Polymers for biomedicine : synthesis, characterization, and applications 9781118967904

"This book highlights dynamic developments in polymer synthesis, focusing on synthetic techniques that yield new bi

302 49 13MB

English Pages [603] Year 2017

Report DMCA / Copyright

DOWNLOAD FILE

Polecaj historie

Polymers for biomedicine : synthesis, characterization, and applications
 9781118967904

Table of contents :
Content: List of Contributors ix Part I. Pseudo-Peptides, Polyamino acids and Polyoxazolines 1 Chapter 1 - Characterization of Polypeptides and Polypeptoides- Methods and Challenges 3David Huesmann and Matthias Barz Chapter 2- Poly(2-Oxazoline): The structurally Diverse Biocompatibilizing Polymer 31Rodolphe Obeid Chapter 3- Poly(2-oxazoline) Polymers synthesis, characterization and Applications in Development of Therapeutics 51Randall W. Moreadith and Tacey X. Viegas Chapter 4- Polypeptoid Polymers: Synthesis, Characterization and Properties 77Brandon A. Chan, Sunting Xuan, Ang Li, Jessica M. Simpson and Donghui Zhang Part II - Advanced Polycondensates 121 Chapter 5 - Polyanhydrides: Synthesis and Characterization 123Rohan Ghadi, Eameema Muntimadugu Wahid Khan and Abraham J. Domb Chapter 6 - New Routes to Tailor-Made Polyesters 149Kazuki Fukushima and Tomoko Fujiwara Chapter 7 - Polyphosphoesters: An old biopolymer in a new light 191Kristin N. Bauer, Hisaschi T.C. Tee, Evandro M. Alexandrino and Frederik R. Wurm Part III. Cationically Charged Macromolecules 243 Chapter 8 - Design and Synthesis of Amphiphilic Vinyl Copolymers with Antimicrobial Activity 245Leanna L. Foster, Masato Mizutani, Yukari Oda, Edmund F. Palermo and Kenichi Kuroda Chapter 9 - Enhanced Polyethylenimine-Based Delivery of Nucleic Acids 273Jeff Sparks, Tooba Anwer and Khursheed Anwer Chapter 10 - Cationic graft copolymers for DNA engineering 297Atsushi Maruyama and Naohiko Shimada Part IV. Biorelated polymers by Controlled Radical Polymerization 313 Chapter 11 - Synthesis of (Bio)degradable Polymers by Controlled/ Living Radical Polymerization 315Shannon R. Woodruff and Nicolay V. Tsarevsky Part V. Polydrugs and Polyprodrugs 355 Chapter 12 - Polymerized drugs a novel approach to controlled release systems 357B. Demirdirek, J. J. Faig, R. Guliyev and K.E.Uhrich Chapter 13 - Structural design and synthesis of polymer prodrugs 391Petr Chytil, Libor Kostka and Toma Etrych Part VI. Biocompatibilization of Surfaces 421 Chapter 14 - Polymeric ultrathin films for surface modifications 423Henning Menzel Chapter 15 - Surface Functionalization of Biomaterials by Poly(2-oxazoline)s 457Giulia Morgese and Edmondo M. Benetti Chapter 16 - Biorelated polymer brushes by surface initiated reversible deactivation radical polymerization 487Rueben Pfukwa, Lebohang Hlalele and Bert Klumperman Part VII. Self-assembled Structures and Formulations 525 Chapter 17 - Synthesis of amphiphilic invertible polymers for biomedical applications 527A.M. Kohut, I.O. Hevus, S.A. Voronov and A.S. Voronov Chapter 18 - Bioadhesive Polymers for Drug Delivery 559Eenko Larraneta and Ryan F. Donnelly INDEX

Citation preview

Polymers for Biomedicine

Polymers for Biomedicine Synthesis, Characterization, and Applications

Edited by Carmen Scholz

University of Alabama in Huntsville Alabama, USA

This edition first published 2017 © 2017 John Wiley & Sons, Inc All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by law. Advice on how to obtain permission to reuse material from this title is available at http://www.wiley.com/go/permissions. The right of Carmen Scholz to be identified as the Editor of this work has been asserted in accordance with law. Registered Office John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, USA Editorial Office 111 River Street, Hoboken, NJ 07030, USA For details of our global editorial offices, customer services, and more information about Wiley products visit us at www.wiley.com. Wiley also publishes its books in a variety of electronic formats and by print‐on‐demand. Some content that appears in standard print versions of this book may not be available in other formats. Limit of Liability/Disclaimer of Warranty In view of ongoing research, equipment modifications, changes in governmental regulations, and the constant flow of information relating to the use of experimental reagents, equipment, and devices, the reader is urged to review and evaluate the information provided in the package insert or instructions for each chemical, piece of equipment, reagent, or device for, among other things, any changes in the instructions or indication of usage and for added warnings and precautions. While the publisher and authors have used their best efforts in preparing this work, they make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives, written sales materials or promotional statements for this work. The fact that an organization, website, or product is referred to in this work as a citation and/or potential source of further information does not mean that the publisher and authors endorse the information or services the organization, website, or product may provide or recommendations it may make. This work is sold with the understanding that the publisher is not engaged in rendering professional services. The advice and strategies contained herein may not be suitable for your situation. You should consult with a specialist where appropriate. Further, readers should be aware that websites listed in this work may have changed or disappeared between when this work was written and when it is read. Neither the publisher nor authors shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. Library of Congress Cataloging‐in‐Publication Data Names: Scholz, Carmen, 1963– editor. Title: Polymers for biomedicine : synthesis, characterization, and applications / edited by Carmen Scholz. Description: Hoboken, New Jersey : John Wiley & Sons, Inc., 2017. | Includes bibliographical references and index. Identifiers: LCCN 2017012622 (print) | LCCN 2017005426 (ebook) | ISBN 9781118966570 (cloth) | ISBN 1118966570 (cloth) | ISBN 9781118967935 (Adobe PDF) | ISBN 9781118967881 (ePub) Subjects: LCSH: Polymers. | Polymerization. | Polymers in medicine. | Macromolecules. Classification: LCC QD381 .P61244 2017 (ebook) | LCC QD381 (print) | DDC 610.28/4–dc23 LC record available at https://lccn.loc.gov/2017012622 Cover image: Wiley Cover design by (Molecular structure) © nopparit/Gettyimages; (DNA) Ingram Publishing/Gettyimages Set in 10/12pt Warnock by SPi Global, Pondicherry, India Printed in United States of America 10 9 8 7 6 5 4 3 2 1

v

Contents List of Contributors  ix Part I 

Pseudo‐Peptides, Polyamino Acids, and Polyoxazolines  1

1 Characterization of Polypeptides and Polypeptoides – Methods and Challenges  3 David Huesmann and Matthias Barz 2 Poly(2‐Oxazoline): The Structurally Diverse Biocompatibilizing Polymer  31 Rodolphe Obeid 3 Poly(2‐Oxazoline) Polymers – Synthesis, Characterization, and Applications in Development of POZ Therapeutics  51 Randall W. Moreadith and Tacey X. Viegas 4 Polypeptoid Polymers: Synthesis, Characterization, and Properties  77 Brandon A. Chan, Sunting Xuan, Ang Li, Jessica M. Simpson, Garrett L. Sternhagen, and Donghui Zhang Part II 

