions of each oil injected intraperitoneally into rats gave an excellent radiopaque outline of the peritoneal cavity. Emulsion technology isquite a mature science withregard to pharmaceutical preparations. Most studies were carried out onmicroencapsulation as a release vehicle. Only a limited number of studies are available on improvements or innovations related to emulsions. Some of the key issueswere fat emulsions for parenteral applications:

1. Studies related to the stability assesment and its relation to the nature of the phospholipids [148-1501. Highly negatively charged droplets were prepared by using purified phospholipids enriched with negatively charged phosphatides; use of amino acids, urea, or carboxylic or sulfonic acid to form stable emulsions even after mixing with blood plasma [151]. 2. Attempts to make bettercharacterized parenteral emulsions(by Coulter counter [152,153]) and to improve their productiontechniques (microfluidizer [1541). 3. Stability studies related to thermal treatments[155,156]. 4. Effect of various nutient ratios on the emulsion stability of total nutrient admixture [157]. 5. Elimination and metabolism of a fat emulsioncontaining mediumchain triglycerides [1581. 6. Transdermal new formulations for special applications have been also studied. For example, new nicotinic devices and their usein the treatment of withdrawal symptoms associated with smoking cessation [1591. C. Lipid Emulsions for Drug Delivery

Low water soluble drugs are difficult to administer and, therefore, are good candidates for formulation into O N emulsion. Fat emulsions are known and used in parenteral nutrition (artificialcylomicra have been developed in the form of fat emulsions). The two-phase emulsion system should havean advantage overa solubilized system(use of cosolvents such as polyols) in that the drugcontained therein will not be able to precipitate as solid (harmful) particles when diluted. Furthermore, the presenceof the bulk of the drug in nonaqueous environment may lead to an increased stability of the drug (e.g., reduced hydrolysis) as well as to a possible controlled-release system. The use of soybean oil emulsionsas carriers for lipid-soluble drugs has

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30.8 LO.0 Dose ( m g I kg I

52.0

Fig. 29. Administration of thiopentone in aqueous and emulsion forms.0 ,emulsion; 0,solution. (After ref. 164.)

been pioneeredby Jeppsson and Ljunbergin Sweden [160-1631. The drugs, dissolved in the oil phase, have included barbituric acids, cyclandelate, diazepam, andlocal anesthetics and the routes used have been intravenous, intra-arterial, subcutaneous, intramuscular, and intraperitoneal. The greatest interest hascentered around the use of fat emulsions asvehicles for the intravenous administration of drugs [1601. For thecase of the barbituric acids [MO], a prolongation of anesthesia was observed when the drug was administered in the oil phase of a soybean emulsion comparedwith a solution of the corresponding sodiumsalt (Fig. 29). The results were explained either by a slow release of the drug from the oil particles or thepossibility of a more specific delivery of the drugsto the central nervous system (CNS) when the drug is confined in the oil droplets, the latter being due to the type of close association of droplets and thevascular lining of the CNS. As evidence for the possible close association of lipid particles with vascular linings, Jeppsson and Ljunberg[l641 cited the work of Schoefl and French [165], who demonstrated that in the mammary glands of lactating mice, chylomicra and artificial fat particles (soybean oil emulsion) were concentrated against the luminal surface of the endotheliumof small blood vessels and appeared to adhere closely to it. In some cases, the fatparticles appeared to have sunk deeply into thecytoplasm, or it appeared that thin flaps had encircled them. In contrast, in the lung, the fat particles were freely suspended in the blood plasma. The role of an enzyme, believed to

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be lipoprotein lipase, in this adhesion process and the transport of fat through the vessel wall was discussed. Many more reports exist in the literature on drug delivery by the intravenous administration of fat emulsions containing drugs. Examples are the administration of lignocaine [164,166] (which protects the heart from electrically induced arrhythmias), diazepam [167-1701, hexobarbital [l701 and phenyl butazone[170]. In spite theextensive work invested in stabilizingthe drug-containing fat emulsions with various peimitted emulsifiers, it is not yet clear whether the emulsifiers and the fats decompose in the system and whether theyplay any role in the release and/or targeting of the drug. Some investigators claim that certain effects (such as sustained release) could well be related to metabolic changes induced by the presence of the emulsion rather than delayed drugrelease or drug targeting [170,171]. Narcotic antagonists are importantcandidates for slow-release action and, therefore, have been tested in various formulations to induce physical dependence in rats following subcutaneousdepot injection [172,173]. Valinomycin [l751 and other anticancer drugs have been introduced into intravenous emulsions [173-1751. Tests in animals showed that the drug contained in an emulsion had effects similar to those of an aqueous suspension, but that the emulsion formulation required a 20-fold lower dose to produce these effects [176]. Artificial intravenous emulsions havealso been tested as “on-target” delivery systems for drugs that arevery oil soluble. The drugis dissolvedin the oil and the emulsion is injected. The fat particles (chylomicrons) can deposit and close contact between the fatparticles, and the endothelium can allow passage of the drug from the oil phase tothe tissue without it necessarily passing into theplasma. The excess chylomicrons willbe cleared by the liver, adipose tissues, heart muscle, and lactating mammary glands. In this connection, it is of interest to review the work of Meyerson [177], who found a prolonged-release effect after dissolving progesterone in soybean oil and thenemulsifymg it to give an O/W emulsion forintravenous administration. Similarly, Mitushima et al. [l781 have employed a lipid emulsion as a novel carrier for corticosteroids. Their studies in rats have suggested that corticosteroids incorporated inlipid emulsions are taken up by the reticuloendothelial system and certain inflammatory cells to a greater extent than free corticosteroids, thus resulting in stronger antiinflammatory activity [178]. Thus, it was believed that corticosteroids in lipid emulsion could be of value in the treatmentof certain diseases such as rheumatoid arthritis, where phagocytic inflammatory cells play an important role in the pathogenesis.