Advanced Polycondensates  121

5 Polyanhydrides: Synthesis and Characterization  123 Rohan Ghadi, Eameema Muntimadugu, Wahid Khan, and Abraham J. Domb 6 New Routes to Tailor‐Made Polyesters  149 Kazuki Fukushima and Tomoko Fujiwara 7 Polyphosphoesters: An Old Biopolymer in a New Light  191 Kristin N. Bauer, Hisaschi T.C. Tee, Evandro M. Alexandrino, and Frederik R. Wurm

vi

Contents

Part III 

Cationically Charged Macromolecules  243

8 Design and Synthesis of Amphiphilic Vinyl Copolymers with Antimicrobial Activity  245 Leanna L. Foster, Masato Mizutani, Yukari Oda, Edmund F. Palermo, and Kenichi Kuroda 9 Enhanced Polyethylenimine‐Based Delivery of Nucleic Acids  273 Jeff Sparks, Tooba Anwer, and Khursheed Anwer 10 Cationic Graft Copolymers for DNA Engineering  297 Atsushi Maruyama and Naohiko Shimada Part IV  Biorelated Polymers by Controlled Radical Polymerization  313 11 Synthesis of (Bio)degradable Polymers by Controlled/“Living” Radical Polymerization  315 Shannon R. Woodruff and Nicolay V. Tsarevsky Part V 

Polydrugs and Polyprodrugs  355

12 Polymerized Drugs – A Novel Approach to Controlled Release Systems  357 Bahar Demirdirek, Jonathan J. Faig, Ruslan Guliyev, and Kathryn E. Uhrich 13 Structural Design and Synthesis of Polymer Prodrugs  391 Petr Chytil, Libor Kostka, and Tomáš Etrych Part VI 

Biocompatibilization of Surfaces  421

14 Polymeric Ultrathin Films for Surface Modifications  423 Henning Menzel 15 Surface Functionalization of Biomaterials by Poly(2‐oxazoline)s  457 Giulia Morgese and Edmondo M. Benetti 16 Biorelated Polymer Brushes by Surface Initiated Reversible Deactivation Radical Polymerization  487 Rueben Pfukwa, Lebohang Hlalele, and Bert Klumperman

Contents

Part VII 

Self‐Assembled Structures and Formulations  525

17 Synthesis of Amphiphilic Invertible Polymers for Biomedical Applications  527 Ananiy M. Kohut, Ivan O. Hevus, Stanislaw A. Voronov, and Andriy S. Voronov 18 Bioadhesive Polymers for Drug Delivery  559 Eneko Larrañeta and Ryan F. Donnelly Index  603

vii

ix

List of Contributors Evandro M. Alexandrino

Brandon A. Chan

Max Planck Institute for Polymer Research (MPIP) Mainz, Germany

Department of Chemistry and Macromolecular Studies Group Louisiana State University Baton Rouge, LA, USA

Khursheed Anwer

Celsion Corporation Huntsville, AL, USA Tooba Anwer

University of Alabama at Birmingham Birmingham, AL, USA Matthias Barz

Institute of Organic Chemistry Johannes Gutenberg‐Universität Mainz Mainz, Germany Kristin N. Bauer

Max Planck Institute for Polymer Research (MPIP) Mainz, Germany Edmondo M. Benetti

Laboratory for Surface Science and Technology Department of Materials ETH Zürich Zürich, Switzerland

Petr Chytil

Institute of Macromolecular Chemistry Academy of Sciences of the Czech Republic Prague, Czech Republic Bahar Demirdirek

Department of Chemistry and Chemical Biology Rutgers University Piscataway, NJ, USA Abraham J. Domb

School of Pharmacy‐Faculty of Medicine The Hebrew University of Jerusalem, and Jerusalem College of Engineering (JCE) Jerusalem, Israel Ryan F. Donnelly

School of Pharmacy Queen’s University Belfast Belfast, UK

x

List of Contributors

Tomáš Etrych

Ivan O. Hevus

Institute of Macromolecular Chemistry Academy of Sciences of the Czech Republic Prague, Czech Republic

Department of Coatings and Polymeric Materials North Dakota State University Fargo, ND, USA

Jonathan J. Faig

Department of Chemistry and Polymer Science Stellenbosch University Matieland, South Africa

Department of Chemistry and Chemical Biology Rutgers University Piscataway, NJ, USA Leanna L. Foster

Macromolecular Science and Engineering Center University of Michigan Ann Arbor, MI, USA Tomoko Fujiwara

Lebohang Hlalele

David Huesmann

Institute of Organic Chemistry Johannes Gutenberg‐ Universität Mainz Mainz Germany Wahid Khan

Department of Chemistry University of Memphis Memphis, TN, USA

Department of Pharmaceutics National Institute of Pharmaceutical Education and Research (NIPER) Hyderabad, India

Kazuki Fukushima

Bert Klumperman

Department of Polymeric and Organic Materials Engineering Yamagata University, Yonezawa, Yamagata, Japan Rohan Ghadi

Department of Pharmaceutics National Institute of Pharmaceutical Education and Research (NIPER) Hyderabad, India Ruslan Guliyev

Department of Chemistry and Chemical Biology Rutgers University Piscataway, NJ, USA

Department of Chemistry and Polymer Science Stellenbosch University Matieland, South Africa Ananiy M. Kohut

Department of Organic Chemistry Lviv Polytechnic National University Lviv, Ukraine Libor Kostka

Institute of Macromolecular Chemistry Academy of Sciences of the Czech Republic Prague, Czech Republic

List of Contributors

Kenichi Kuroda

Randall W. Moreadith

Macromolecular Science and Engineering Center and Department of Biologic and Materials Sciences School of Dentistry University of Michigan Ann Arbor, MI, USA

Serina Therapeutics Huntsville, AL, USA

Eneko Larrañeta

School of Pharmacy Queen’s University Belfast Belfast, UK Ang Li

Department of Chemistry and Macromolecular Studies Group Louisiana State University Baton Rouge LA, USA Atsushi Maruyama

Department of Life Science and Technology Tokyo Institute of Technology Yokohama, Japan Henning Menzel

Institut für Technische Chemie Technische Universität Braunschweig Braunschweig, Germany Masato Mizutani

Department of Chemistry and Chemical Biology Baker Laboratory Cornell University Ithaca, NY, USA

Giulia Morgese

Laboratory for Surface Science and Technology Department of Materials ETH Zürich Zürich, Switzerland Eameema Muntimadugu

Department of Pharmaceutics National Institute of Pharmaceutical Education and Research (NIPER) Hyderabad, India Rodolphe Obeid

R&D/Process Development & Manufacturing Scale‐Up, IntelGenx Corp. St. Laurent Quebec Canada Yukari Oda

Department of Applied Chemistry Kyushu University Fukuoka, Japan Edmund F. Palermo

Department of Materials Science and Engineering Rensselaer Polytechnic Institute Troy, NY, USA Rueben Pfukwa

Department of Chemistry and Polymer Science Stellenbosch University Matieland, South Africa

xi

xii

List of Contributors

Naohiko Shimada

Andriy S. Voronov

Department of Life Science and Technology Tokyo Institute of Technology Yokohama, Japan

Department of Coatings and Polymeric Materials North Dakota State University Fargo, ND, USA

Jessica M. Simpson

Stanislaw A. Voronov

Department of Chemistry and Macromolecular Studies Group Louisiana State University Baton Rouge, LA, USA

Department of Organic Chemistry Lviv Polytechnic National University Lviv, Ukraine

Jeff Sparks

Tacey X. Viegas

Celsion Corporation Huntsville, AL, USA

Serina Therapeutics Huntsville, AL, USA

Garrett L. Sternhagen

Shannon R. Woodruff

Department of Chemistry and Macromolecular Studies Group Louisiana State University Baton Rouge, LA, USA

Department of Chemistry and Center for Drug Discovery, Design, and Delivery, Southern Methodist University Dallas, TX, USA

Hisaschi T.C. Tee

Max Planck Institute for Polymer Research (MPIP) Mainz, Germany Nicolay V. Tsarevsky

Frederik R. Wurm

Max Planck Institute for Polymer Research (MPIP) Mainz, Germany

Department of Chemistry and Center for Drug Discovery, Design, and Delivery, Southern Methodist University Dallas, TX, USA

Sunting Xuan

Kathryn E. Uhrich

Donghui Zhang

Department of Chemistry and Chemical Biology Rutgers University Piscataway, NJ, USA

Department of Chemistry and Macromolecular Studies Group Louisiana State University Baton Rouge, LA, USA

Department of Chemistry and Macromolecular Studies Group Louisiana State University Baton Rouge, LA, USA

1

Part I Pseudo‐Peptides, Polyamino Acids, and Polyoxazolines

3

1 Characterization of Polypeptides and Polypeptoides – Methods and Challenges David Huesmann and Matthias Barz Institute of Organic Chemistry, Johannes Gutenberg‐Universität Mainz, Mainz, Germany

1.1 ­Introduction Materials made from polypeptides, and recently also polypeptoids, have received considerable and growing attention in recent years. Since synthetic polypeptides, just like natural proteins, are made up of amino acids, they can be non‐toxic, biocompatible, and degradable in the body while they remain stable in aqueous solution. The multitude of different side chains enables the design of peptidic superstructures like polyion complexes [1,2], polymer micelles [3,4], polymer vesicles [5,6], nanofibers or ‐tubes [7], and hydrogels [8]. Apart from exactly defined polypeptides (i.e., proteins) that show a defined sequence of amino acids, there are also natural polypeptides that resemble less  defined classic synthetic polymers. One of these polypeptides is poly(ɣ‐ glutamic acid) [9,10], which is produced by bacteria and cnidaria [11]. It is the major constituent of nattō (Japanese food from fermented soy beans) and approved by the FDA for cosmetic applications.

1.2 ­Synthesis of Poly(peptide)s Synthetic polypeptides were first described by Leuchs in the beginning of the twentieth century, although their polymeric nature was not acknowledged at that time [12–14]. Many researchers have explored synthetic polypeptides through the twentieth century [15,16] partially with poor results regarding polymerization kinetics, end‐group integrity, or dispersity, in particular with more complex systems such as block copolypeptides, star‐like polypeptides, or bottle‐brush polymers. The reasons for this are manifold, including monomer Polymers for Biomedicine: Synthesis, Characterization, and Applications, First Edition. Edited by Carmen Scholz. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

4

Polymers for Biomedicine

purity, monomer instability over prolonged periods of time and the fact that the polymerization does not necessarily follow a single mechanism (Figure 1.1). The most prominent competing pathways are the normal amine mechanism (NAM, which leads to a classical chain growth) and the activated amine mechanism (AMM, which leads to undefined polymers through condensation of polymer chains). Further, addition of an N‐carboxyanhydride, NCA, monomer O R′ Rearrangement –CO2 O R′

N H

R

O

O

O

O O Carbamate

N H

NH2

N H

R

OH +NCA

n

+NCA O

R′

H2NR′

O

OO +



N R

N H

O

O

O

NH R

O

O R′

O

H N R

OH

R′

–CO2

NH2

N H

O

R

AMM

O

O

OH

R R O Urea Derivative

NCA

–R′NH3+

O

H N

O

NAM

O NH

R

R

H N

N H

H N

O

O N

R

O

H N R

O– –CO2 O

O

O NH–

N

O

R

R

+NCA

O– O

N C O

O

R Isocyanate

O N

O

R

R O

Condensation Products

O RNH2 O

O N



R

O

H N

N R

O

NH2 O +

R

H n

Figure 1.1  Mechanisms of NCA polymerization: Normal amine mechanism (NAM) and activated monomer mechanism (AMM).

Characterization of Polypeptides and Polypeptoides

before decarboxylation can lead to carbamates, which can rearrange into urea units, while a deprotonated NCA can open to form an isocyanate. A more in‐ depth discussion of the reaction mechanism is outside of the focus of this chapter and can be found in excellent reviews and books [15–17]. The complex reaction mechanism has led to the development of controlled NCA polymerization methods starting in the end of the last century. In the late 1990s, the group of Timothy Deming was the first to demonstrate that the NCA polymerization using transition metal catalysts proceeds in a living manner and yields well‐defined polypeptides (Figure 1.2) [18]. While this approach has been very successful for the preparation of well‐defined and complex polypeptide architectures [5,6,19], it has the need for a transition metal catalyst. Additionally, the synthesis of hybrid structures remains challenging since the transition metal catalyst needs to be modified [20]. As a complementary approach, Cheng and coworkers have reported silylated amine initiators, which allow control over NCA polymerization [22,23]. The trimethylsilyl residue remains at the polymer terminus over the course of the polymerization, allowing the preparation of defined polypeptides (Figure 1.3). The rate of polymerization is not slowed down by this polymerization technique as the polymerization (M/I = 300) was reported to be completed within 24 h or less. Amine initiated polymerization has been reported to be complete Initiation: (L)nM

O +

O

–CO

HN

O

R HN

N M (L)n R

O

O N H

Propagation: R HN

proton migration

R

O NH

R NCA –CO2 R O HN M(L)n NH R R HN O

R

O N

HN NCA M(L)n –2 CO2

HN

O

M = Co, Ni

R

(L)n M O

R

R

NCA

Polymer M (L)n

O

HN

M(L)n NH R Polymer N

proton migration

R

O

HN

N

M (L)n R

O N H

Polymer

O

Figure 1.2  Initiation and propagation of metal catalyzed NCA polymerization. Source: Deming 2000 [21]. Reproduced with permission of American Chemical Society.