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D. Vaccine Emulsions

Adjuvant methods of immunization havethe advantage of achieving both the primary and secondary immune responses with only one injection of antigen and of maintaining the antibody level over a long period of time. Adjuvants have been used recently in tumor immunotherapy [179]. An adjuvant effect is obtained by adsorbing the antigen onto a mineral gel (aluminum hydroxideor phosphate). This is widely usedas an immunizing agent for human use [179]. Water-in-oil emulsions, consisting of anaqueous antigen solution dispersed in a mineral oil phase with the aid of a lipophilic emulsifier (mannide-monooleate), have been prepared to act as efficient immunological adjuvants. These emulsified systems presumable enhance the antibody response (slowly releasing secondary stimulation of antibodies) and the stability of the vaccine on storage [180,181]. Various mechanisms have been suggested to explain the high antibody response which these adjuvants produce, and it has been assumed that the emulsion acts as an inert depot, whichslowly releases antigen over a long period of time [179,182-1841. Steinberg [l831 has suggested that in addition to the depoteffect, the oil attracts mononuclear cells about the antigen which can take part in antibody production andgive a high antibody level. Freund and McDermott [l851 were some of the first to demonstrate adjuvant activity of mineral oil emulsions. These simple water-in-oil emulsions, subsequently termed incomplete adjuvants, were later modified to give a greater antigenic response by incorporating dried, heat-killed Mycobacterium tuberculosis in the mineral oil phase. Such emulsions are termed Freund's complete adjuvants [186]. The complete adjuvantinitiates a cellular response andis too reactive for immunization in humans, although it is used in experimental work [186-1891. Problems have been encountered in the use of mineral oil emulsion adjuvants, since mineral oil isnot metabolized andwill persist at theinjection site [179,187,188]. This has limited the use of mineral oil emulsions in humans. Studies have, therefore, beencarried out in the search for alternative oil phases for adjuvant preparations (hexadecane, squalane, sesame oil, and peanut oil. Stewart-Tu11and others have found that low molecular weight hydrocarbons of chain length C,&,, were goodalternatives to mineral oil). Early studies with vegetable oils .were unsuccessful, as the oils were unstable and gave lessantibody response thanmineral oils [189]. However, further work produced a suitable adjuvant of peanut oil [187]. This adjuvant consists of a water-in-oil emulsion of aqueous antigen together

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with an equal volume of an oil phase. Arlacel A is the emulsifier and aluminum monostearateis the stabilizer. A more recent report by Reynolds [l901 describes the adjuvantactivity of a refined peanut oil base emulsified inaqueous vaccines with glycerol and lecithin. This formulation was found to be metabolizable and caused no granulomasor abscess formation. E. Emulsions as Adjuvants

Oil-in-water emulsions have beenused as adjuvant-type systems in cancer chemotherapy [191].Successful tumor regression has been achieved by intratumoral injections of viable Bacille Calmette-GuCrin (BCG)vaccine, and a potential antitumor preparation has been constructed by combining trehalose dimycolate purified from BCG with aqueous soluble extracts of Re mutants of Salmonella species (Re glycolipid). An essential requirement of activity has been the formulation of an O/W emulsion. A drug injected into the interstitial spaces in muscle tissue is transported away from the injection site by the circulating blood, but someof it also reaches the regional lymph nodes. The amount of drug so absorbed depends on the injection formulation and the site of the injection. It is known that lipids, fatty acids, and high molecular weight substances are absorbed mainly to the lymph system and lower molecular weight compounds are absorbed into the blood. Many important drugs are water soluble with relatively low molecular weights; therefore, a specific drugdelivery system in the form of an emulsion or other colloidal system is necessary to deliver such agents to thelymphatic system. Sezaki, Hashida and other workers [192-1981 have investigated the use of a water-in-oil emulsion to provide drug-delivery systems using a series of anticancer drugs in a series of formulations; water-in-oil emulsion, water-in-oil emulsion containing gelatin, and also simple oily and aqueous solutions and aqueous intravenous injections for comparison. Injections were given by intramuscular, intraperitoneal, and intragastic routes. Concentrations of both the drug and the oil were measured in the blood, regional lymph, thoracic lymph, and lymph in other parts of the body and also at the injection site following injection at various sites. It was suggested that absorption of a drug from its injection site could be absorbed into the bloodstream or into the regional lymph nodes. It passes to the thoracic duct and thento thebloodstream. It can travel as theisolated drug or in the emulsion formulation. Using mitocycin C and bleomycin, it was noted that the water-in-oil emulsion produceda much greater specific delivery of drug into the lymph system than did an oil-in-water emulsion [195-196]. Sezaki et al. [195],