5

6

Polymers for Biomedicine O R HN

Si

O O H2N O

HN Si

H Si

TMS

O

H N

O

Si NH2

O

R

N H n

O

TMS

O O

N

Si

O TMS

O

R

H N

N

TMS

R

H N O

TMS NCA

O N H O

R

O SiMe3

O

R

H N O

N H

O

TMS

O N H

O

Figure 1.3  Mechanism of trimethylsilyl‐mediated NCA polymerization. Source: Lu 2007 [22]. Reproduced with permission of American Chemical Society.

within the same time frame (17 h for Xn = 438, poly(benzyl glutamic acid), (PGlu(OBn)) [24]. On the other hand, several approaches have been investigated to optimize the conditions of conventional amine initiated NCA polymerization. Vayaboury et al. used non‐aqueous capillary electrophoresis to show a dramatic increase of living chain ends by lowering the polymerization temperature to 0 °C [25]. Unfortunately, no GPC plots and polymer dispersities of the obtained polymers were presented. Heise and coworkers investigated the influence of reduced temperature further [26] and used vacuum for the removal of CO2 from the reaction to increase its speed [27]. CO2 liberation is a step in NCA polymerization, which depends highly on the pressure in the reaction vessel. Wooley and coworkers reported the removal of CO2 by nitrogen flow through the reaction mixture, thereby increasing also the polymerization speed [28]. Both findings are surprising since theoretical studies and experiments have shown that CO2 liberation is not the rate determining step of the polymerization [15,29–31]. However, the performed control polymerizations (no nitrogen flow) yielded polypeptides with high dispersities of 1.38 and 2.19 for Xn of 50 and 100, respectively, while dispersities of PGlu(OBn) initiated by primary amines are usually well below 1.2 [32]. Scholz and Vayaboury tackled the issue of different secondary structures in the growing peptide by introducing thiourea to suppress hydrogen‐bond formation [33]. It was found that the dispersity of polypeptides decreased markedly, independent of whether macroinitiators (PEG‐NH2) or low molar mass initiators (hexylamine) were used.

Characterization of Polypeptides and Polypeptoides

In a different approach, Schlaad and coworkers introduced HCl salts of ­ rimary amines as initiators, lowering the reactivity of the growing chain end [34]. p Elevated temperatures (40–80 °C) were used to counteract the slow polymerization. This method was complemented by other ammonium salts, namely different acetates [35] and recently the non‐nucleophilic tetraflouroborates by Vicent and coworkers [36]. Finally, Hadjichristidis and coworkers reported on the use of highly purified  monomers, solvents and reagents under high vacuum techniques [24]. Interestingly, these results suggest that all the previously mentioned potential side reactions are impurity related and that control can be achieved by working with highly pure solvents, monomers, and initiators.

1.3 ­Characterization of Poly(peptide)s The aim for better and better control over the NCA polymerization over the last century was also accompanied by the development of analytical methods that allowed for a better characterization of polypeptidic materials. As with many classes of polymers, polypeptides are often characterized by the most widely used analytical techniques NMR and GPC to determine composition, size and dispersity of the polymers. However, due to the periodical peptide bond in the polypeptide backbone, these polymers are often not in a random coil conformation – as is usually the case for other polymers. This leads to two major challenges in the characterization of polypeptides: (1) The secondary structures must be characterized using for example NMR, IR, CD spectroscopy, or X‐ray diffraction and (2) the different secondary structures lead to a change in the hydrodynamic radius of the polymers, limiting the usefulness of  methods that rely on the hydrodynamic radius to deduce other physical parameters (e.g., GPC). It is worth noting, that these challenges do not apply to most polypeptoids, since they lack the free hydrogen at the amide bond and ­therefore do usually not form secondary structures. The combination of a complex polymerization mechanism with many ­potential side reactions on one hand and challenging characterization on the  other calls for extremely careful interpretation of obtained data. In the ­following sections, we will introduce different analytical methods for analyzing ­polypeptides highlighting their advantages and limitations.

1.4 ­Gel Permeation Chromatography (GPC) Gel permeation chromatography is certainly one of the most important analysis methods in polymer chemistry yielding not only average molecular weights, but also a value for polymer dispersity, describing the width of the molecular

7

8

Polymers for Biomedicine

weight distribution. However, these molecular weight distributions are often not obtained directly, but indirectly using a calibration by polymer standards. The separation in the GPC column itself is enabled by polymer beads with different pore sizes. Large molecules cannot enter the pores and elute first from the columns, while smaller molecules can diffuse into the pores, thus remaining in the column for a longer time. However, the separation does not occur by molecular weight, but by polymer size (i.e., hydrodynamic volume) and molecular weight is only inferred from calibration. To obtain correct molecular weights from this method, two conditions should be fulfilled: (1) The polypeptide has to be in one conformation and (2) the standards for the calibration have to be the same polymer (or at least very similar in structure) and have to exhibit the same secondary structure as the polymer measured. Both conditions are virtually never fulfilled when working with polypeptides. Even very good solvents for polypeptides like dimethylformamide (DMF), dimethylacetamide (DMAc), N‐methylpyrolidone (NMP), and hexafluoroisopropanol (HFIP) are usually not able to suppress secondary structures in protected polypeptides, leading to a strong change in hydrodynamic volume [37]. Further, the standards used for calibration are in many cases poly(ethylene glycol) (PEG) or poly(methyl methacrylate) (PMMA), both molecular structures are far from that of polypeptides (Figure 1.4). Therefore, it appears reasonable to produce GPC standards for each class of polypeptides, as done by Hadjichristidis and coworkers. They synthesized narrowly distributed PGlu(OBn) and determined their molecular weight (Mn) by membrane osmometry. They then used these samples for the calibration of their GPC [24]. Using a calibration with polypeptide standards, the molecular weights from GPC measurements can be used to create a meaningful kinetic plot (Figure 1.5). Unfortunately, calibration with polypeptides is seldom performed, since polypeptide standards are not commercially available. In some cases, bimodal molecular weight distributions can be observed due to a change in secondary structure, which results in a pronounced change in hydrodynamic volume. This can, for example, be seen in growing poly(N‐ ε‐benzyloxycarbonyl‐L‐lysine) (PLys(Z)) chains that undergo a transition in secondary structure at a degree of polymerization around 15 [37]. Observing the polymer at different time points of the polymerization shows a monomodal distribution (random coil) changing to a bimodal distribution when two ­secondary structures (random coil and α‐helix) are present (Figure 1.6). Once O H3C

O

O

PMMA

R

H N O

PEG

Polypeptide

Figure 1.4  Molecular structures of common GPC standards compared to polypeptides.

Characterization of Polypeptides and Polypeptoides

2.4 × 10–2

[M]/[I] = 743

1,6 × 105 1,4 × 105 1,2 × 105

kp, obs (1 mol–1 min–1)

2.0 × 10–2

[M]/[I] = 438

1,0 × 105 8,0 × 104

1.6 × 10–2

[M]/[I] = 223

6,0 × 104 4,0 × 104

[M]/[I] = 110

2,0 × 104 0,0

1.2 × 10–2

[M]/[I] = 52 10 20 30 40 50 60 70 80 90 100 Polymer conversion

8.0 × 10–3 4.0 × 10–3 0.0 9.0 × 10–3 1.8 × 10–2 2.7 × 10–2 3.6 × 10–2 4.5 × 10–2 5.4 × 10–2

0.0

[I], mol–1

Figure 1.5  Kinetic plot from the amine initiated polymerization of poly(glutamic acid). Molecular weights were obtained from GPC data, calibrated with poly(glutamic acid) standards. Source: Aliferis 2004 [24]. Reproduced with permission of American Chemical Society.

Polymerization time

1 0.8

RID

0.6 0.4 0.2 0 16

18

20

22 Vol/mL

24

26

28

Figure 1.6  DMF GPC of a growing PLys(Z) chain showing the transition from random coil to α‐helix. Source: Huesmann 2014 [37]. Reproduced with permission of American Chemical Society.

9

Polymers for Biomedicine 1

PLys(Z)25 PLys(Z)50

0.8

PLys(Z)100 PLys(Z)200

0.6 RID

10

0.4

0.2

0 16

17

18

19

20

21

Vol/mL

Figure 1.7  HFIP GPCs of PLys(Z), from right to left: P(Lys(Z)25, P(Lys(Z)50, P(Lys(Z)100, P(Lys(Z)200. Source: Huesmann 2014 [37]. Reproduced with permission of American Chemical Society.

all chains have reached the appropriate length to form only α‐helices, the distribution becomes monomodal again. The same behavior is also visible for PLys(Z) of different degrees of polymerization (Figure 1.7). An estimation of the hydrodynamic radius (Rh) of a PLys(Z) random coil (worm‐like chain model) and α‐helix (cylinder) with DP = 15 shows an increase in hydrodynamic volume of α‐helix over random coil [37]. Taking this into account, it is clear that molecular weights of polypeptides from GPC have to be treated with extreme care. A molecular weight obtained with PMMA standards should never be treated as the real molecular weight of polypeptides, nor should every broadening or bimodal molecular weight distributions be directly related to poorly defined polymers. Bimodal molecular weight distributions are not necessarily caused by side reactions but might be attributed to the coexistence of different secondary structures. Vice versa, it would not be wise to infer the coexistence of different secondary structures just from bimodal GPC distributions. Thus, in addition to standard GPC ­analytics it seems highly beneficial to investigate solution conformation by complementary characterization methods.

Characterization of Polypeptides and Polypeptoides

1.5 ­Infrared (IR) Spectroscopy Fourier transformed infrared (FT‐IR) spectroscopy is one of the oldest analytical methods in organic chemistry and has become an established tool for the structural characterization of proteins. Using infrared light (wave numbers 4000–400 cm−1) vibrational modes in molecules are exited, leading to the absorption of light of a characteristic wavelength. Samples can be measured in solution or in the solid state. The use of attenuated total reflection (ATR) units allows easy measurements of small sample volumes. For the analysis of polypeptides, the absorption of the amide I band (CO stretching, around 1650 cm−1) and amide II band (NH bending, CN stretching, around 1550 cm−1) are typically used. The typical amide I frequencies of protein secondary structures are shown in Table 1.1. α‐Helices show absorption between 1650–1657 cm−1, while random coils shift the absorption to lower wave numbers (1640–1651 cm−1) and turns show an adsorption at higher wave numbers (1655– 1675 cm−1, 1680–1696 cm−1). The distinction of parallel and anti‐parallel β‐sheets is not easy, since their absorption is very similar. Parallel β‐sheets absorb at 1626–1640 cm−1 and anti‐parallel β‐sheets at 1612–1640 cm−1. However, anti‐ parallel β‐sheets have a second, weak band at 1670–1690 cm−1, which allows discrimination between the two conformations. It should be stressed that these numbers are guidelines and may vary with changing amino acids [38,39]. IR spectra can of course only provide a qualitative picture of the secondary structures present in the polypeptide. Further, the bands are sensitive to the applied solvent and its water content. For the characterization of synthetic polypeptides organic solvents are often necessary to dissolve protected polypeptides like DMF or DMAc. These solvents have strong amide bond Table 1.1  Typical amide I frequencies of different secondary structures. Source: Pelton 2000 [40]. Reproduced with permission of Elsevier. Conformation

Amide I/cm−1

α‐helix

1650–1657

β‐sheet (anti‐parallel)

1612–1640, 1670–1690 (weak)

β‐sheet (parallel)

1626–1640

turn

1655–1675, 1680–1696

random coil

1640–1651

11

Polymers for Biomedicine

absorption bands themselves and can therefore not be used when looking at secondary structures. Water is also problematic, due to absorption at 1650 cm−1, which can be suppressed using high peptide concentrations, short path length and careful baseline subtraction. Urea and thiourea absorb between 1600– 1700 cm−1 and also trifluoroacetates are problematic due to their absorption at 1673 cm−1. HFIP on the other hand, does not absorb between 1500–3000 cm−1 and can easily dissolve most polypeptides. Whenever measurements are performed in dry state, it should be kept in mind that there might be a pronounced change in secondary structure due to the absence of interaction with the ­solvent, which leads to a partial or complete collapse of the polypeptide. Zhang and coworkers synthesized a mannose modified poly(glutamic acid) which, like its precursors, adopts an α‐helical conformation [41]. Among other techniques, IR spectroscopy was used to confirm this conformation in the solid state (Figure 1.8). HO

O

H N

O

NaN3, DMF n

HO HO

O

H N

60 °C, 48 h Cl

O

O

OH O H N

PMEDTA, CuBr DMF, rt, 24 h N3

O

A

O

O

C n

B

amide II

O

OH OH

n N N N

O

D

amide I

νC-H

νC=O

O O

OH OH

νO/N-H

νO-H

D

Absorbance (a.u.)