L

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therefore, suggested that encapsulating a water-insoluble drug in oil was highly advantageous forthe transport of drug fromthe interstitial spaces to the lymphaticcapillaries. They also noted that theconcentration of oil was much higher in the lymph nodes thanin the thoracic lymph and that it rose to a maximum quickly and then decreased in the former, whereas it was relatively constant in the latter. This suggested that theoil was transported to the lymph nodes, where it accumulated and was gradually released downstream. An oil-in-water emulsion containing gelatin as an emulsifying agent was found to be intermediate in its lymphatic targeting ability between the water-in-oil emulsionand the oil-in-water emulsionprepared by using polysorbate 80 [195-1961. The gelatin-containing emulsion was investigated, andit was found that themitomycin C was partially bound to theoil drops [195]. In subsequent experiments, a water-in-oil emulsion, using a relatively high concentration of gelatin as part of the emulsifying system, was compared with the simple water-in-oil emulsion. The concentration of the gelatin used was20% of the aqueousphase, and the gelatin was thought to form solidified microspheresin the oil. It was noted thata high drug concentration surroundingthe injection site was prolonged with the emulsion formulations, especially the gelatin emulsion. A sustained release of drug into both lymph and blood systems was significantly improved with the two emulsions comparedwith the aqueoussolutions [194-1981. The time during which the drug concentration in the regional lymph node was higher than that in other lymph nodes (in the orderof several hours with both emulsion formulations as compared with halfan hourfor the aqueoussolution). Also, the time delay to reach a maximum or peak concentration of drug in both blood and lymph was greater with the emulsion formulations. Microscopical observations made of the muscle fibers and the regional lymph nodes showed that multiple water-in-oil-in-interstitial fluid emulsion was formed in the muscle tissue on injection. With the aqueous gelatin-in-oil emulsion, microspheres (1-2 pm in diameter) were observed to be dispersed in the oil droplets. The investigators [193,194] suggested that this system wasmore stable than the simple water-in-oil emulsion and that itprolonged the release of the drug forthis reason. This suggests that the release of drug is due primarily to the breakdownof the emulsion. Measurements of the total amountsof drug absorbed intothe lymph system revealed that the microsphere-in-oil emulsion was superior to the water-in-oil formulation in promoting specific lymphatic absorption. It was concluded [l941 that water-in-oil and microsphere-in-oil emulsions would be advantageous for use in cancer chemotherapy. In the light of these experiments, bleomycin in the form of a microsphere-in-oil emulsion was injected into the appendix of rabbits having a carcinoma on the organ after surfkal removal of the carcinoma [198]. A

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mixture of medium-chain triglycerides (MCTs) wasused as theoil phase of the vehicle, since this had a lower viscosity than the sesameoil previously used and, therefore, was easier to inject. The MCT-based emulsion improved the original delivery characteristics of the microsphere-in-oil emulsion with sesame oil as the oil base. It produced a definite improvement in the extentof metastasis. The metastatic cells were seento bedestroyed after 1month (observedmicroscopically) compared with an aqueousformulation which slightlydamaged diseased cells, but these recovered within a month. It also delivered little drug IO the lung or liver, where it is especially toxic. Takahaski et al. [l991 investigated the use of water-in-oil emulsions as a vehicle for the injection of anticancer agents (bleomycin and mitomycin C) directly into tumorsfor antitumor activity and also for the preventionof lymphatic metastasis, thereby making use of the sustained-release effect of such a formulation. They found in experimental animals a prolonged level of drug in the tumor using a water-in-oil emulsion, and also a decrease in tumor size of 50% in 16 days and 75% survival rate compared with a 5% decrease in tumor size and a 40% survival rate with an aqueous injection (Fig. 30). Clinical trials on carcinomas produced encouraging results. In some cases, a multiple emulsion was employed as the delivery system.

-1 00

2

4

6

8

1014 12

16

Time after injection (davrt

Fig. 30. Changes in mean tumor diameterof rat carcinoma aftera single 0.5-mg dose of bleomycin in various forms. 0 , untreated control; A, aqueous solution intravenous; V, aqueous solution intratumoral; 0, emulsion intratumoral. (After ref. 199.)

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The chemical composition of a digestible oil may also be critical, as this will determine the physiological mechanisms stimulated during digesof tion. Forexample,thehydrocarbon chain length andthedegree unsaturation of the fatty acids liberated by triglyceride hydrolysis determine the extent of changes in gastrointestinal motility, with the longerchain unsaturated fatty acids havingthe greaterinhibitory effect [200,201]. Incorporation of long-chain fatty acids into the bile salt micelle increases the solubilizing capacity of the micelle for lipophilic materials within the intestinal lumen [202,203], and incorporation of unsaturated long-chain fatty acids may result in an increase in the permeability of the mucosal membrane [204,205]. Short- and medium-chain fatty acids are absorbed via the portal route, whereas long-chain fatty acids (>C,,) are absorbed into the lymph, with the unsaturated acids stimulating chylomicron synthesis. Lipophilic compounds selectively absorbed into the lymph are carried in the chylomicron [206], the natural fat particles, and therefore administration of long-chain unsaturated fatty acids may stimulate lymphatic absorption. It is apparent that the chemical composition of the oil phase of the emulsion is a major factor in determining the extent of drug absorption following oral administration. F. Oral Emulsions

Oral administration of drugs in an emulsion formhas been known for over 100 years and is easier to use than intravenous injection. Its main advantage is the morefacile absorption of certain drugs from the gastrointestinal tract. The ONV emulsions arerelatively easy to prepare, since there are no strict requirements on droplet size, nature of surfactant and its purity, nature of its fat, norphysical, chemical and biological properties. The range of emulsifying agents is greater than that available for intravenous emulsions. Materials suchas acacia, tragacanth gums, and methylcellulose as well as various nonionic surface-active agents have been employed in pharmaceutical formulations for oraluse [207,208]. The ionic surfactants are notnormally usedfor internal preparations but arereserved for topical and cosmetic applications. The G U S (generally regarded as safe) listings of the U.S. Food and Drug Administration are a useful source of information [209]. Vegetable oils suchas cornoil, peanut oil, and soybeanoil are used as emulsion vehicles for drugdelivery. Mineral oils (liquid paraffin) are emulsified to provide a laxative effect but are not used for drug administration unless the drugis intended forthe same pharmacologicaleffect. Many attempts have been madeto entrapvarious drugs in O N emulsions for oral adsorption.