12

νalkyne C νN=N=N

B A 1000

1500

2000

2500

3000

3500

Wave number (cm–1)

Figure 1.8  IR spectrum of mannose modified poly(glutamic acid) as well as its precursors, showing typical amide I and II bands of an α‐helical conformation. Source: Tang 2010 [41]. Reproduced with permission of American Chemical Society.

Characterization of Polypeptides and Polypeptoides N-hexylamino-p(L-Lys) N-hexylamino-p(L-Cys) N-hexylamino-p(L-Glu)

95

85 α-helix β-sheet α-helix

Lysine Cysteine

75 Glutamate

65

1850

3650

3150

1650

2650

1450

2150

55

1650

1150

45 650

Figure 1.9  IR spectrum of protected lysine, cysteine, and glutamic acid 20‐mers. The arrows indicate the amide I band. Source: Ulkostki 2013 [42]. Reproduced with permission of American Chemical Society.

Scholz’s group investigated the secondary structure of oligopeptides [42]. IR was used to identify the different secondary structures of 20‐mers of protected lysine, cysteine, and glutamic acid. A shift of the amide I band can be seen between the α‐helical lysine and glutamic acid on one hand and the cysteine in a β‐sheet conformation on the other hand (Figure 1.9). Another very useful application of IR spectroscopy in NCA polymerization lies in measuring the progress of the reaction. The monomer conversion can be easily visualized by the disappearance of the strong absorption band of the NCA at around 1860 and 1785 cm−1 (in DMF) (Figure 1.10). Like FT‐IR spectroscopy, Raman spectra can be recorded by measuring the scattered instead of the absorbed light. In this case, different selection rules apply and different bands might be visible. In conclusion, IR‐spectroscopy is a very powerful and easy to use tool to get a qualitative insight into the secondary structure of polypeptides, while quantitative interpretation needs further treatment of the raw data by peak deconvolution and is strictly only applicable to natural proteins. When measuring in solution, the absorption of the solvent must be kept in mind, while the dry state may not represent the conformation of the polypeptide in solution.

13

Polymers for Biomedicine 1.2

1

Transmittance

14

DMF Monomer in DMF

0.8

0.6

Polymer in DMF 1.1 1.1

1.05

0.4

1

1.05

0.95 0.9

0.2 2000

0.85

1 1900

1800

1850

1800

1600

1750

1400 Wavenumber

1000

950

1200

900

1000

850

800

600

(cm–1)

Figure 1.10  Useful IR NCA peaks to control monomer conversion (Lys(Z) in DMF).

1.6 ­Nuclear Magnetic Resonance (NMR) Spectroscopy NMR spectroscopy is a complementary technique commonly used in polymer synthesis, as well as protein structure determination. The technique is based on the absorption of electromagnetic radiation by nuclei with a spin different from zero (1H, 13C, 15N) in a strong magnetic field. In this section, only the application of relatively simple 1D NMR experiments for the determination of chain length, composition, and secondary structure will be discussed. More complex or multidimensional NMR techniques for the determination of protein tertiary structures like NOESY will not be discussed here but can be found in literature [43,44]. The simplest application is the determination of chain length, by comparing protons of the initiator or end group to protons of the repeating unit. To receive accurate results from this method, all chains need to carry the reference groups. To reference to the initiator, all chains should be started by the initiator and not by impurities in the solvent (e.g., water). For the end group the post polymerization modification reaction performed needs to be quantitative. Further, the reference signals have to be well separated from the signals of the repeat unit to allow separate integration. To receive an accurate measurement even for larger polypeptides, a strong initiator signal is beneficial. Neopentylamine has proven to be a very reliable initiator that features nine equivalent protons,

Characterization of Polypeptides and Polypeptoides

b b

b

O

c O

NHb f f

g

e af H d N NH2 n O

b b g g

9H

0.82

840 H

a,b

c

d

e

f

g

8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 𝜕 (ppm)

Figure 1.11  NMR of poly(N‐ε‐benzyloxycarbonyl‐L‐lysine) (PLys(Z)), degree of polymerization 420, initiator: neopentylamine.

which do not overlap with any amino acid signals [37]. Using this initiator, polypeptides with length of more then 420 repeating units can be identified by NMR as can be seen in Figure 1.11. NMR spectroscopy can further be used to determine the secondary structure of polypeptides in solution. Especially the α‐position of amino acids is very sensitive to changes in the secondary structure. This can be seen in the chemical shift of the α‐proton as well as the α‐carbon in 1H and 13C NMR spectroscopy, respectively [45–47]. The chemical shifts for 13Cα, 13Cβ, 13C’, 15N, 1 HN, and 1Hα in α‐helix, β‐sheet, and random coil conformation have been collected and can be used for easy reference (Table 1.2) [47]. These chemical shifts given in Table  1.2 are attributed to unprotected amino acids in water, but  only shift slightly when measuring protected polypeptides in organic ­solvents [37]. The effect of secondary structure in a growing PLys(Z) chain can be observed in Figure 1.12 (left). While the NMR spectrum shows two Hα peaks for short PLys(Z) the peak of the random coil disappears with increasing degree of polymerization. 13 C‐NMR spectroscopy was extensively applied for the study of secondary structures in acetonitrile, TFA and in the solid state [48–50], as well as for sequence analysis [51] by Kricheldorf and coworkers. Figure 1.12 (right) shows a solid state 13C CP/MAS (cross polarization/magic angle spinning) NMR spectrum of poly(norvaline). The peaks corresponding to Cα, Cβ, and C’ show split peaks indicating the coexistence of α‐helix and β‐sheets.