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Higher levels of drug in the plasma during the first 4 h after oral administration of acetyl sulfisoxazole O/W emulsion were observed in Svenson's early studies [210]. The emulsion was also shown to be three to five times more active against streptococcal and pneumococcal infections in mice. Similar results were reported with sulfonamides [211]. In most formulations, greater advantages were found in drugs with limited water solubility (poor dissolution), such as indoxole [212], griseofulrin [213,214] (Fig. 31), phenytoin [215], and theophylline [216]. Formulation of nitroglycerin in a sesame oil-in-water emulsion yielded lower and later peak plasma levels than the equivalent aqueous solution, butbioavailability was unaffected [217]. The delay in drug absorption from the emulsion was attributed to the time required forthe drug to move from the oil phase into the aqueousphase prior to absorption andto effects of the oil on gastric emptying. Thus, the emulsion provided some sustained-release characteristics to nitroglycerin absorption, whereas not affecting the extent of absorption.

3 r

'1

Time (hrl

Fig. 31. Administration of griseofulvin in different dosage forms (30 mglkg of micronized griseofulvin inrats). 0, aqueous suspension;A,corn oil suspension; 0 , corn oil-in-water emulsion containing suspended griseofulvin.(After ref.213.)

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Absorption of the fat-soluble vitamin, vitamin A, occurs primarily via the lymphatic pathway [215,218]. Emulsification of vitamin A palmitate and vitamin A alcohol in corn oil has been found to increase the total absorption of vitamin A in normal patients and in those with cysticfibrosis [219,220]. These observations have important implications on both the dietary management of patients with cystic fibrosisand, moregenerally, on the use of the lymphatic system for drugdelivery. Drug absorption into the lymph has twomain advantages overthe portal route. First, the possibility of targeting high concentrations of anticancer agents into the lymph to prevent metastasis along the lymphatic pathway. Second, reduction of firstpass metabolism by delivery of the drug directly into the general blood circulation. The lymph is the primary route of absorption for lipophilic dietary compounds suchas cholesterol, long-chain fatty acids, and the fatsoluble vitamins. As seenwith vitamin A, adsorption may be improved by administration with lipids. Similarly, it has been suggested that the lymphatic absorption of lipophilic drugs may be stimulated by coadministration of lipids [221]. The oral absorption of heparin was shown to be insignificant in rats and gerbils following the intraduodenal administration in aqueous solution in micellar solutions of monoolein or sodium taurocholate or in sodium taurocholate-stabilized mineral oilemulsion[222]. However, significant absorption was achieved by administration in trioctanoin, corn oil,or peanut oil emulsions stabilized with sodiumtaurocholate. Although no mechanism for the enhanced absorption of heparin was proposed, the apparent dependence of this effect on thedigestibility of the oil used in the emulsion was noted. Thiswas seen again in absorption studies with the glycoprotein, urogastrone [223]. Absorption was unaltered following administration in aqueous solution, 0.2% (w/v) Ween 80 solution, liquid paraffin emulsion, and diethylphthalate emulsion but was significantlyincreased by administration in trioctanoin and in olive oil emulsions. Studies byBloedow and Hayton [224], using lipids rather than emulsions, suggested that generally polar digestible lipids increase the bioavailability of lipophilic, poor watersoluble drugs, whereas nonpolarnondigestible lipids have no effect on the bioavailability but may reduce the rateof absorption [225,226]. The mechanism of intestinal absorption of drugs from oil-in-water emulsion systems has been explored by Kakemi et al. [227-2291 using an in situ large intestinal rat gut loop model. Their results suggested that the partition coefficient of the drug and the absolute volume of the aqueous phase is of importance, with absorption occurring mainly from the aqueous phase. For a drug with a partition coefficient of less than 1, drug transfer into the aqueous phaseis rate limiting rather thantransfer from the water

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to theabsorptive membrane. However,Noguchi et al. [230]proposed that for alipid-soluble, water-insoluble drug, the absorption process consists of adsorption of vehicleoils to the absorptive membrane, partitioning of drugs intothe membrane lipids with simultaneous partition into hydrolyzed oils, release from the membrane lipids or hydrolyzed oils with movement into the inner compartments, and then transport into the portal or lymphatic systems. The absorption of ephedrinefrom orally administered emulsions given to dogs was investigated byLin et al.[231], whoattempted to rationalize their results using the HLB concept. Correlations of ephedrine present in urine in vivo with the total amount released in vitro using a dialysis method were significant at five different HLB values in the range 10-14. Although emulsionsmay enhance oral drug absorption and biological availability, the underlying mechanisms causing this effect remain unclear. Both physiochemical and physiological functions appear to be involved; for example, dissolution rate, oil-water partition coefficient, gastrointestinal mobility, bile flow, lymphatic absorption, and membranepermeability. The exact way in which these factors interact to affect drug absorption will depend largely on the natureof the drug and theoil phase. Oral administration of drugs in emulsions, therefore, seems to have great potential as a dosage form provided the problems of stability and drug release can becontrolled. G. Topical Emulsions