15

Table 1.2 Statistically derived chemical shifts in secondary structures. Source: Wishart 2011 [47]. Reproduced with permission of Elsevier. 13

13



13



15

C’

1

N

1

HN



Residue

Helix

Sheet

Coil

Helix

Sheet

Coil

Helix

Sheet

Coil

Helix

Sheet

Coil

Helix

Sheet

Coil

Helix

Sheet

Ala

54.8

51.5

52.8

18.3

21.1

19.1

179.4

176.1

177.7

121.4

124.5

123.6

8.08

8.44

8.15

4.03

4.77

Coil

4.26

Cys (ox)

58.0

55.0

55.6

39.4

43.9

41.0

176.2

173.6

174.9

117.7

121.0

118.0

8.20

8.80

8.25

4.15

5.15

4.65

Cys (red)

61.3

56.9

57.5

27.8

30.2

29.4

176.2

173.6

174.9

117.7

121.0

118.0

8.20

8.80

8.25

4.15

5.15

4.65

Asp

56.7

53.9

54.2

40.5

42.3

40.9

178.0

175.5

176.3

119.2

122.2

120.0

8.18

8.51

8.36

4.43

4.94

4.60

Glu

59.1

55.5

56.9

29.4

32.0

30.2

178.6

175.4

176.4

119.0

122.1

120.4

8.22

8.53

8.37

4.01

4.78

4.28

Phe

60.8

56.7

58.0

38.8

41.5

39.5

177.1

174.3

175.6

119.2

121.1

119.7

8.18

8.75

8.17

4.16

5.09

4.54

Gly

46.9

45.2

45.5

N/A

N/A

N/A

175.5

172.6

173.9

107.5

109.3

109.1

8.29

8.34

8.33

3.81

4.20

3.96

His

59.0

55.1

55.9

29.5

31.9

30.0

177.0

174.2

174.8

118.0

120.5

118.7

8.10

8.62

8.21

4.33

5.06

4.53

Ile

64.6

60.1

61.0

37.6

39.9

38.7

177.7

174.9

175.6

119.7

122.8

120.9

8.02

8.68

7.98

3.67

4.68

4.15

Lys

58.9

55.4

56.6

32.3

34.6

32.8

178.4

175.3

176.3

119.2

122.2

120.5

7.99

8.48

8.23

3.99

4.69

4.26

Leu

57.5

54.1

54.9

41.6

43.8

42.4

178.5

175.7

176.9

119.6

124.1

121.5

8.05

8.60

8.08

4.00

4.82

4.36

Met

58.1

54.6

55.7

32.3

35.1

33.4

178.0

174.8

175.4

118.2

121.7

119.7

8.09

8.64

8.18

4.07

4.96

4.38

Asn

55.5

52.7

53.2

38.6

40.1

38.6

176.9

174.6

175.1

117.3

121.6

118.2

8.22

8.60

8.40

4.48

5.06

4.66

Pro

65.5

62.6

63.5

31.5

32.3

31.9

178.3

176.2

176.9

N/A

N/A

N/A

N/A

N/A

N/A

4.22

4.60

4.37

Gln

58.5

54.8

56.1

28.5

31.3

29.1

178.0

174.9

175.9

118.4

121.1

119.5

8.04

8.48

8.23

3.99

4.80

4.26

Arg

58.9

55.1

56.4

30.1

32.2

30.7

178.3

175.1

176.0

118.9

122.3

120.4

8.07

8.56

8.25

3.99

4.74

4.24

Ser

60.9

57.5

58.4

63.1

65.2

64.0

175.9

173.6

174.5

114.9

116.9

115.6

8.14

8.50

8.23

4.25

4.91

4.47

Thr

65.6

61.1

61.6

68.9

70.8

70.1

175.9

173.7

174.7

114.6

116.5

113.4

8.04

8.51

8.16

4.00

4.86

4.45 4.12

Val

66.2

60.8

62.1

31.5

33.9

32.7

177.7

174.8

175.7

119.2

121.9

119.8

8.02

8.62

8.04

3.58

4.60

Trp

60.0

56.4

57.8

29.3

31.5

29.7

178.1

175.4

176.2

119.8

122.1

120.2

8.12

8.59

7.92

4.38

5.19

4.55

Tyr

61.0

56.8

58.0

38.3

41.0

39.0

177.4

174.5

175.4

119.2

121.4

119.5

8.07

8.68

8.06

4.09

5.10

4.52

Characterization of Polypeptides and Polypeptoides

Xn d CH2

200 100

b CH2

..

f

e

βs

x αh

4.2

d

NH CH CO NH CH2 n a x

50 25

c

c CH2

αh

βs f

ss

a

b αh βs

ss

3.8 δ (ppm)

δ (ppm)

150

100

50

0

Figure 1.12  Left: α‐proton in growing PLys(Z) chains changing from random coil to α‐helix. Source: Huesmann 2014 [37]. Reproduced with permission of American Chemical Society. Right: 13C CP/MAS spectrum of poly(norvaline) (DP = 20). The split peaks indicate different secondary structures. Source: Kricheldorf 1983 [48]. Reproduced with permission of American Chemical Society.

In summary, it can be concluded that NMR is an excellent technique to not only investigate the length of polypeptides but also to get an insight into their secondary structure. The fact that (block) copolypeptides contain far less amino acids than natural proteins allows the determination of secondary ­structures already with very simple methods like 1H NMR.

1.7 ­Circular Dichroism (CD) Spectroscopy CD spectroscopy is one of the oldest methods to determine secondary s­ tructures in proteins. The principle is based on the difference in absorbance of right and left handed circular polarized UV‐light by chiral molecules. Specifically the absorptions of the amide bond between 180–250 nm (excitations: n → π* around 190 nm; p → π* around 220 nm) are of interest. The reference spectrum used most often is a polylysine spectrum (Figure 1.13) since polylysine can be forced into the three different conformations under different conditions (random coil: pH 7, α‐helix: pH 11, β‐sheet: pH 11, and 50 °C for 10 min) [52]. Using reference spectra like these, experimental CD spectra can be fitted to determine the content of the different secondary structures [53–55]. However, caution must be taken, as this method assumes that secondary structures in the reference proteins are the same in crystals (X‐ray crystallography) and solution (CD spectroscopy). Further, it has to be kept in mind that these

17

Polymers for Biomedicine

50

α - HELIX 90 : 10

40

30 β - FORM

20 [m2] ×10–3

18

50 : 50 10

0 COIL –10

–20

–30

190

200

210 220 λ, mμ

230

240

250

Figure 1.13  CD spectrum of poly(lysine) in random coil (left), α‐helical (right, open symbols) and β‐sheet conformation (right, full symbols). Source: Sarkar 1966 [52].

reference spectra come from proteins and might not be applicable to homopolypeptides with protecting groups in organic solvents. For example, the TFA‐ protecting group in Poly(N‐ε‐trifluoroacetyl‐L‐lysine) (PLys(TFA)) includes an amide bond in the side chain whose absorption overlays that of the amides in the backbone. Absorption is also problematic with respect to  ­solvents and salts. Solvents containing amide bonds (DMF, DMAc, NMP (N‐methyl‐2‐pyrrolidone)) as well as buffer salts like chlorides, phosphates, sulfates, carbonates acetates, or TRIS cannot be used due to UV absorption. Thus, quantification of different secondary structures within one polypeptide sample remains challenging and CD spectroscopy may only provide a very rough estimate of existing structures.