Emulsion systems can be administered to theexternal surfaces of the body and to body cavities as topical formulations in the form of O/W or W/O creams containing drugs. General reviews on the structureof the skin, the absorption of drugs through the skin, and pharmaceutical and cosmetic products for topical administration can be found in the relevant textbooks [232-2341. A good topical formulation should be one in which the dosage form has both physical and chemical stability, has cosmetic acceptability, and also provides the optimum environment for the active ingredient to reach the skin surface [235]. For the corticosteroids, which may be regarded as representative molecules, the topical therapeutic activity may be considered as the result of three interactions: release, penetration, and antiinflammatory activity. These, in turn, result from the interactions between corticosteroid, vehicle, and skin. In the design of topical formulations, all three factors must be considered to optimize drug delivery [233].

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In general, a suitable topical preparation may be produced by:

1. Optimization of the drug concentration in order that all the drug be in the solution 2. Minimizing the amount of solvent used so that thechemical potential of the drug is maximized inthe solution 3. Ensuring that the vehicle components affect the permeability of the stratum corneumin a favorable manner In emulsion systems, the distribution of the drug between the various phases and the total drug concentration will define the overall concentration gradient that exists across the skin. Also, the relative proportions of the lipid and aqueous phases in the emulsionwill affect the degree of hydration of the stratum corneum and hence drug penetration rates [236]. In general, emulsions will be less occlusivethan greasy materials. The pharmaceutical andcosmetic of a topical product is related toits viscosity, which will affect its consistency, spreadability, and extrudability and will determine the rate which at the active drug candiffuse to the outer layers of the stratum corneum [234,237]. If a thick layer is applied to the skin, the drug in the upper regions of a viscous preparation will not reach the skin during the normal application time. Surfactants present in emulsion systems mayalso influence the rateof release from the formulation as well as the rate of absorption [238]. Many publications have indicated that the presence of soaps can enhance the penetration rates of poorly absorbed materials [239-2421. The surfactants . may also promote the diffusion from the base itself and hence influence therapeutic performance[236,242]. However, simple relationships between the HLB and surfactant action could not be established, and it was concluded that the surfactants had a direct action on the protein rather than the lipid components of the stratum corneum [243]. The following types of emulsifying agent are used pharmaceutically:

1. Zonic: examples of anionic surfactants commonlyused are the alkali salts of the higher fatty acids and sulfate esters of the higher fatty alcohols; for example, sodium cetyktearyl sulfate. Cationic surfactants employed include the long-chain quaternary ammonium compounds; for example, cetrimide. If ionic surfactants are included in a formulation, care must be taken that they do not interact with the active ingredient [243]. 2. Nonionic: a large number of nonionic emulsifiers existfor stabilizing topical emulsions. The long-chain alcohols (e.g., stearyl) are very hydrophobic and are used to stabilize W/O emulsions. The

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corresponding polyoxyethylene glycol ethers have higher HLB values and therefore better emulsifying capacity and are used in the preparation of O/W emulsions. The overall properties of nonionic surface-active agents depend onthe relative proportions of the constituent hydrophilic and lipophilic moieties of the molecule (HLB value). 3. Mixture of emulsifiers: to obtain a stable emulsion, it is often useful to use mixed-surfactant systems. A combination provides a more rigid film at the lipid-aqueous interface and an increased reduction in the interfacial tension. An example of a complex emulsifiersystemiscetyl alcohol, sodium lauryl sulfate, and glyceryl monostearate. The rheological properties of semisolid emulsions made frommixed emulsifiers have been studied extensively by Barry and Eccleston [2442471. The system comprising oil-alcohol (usually a mixture of cetyl and stearyl alcohols)-surfactant (cetrimide, sodium lauryl sulfate, or Cetomacrogol) was found to producea viscoelastic structurethatgavea bodying effect to theformulation through the formation of a gel network of liquid crystals. The rheological properties were highly dependent on the quantity and natureof the surfactant and the alcohol. Interestingly, the use of pure alcohols of different chain lengths gave rise to the formation of semisolid products with poor stability. Oil-in-water creams are water miscible and hence washable andmay be used in a range of active ingredients. After application, the film produced is less occlusivethan a W/O emulsion system.They spreadeasily and their advantage is that as the water evaporates, they exert a cooling effect at the skin surface. They also produce little irritation, yet their major disadvantage is that despite their fat content,they dry out. Water-in-oil emulsions have some advantages in certain disease conditions, where an emollient effect on the stratum comeurn is required. They also provide some protection barrier for the skin and generally provide occlusion. The occlusion effect increases the hydration of the stratum of some drugs. corneum and hence enhances the permeation Examples of the effect of formulation on thepercutaneous absorption of methyl nicotinate are shown in Table 9. The data show that aqueous cream BP releases methyl nicotinate more readily than oily cream BP. The table also showsthe effect of decreasing the drugconcentration on thetime of onset of erythema. Simple formulation changes can markedly affect drug release, as shown by comparing the time of onset of erythema after the addition of glycerol to aqueous creamBP.

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Table 9. Percutaneous Penetration of Methyl Nicotinate from Four Emulsion Vehicles Concentration of methyl nicotinate 1 4.1 0.5 0.1 0.05 0.01

Time of onset of erythema (min) aqueous cream BP 3.3 6.3 4.0 6.3 4.6 16.7 5.9 7.4

oily cream BP

aqueous cream

aqueous cream glycerol

+ 40% glycerol + 60%

4.5 5.2 6.8 10.5

4.6 5.6 6.7 11.7 17.6

-

-

Many publications have discussedthe effect of formulation (including emulsion systems) on the activity of topical corticosteroids [248,249]. The factors influencing activity concern the physicochemical characteristics of the steroid and the choice of the vehicle. Simple constants such as the partition coefficient and the solubility of the steroid are important. It appears that the greatest release is obtained from preparations in which the drug is dissolved at its maximal solubility concentration. Other factors, such as the viscosity of the vehicle and the effect of the vehicle on the degree of hydration of the stratum corneum, arealso significant. The influence of vehicle on the penetration of individual potent corticosteroids is also very significant. Marked differences in the activity of the same corticosteroid in preparations of different companies could be attributedto vehicle-dependent effects showing the significance of formulations. Many formulations forcreams and semisolid products areavailable in the literature and patents. The scientific studies, however, concentrate on the various aspects of the release of drugs or active matter from theemulsion to the skin or to various “skins” such as membranes and cells [2502521. Most mechanisms treat the release in terms of Fick‘s second law of diffusion controlled process [250,253,254]. The effect of various nonionic emulsifiers [255-2581 on the release of various antimicrobial agents, various oils and various adjuvants on the rate of release was studied. The release seems to be diffusion controlled [250].

H. Perfluoro Emulsions for Artificial Blood

A formulation for artificial blood (total blood replacement) has been a goal for many investigations [259-2791. Various perfluoro emulsions were tried, including perlluorotributylamine, perfluorobutyltetrahydrofuran,

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perfluoro decalin, perfluoromethyl decalin, and perfluoromethylcyclohexane. The emulsified fluorocarbon provides the necessary oxygenexchange, and the hydroxyethyl starch acts as a plasma expander. The emulsifier isthe polyoxyethylene-polyoxypropylene copolymer (Pluronic F68) [261]. Some encouragingresults were reported by Geyer [261] on complete replacement of blood in rats. The formation of a suitable perfluorochemical emulsion has been difficult, since those oils that provide stable emulsions are not cleared from the body in a satisfactory way and the choice of surfactants is extremely limited [262,263]. Long-term room temperatureshelf stability of ready-for-use concentrated fluorocarbon emulsions is necessary in order fully to exploit their rherapoic potential. The degradation mechanisms of the fluorocarbons, the factors that have impact on their stability and the means of proplonging their shelf life by using fluorinated surfactants or coemulsifiers, have seen carefully studied [259]. Generally, the fluorinated amphiphiles addmuch to the stability. Davis et al. [262,263] found that mixtures of lecithin and poloxamer were the best systems for emulsion stability. Of interest was the fact that the preferred oil, perfhorodecalin, although originally giving a very fine emulsion onemulsification, had poor stability. The fluorocarbon-emulsifying and emulsion-stabilizing properties of F-alkylated surfactants when used alone or in combination with Pluronic F68 or egg yolk phospholipids (EYP), the two surfactants used in the presently developed fluorocarbon emulsions, were investigated. The stability of these emulsions was evaluated by monitoring the increase in average particle sizes and changes in particle size distribution over time [264,265]. Generally speaking, F-alkylated amphiphiles aremuch more efficient as emulsifiers or coemulsifiers or coemulsifiers for fluorocarbons than their hydrocarbon analogues. Emulsions significantly more stable, than those obtained with egg yolkphospholipids (and a fortiori, Pluronic F68) can be obtained with several of the monodisperse F-alkylated amphiphiles even when used alone, which by itself is remarkable; mixtures of amphiphiles (which includes Pluronic and EYP) are indeed known to be considerably more efficient emulsifiers than any of their constituents taken separately. However, the exact prevision of the behavior of individual surfactants is still uncertain. Some structurally closely related surfactants were found to have widely different emulsion-stabilizing characteristics [266,267]. Examples of various distinct types of behavior aregiven below. Stable, medium concentrated (50% w/v; i.e., 27% v/v) to highly concentrated (up to 100% w/v; i.e., 52% v/v) fluorocarbon emulsions were

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468 0.8 0.7

4%

EYP

-

/

L

m

aJ

0.4

M

m

L

aJ

e

0.3

0.2 0.1

I 0

I

I

100

200

Days Fig. 32. Comparison of average particle sue vs time variation for sterilized Foctyl bromideemulsions (90%w/v) prepared(a) with 1% of sodium D-glucose 6-[2(F-octy1)ethyl phosphate]and, (b) with 4%of egg yolk phospholipides(EYP). (After ref. 269.)

obtained with several F-alkylated amphiphiles, including F-alkylated glucosyl-phosphates [268,269], trehalose ester[270], carnitine esters[271], some amino acids of type, amine oxides, phosphocholines, and phosphatidylcholines [272]. Fig. 32 compares, as an example, the average particle size increase over time for a 90% w/v concentrated perfluoro octyl bromide (PFOB) emulsion prepared with the F-alkylated glucosyl-phosphates and with EYF! It is noteworthy that even very low amounts of the fluorinated surfactant result in more stable emulsions than when four times larger amounts of EYP are used [268]. F-Alkylated fluorinated surfactant polymers were shown to be more effective emulsifiers than theirnonfluorinated parent compounds [273]. Long-term shelf lives of perfluordecalin (FDC) emulsions (20% w/v) were, however, not achieved. Somewhat better results were obtained with the F-alkylated Tris (hydroxymethyl) acrylaminomethane (THAM) telomers [274,275]. On the other hand, the anionic poly(ethyleneglyco1) phos-

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phate and theneutral ones were foundto be less efficient emulsifiersthan Pluronic F68 [276]. Further improvement in fluorocarbon emulsion stability is expected when mixtures of surfactants are used, especially if they present complementary attributes. Therefore, F-alkylated amphiphiles havealso been investigated as cosurfactants with Pluronic F68, EYP, or other F-alkylated surfactants. Pluronic F68-type poloxamers are not very effective in reducing the interfacial tension between a fluorocarbon and water, but they exercise considerable steric stabilization. The addition of a fluorinated cosurfactant was expected to compensate forthis lack of effectiveness. Polyhydroxylated surfactants were chosenfor this purpose, because they could hydrogen bond Fto Pluronic. Considerable stabilization was indeedachievedwhen alkylated maltoside- or xylitol-derived surfactants were usedin conjunction with Pluronic F68 [277]. This synergistic stabilization effect is illustrated in Fig. 33 for a 50% w/v concentrated FDC emulsion preparedwith 5% w/v ofa mixture of 2-(F-octyl)ethyl maltoside and Pluronic F68. The stability of the emulsion, which is rather poorwhen Pluronic F68 is used alone, is significantly improved when 20430% of the' poloxamer is replaced by the Falkylated maltoside in the surfactant mixture. It should be noted that this Falkylated surfactant alone (right side of Fig. 33) is totally unable to stabilize the emulsion. The glucosyl-phosphates were found to act synergistically with EYP [268]. Therefore 90% w/v PFOB emulsions were prepared in which the EYP/F-(octy1)ethyl phosphate ratio was decreased stepwise, with the total amount of surfactant being held constant (Fig. 34). The replacement of only one eighth of the EYP by F-(octy1)ethyl phosphate was sufficient to obtain both a definite stabilizing effect and, interestingly, a decrease in particle size which, moreover, is better preserved during sterilization and ageing than in the emulsions preparedwith EYP alone. Synergistic stabilization was also observed when mixtures of two Falkylated phosphocholines were used in the formation of 100% w/v FDC emulsions [278]. These concentrated emulsions were significantly more stable than those formulated with EYP. Mixtures of F-alkylated THAM telomers of different length or of these telomers with the F-alkylated xylitol derivative also led to some improvementsin FDC or PFOB emulsion stability [275].

1. Biological Evaluation of F-Alkylated Amphiphiles (in Vivo Tests) The acute toxicity of F-alkylated surfactants has been evaluated by intravenous injection in mice of about 0.5 mL (i.e., 25 mWkgbody weight, caudal vein) of their solutions and/or dispersions. The reported maximum

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'

0.0 4

0

20

40

60

80

100 %

Maltoside / Pluronic F-68(w/w)

Fig. 33. Average particle sizes initially (0),after 1 month (O), after 1 year (m) at 25"C, in 50% w/v F-decalin emulsion asa function of a ratio of 2-(F-octyl)-ethyl maltoside to Pluronic 68 in thesurfactantmixture, with thetotalamount of surfactant being held constant(5% w/v). tolerated dose (MTD) compatible with the .survival of all injected mice (n=10) and/or lethal dose causing the deathof 50% of the mice (LDS0)are collected in tables [279]. The highest tolerance, by far, is observedfor the F-alkylated double-chain glycerophosphocholines. They display considerably higher intravenous tolerance than the correspondingsingle-chain analogues, with MTD values for compounds superior to 8000 mg/kgbodyweightcompared with 25-125 mgkg for compounds. Obviously, the nature of the hydrophilic head is also important. In the single-chain compound series, the following sequence of increasing tolerance was observed: phosphocholine phosphocholine mono-N-ethyl < amino acid < trehalose sucrose

-

-

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n.xn T I 40°C

x % wlv of 81b

After 6.7 nrollths

T

IL!

L

After 3 m o n t h s

+

I

0.00 0

1

2

3

4

X%

Fig. 34. Averageparticlesizesin 90% perlluorooctyl bromide emulsions, prepared by microfluidation, asa function of the amount, x%, of sodium D-glucose6[2-(F-octyl)ethyl phosphate], with the remaining (4-x)% consisting of egg yolk phospholipides.

< maltose < xylitol < poly(ethy1ene-glycol) phosphate < THAM telomer. The high in vivo tolerance of the poly(ethyleneglyco1) phosphate derivatives is seen further to increase with increasing the size of the polar head and its hydrophilic character. These trends indicate that for similar hydrophobic tails, intravenous tolerance improves along the sequence: zwitterionic single-chain < neutral single-chain < zwitterionic double-chain compounds. It also appears that increased hydrophilic character (from neutral to zwitterionic or toanionic compounds) needs to bebalanced by an increase in hydrophobic character or by an increase of the size of its hydrophilic head. No obvious relation is, however, found between hydrophobic chain length an in vivo toxicitywhatever the hydrophilic head. , All in all, F-alkylated glycerophosphocholines, poly(ethy1ene-glycol) phosphates and THAM telomers, l-O-[3-(F-octyl)-Zpropenyl]xylitol,2(F-octy1)ethyl maltoside, and 6-0-[3'-(F-octyl)propanoyl]trehalose stand out as themost promising candidates for biomedical preparations.

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One of the primary goals for which many of the F-alkylated amphiphiles were synthesized was the obtaining of stable fluorocarbon emulsions to be used as injectable oxygen carriers and in particular as blood substitutes. Consequently, some of the new surfactants were incorporated in emulsions which were tested in isovolemic exchange-perfusion experiments Therein rats. This was the case for 1-0-[3’-(F-octyl)-2’-propenyl]xylitol. fore, adispersion of 0.5& of the octyl-xylitol in grams per literof Pluronic F68 was used to prepare a 10% (W/.) emulsion of lY2-bis(F-butyl)ethene (F-ME). This emulsion was tested on a series of 10 conscious rats whose red blood cell count was reduced to about one third of normal (hematocrit 15%) [280].The behavior of the animals during and after the exchange was comparable to thatof control animals, and the survival ratio was 10/10 after the 6-month observation period. This test was also applied to the mixed fluorocarbon-hydrocarbon “dowel” molecules. One of these compounds, C,Fl,CH=CHCl,H21, was administered intravenously inemulsifiedform at doses 300-600 times larger than those required to stabilize a clinically relevant dose of fluorocarbon emulsion. None of the 33 treated animals died. ”F-NMR (nuclear magnetic resonance) analysis of the organtissues indicated no metabolism; the halfof the compound in the liver (where fluorocarbon droplets are transiently stored) is about 25 5 days. In conclusion, compared with their hydrogenated analogues, fluorinated surfactants display significantly enhanced surface activity both in terms of effectiveness and efficiency and, in particular, strongly reduced water-fluorocarbon interfacial tensions. Fluorinated amphiphiles also show an increased tendency to form organized supramolecular assemblies, including liposomes, and impart unique properties to these assemblies. There is no doubt thattheir potentialas membrane components and modifiers should be further explored. Their fluorocarbon emulsion-stabilizing capacity, when they are used as surfactants andor as cosurfactants, has been demonstrated butremains unpredictable. The assessment of thebiological properties of fluorinated surfactants has, however, hardly begun. Preliminary evaluation indicates somewhat reduced acute toxicity and definitely lower hemolytic activity compared with their hydrogenated analogues. It is now necessary to establish their fate in the organism and to explore theirbiological effects both when alone and when incorporated in emulsions, liposomes, or other supramolecular assemblies. The influence of fluorinated surfactants on the recognition of such particulate matterin vivo deserves special attention. The stage now. is set forsuch investigations. Only few detailed studieshave been published so far concerning the biological evaluation of F-alkylated amphiphiles. Some attention has, how-

*

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ever, been given to F-alkanoic acids because of their many industrial applications related to their thermal and chemical stability and to their antiwetting and surface activity. Some simple biological tests (in vitro action on cell cultures and on red blood cells) and in vivo tests (acute toxicity in mice after intravenous injection) havebeenreported on a range of other F-alkylated amphiphiles. In a few cases, isovolemic blood exchange-perfusion experiments in rats with emulsions containing such surfactants have been performed. Concerning the F-alkanoic acids, these were found to be toxic with, for example, intraperitoneal LD, (rat) values of 189 and 41 mgkg for Foctanoic and F-decanoicacid, respectively. The major symptoms observed include anorexia, marked bodyweight loss, andhepatomegaly with a concomitant increase in peroxisomesand swelling. But the fluorinated metabolites of these F-acids have not been identified yet. It was reported that F-alkanoic acids covalenty bind to proteins both in vivo and in vitro, but the relevance of this binding to toxicity remains unknown. An increased incidence of Leydig cell adenomas was observed in rats fed with ammonium F-octanoate (300 ppm daily) for 2 years; an endocrine-related mechanism was proposed. The use of diverse F-alkanoic acids was nevertheless deemed acceptable for stabilizing micronized inhalation drug suspensions in 1,1,1,2-tetrafluoroethane7one of the CFC substitutes presently being developed. Studies on F-glucopyranosides also indicate the existence of toxicity due torapid hydrolysis and cleavage. These studies indicate that it is preferable to avoid the direct connection of a F-alkyl chain on a metabolic site which would then give either F-alkanoic acids or unstable F-alcohols. It should be preferable to insert a hydrocarbon spacer between the F-alkyl chain and the possible metabolic sites of the polar head.

2. Effect on the Growth and Viability of Cell Cultures (in Vitro Tests) The growth and viability of cells in the presence of the fluorinated surfactants tested are compared with those of control cultures grown under the same conditions without the surfactant. Cell culture tests were used both to assess direct cell toxicity and as a means of monitoring the final purification steps. Different cell strains were used, including lymphocytes, Hela, Molt-4, hybridoma, K562, or Namalva lymphoblastoid cells. No significant toxic effect of the Namalva cell's growth and viability anionic phosphate esters, the was found at a 1 g L concentration for the three 2-(F-octyl)ethyl D-maltoside, the 6-0-[5'-(F-hexyl)pentanoyl]trehalose, and for the 6-O-[ll'-(F-hexyl)undecanoyl]trehalose in spite of their high surface activity.

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3. HemolyticActivity Unexpectedly, avery drastic difference in hemolytic potency was found betweensurfactants with hydrocarbon chains and with fluorocarbon chains of similar length. The hemolytic activity is consistently and considerably lower and often totally suppressed for the F-alkylated surfactants as compar