Characterization of Polypeptides and Polypeptoides [Θ] = MOLAR ELLIPTICITY

42 38

1 DECA

DI

34

0

30

[Θ] x 10–4

22 OCTA

18 14

–2 –3 –4 –5

10

HEPTA

6 2

DI

–2

TRI

–6 –7 200 210 220 230 240 250 TETRA Wavelength, mμ HEXA HEPTA OCTA

–6 –10

[Θ] x 10–4

26

TRI TETRA PENTA HEXA

–1

NONA

HEXA PENTA

NONA

Figure 1.14  CD spectra of exactly defined N‐carbobenzoxy‐γ‐ethyl‐L‐glutamate oligomers showing a sharp transition from random coil to α‐helix at six repeating units. Source: Goodman et al., 1969 [56]. Reproduced with permission of Goodman.

CD spectroscopy was, for example, used by Goodman et al. on defined N‐carbobenzoxy‐γ‐ethyl‐L‐glutamate oligomers (Đ = 1) to show the sharp transition from random coil to α‐helix at six repeating units (Figure 1.14) [56]. Deming’s group synthesized di(ethylene glycol)‐modified poly(serine) [57]. While these polymers show a random coil conformation in water, they convert to a β‐sheet conformation on exchange of the solvent to methanol (Figure 1.15). Schmidt and coworkers synthesized brushes with poly(lysine) side‐chains [58]. Upon addition of NaClO4, a coil to helix transition of the side‐chains could be observed in the CD spectrum (Figure 1.16). We can conclude, that CD spectroscopy is a very valuable method for gaining an insight into the secondary structures of polypeptides. The quantification of secondary structures by fitting spectra with data from natural proteins is, ­however, problematic, since protecting groups do not allow measurement in water and can influence absorption bands.

19

Polymers for Biomedicine 30 E

[Θ] x 10–3

20

10

D C

0 B

–10

A 180

190

200

210 220 230 Wavelength (nm)

240

250

260

Figure 1.15  CD spectrum of di(ethylene glycol)‐modified poly(serine) showing a coil to sheet transformation with increasing methanol content from A to E. Source: Hwang 2001 [57]. Reproduced with permission of American Chemical Society.

40 30 20 Θ/mdeg

20

10 coil 0 –10 helix

–20 190

200

210

220

230

240

λ/nm

Figure 1.16  CD spectrum of coil to helix transition with increasing NaClO4 concentration in polymer brushes with poly(lysine) side‐chains. Source: Sahl 2012 [58]. Reproduced with permission of American Chemical Society.

Characterization of Polypeptides and Polypeptoides

1.8 ­Mass Spectrometry

Intensity a.u.

MALDI‐TOF (matrix assisted laser desorption/ionization  –  time of flight) spectrometry is usually the mass spectrometry method of choice for polymers. The polymers are first mixed with a matrix material, which is then irradiated by a laser, leading to desorption of the polymer. Next, the polymer is ionized in  the gas phase and analyzed by a time of flight detector. This method is especially mild for large molecules so that usually no fragments and multiply charged ions are obtained [59]. Using MALDI‐TOF, the determination of absolute average molecular weights and dispersities can also be achieved. However, desorption of polymers with high molecular weights is often weaker than for their low molecular weight counterparts, leading to an underestimation of high molecular weight chains and overestimation of low molecular weight chains [60]. While narrowly dispersed samples (Ð 1.20 showing a high molecular weight (HMW) shoulder and peak tailing; and (b) a controlled process with PDI values of  66.7 hr SER-214 Human = t1/2 > 3 days SER-214 Monkey = t1/2 > 3 days

80 60 t1/2 40 20 0

0

24

48

72

Time (h) Note: 37 °C and n = 3 (mean ± SD)

Figure 3.9  Hydrolysis of SER‐214 and release of Rotigotine in various plasma from different species – rat, dog, monkey, and human: Hydrolysis of Rotigotine in plasma from different species – rat, dog, monkey, and human. There is a marked difference in the release rates of Rotigotine in the different plasmas, with rapid release in rat (~11 h) and slow release in monkey and human (>3 days).

monkey and human were almost identical. These release profiles helped identify the most likely candidate drug for the desired target product ­ ­profile  –  which in this instance was a single weekly subcutaneous injectable in a standard insulin syringe that would provide continuous drug delivery of the attached Rotigotine within a relatively narrow therapeutic window (see Figure 3.11 later). SER‐214 as a single subcutaneous injection promoted prompt contraversive rotations and rescued forelimb asymetry in the 6‐hydroxy dopamine (6‐ OHDA)‐lesioned model in rats for approximately 1 week, showing that the release of Rotigotine from the polymer was sustained at pharmacologic levels [38]. When this was extended to the monkey model of Parkinson’s dis­ ease (monkeys will develop characteristic Parkinsonism when fed 1‐methyl‐4‐ phenyl‐1,2,3,6‐tetrahydropyridine (MPTP) in their diet for 3–4 months), a ­single weekly injection of SER‐214 at a human equivalent dose (HED) of around 0.25 mg equivalents Rotigotine per kg rescued Parkinsonism in a manner com­ parable to a twice daily dose of levodopa (L‐DOPA, as a levodopa:carbidopa suspension). This is shown for the Week 2 data in Figure 3.10. The submission of an IND requires that one performs toxicology studies in two species, one rodent (typically rat) and one non‐rodent (we employed cynomolgous macaque monkeys). Ninety‐day chronic dose toxicology studies in rats revealed that SER‐214 was safe and tolerable at doses 40 times the HED, with no

Poly(2-Oxazoline) Polymers in Drug Delivery 200

Vehicle L-DOPA (15 mg/kg)

ON-time (min)

150

SER-214 (1 mg/kg equiv.)

100

50

0

Figure 3.10  Good “On‐Time” in monkeys who receive either placebo polymer, L‐DOPA twice daily or SER‐214 as a single weekly injection: Monkeys were videotaped for 6 hours in an observation cage, and the “on‐time” was determined in a blinded review by a neurologist using a scoring system analogous to that used in humans. Good “On‐Time” is the total cumulative time during the 6‐hour observation period with a bradykinesia score of zero (the monkeys are moving around purposefully without any loss of balance) and without any evidence of dyskinesia (which is an involuntary movement). Vehicle control in this study was a 20 kDa POZ polymer with pendent propionic acid, no attached Rotigotine.

histologic or biochemical abnormalities. Specifically, there was no histologic evidence of vacuole accumulation of POZ in any tissue in the rat. In the monkey, at greater than 20 times the HED, minimal vacuolation of only the renal tubular epithelial cells was noted in the proximal convoluted tubule; no other tissues showed any accumulation and there were no changes in hematology, clinical chemistry, liver function tests, or behavior. The vacuolation resolved following cessation of administration and was of no clinical significance [17]. Figure 3.11 shows the pharmacokinetic profiles for weeks 1, 5, 9, and 12 of the 13‐week study for the high dose cohort of monkeys. In this dose cohort monkeys were administered a single weekly subcutaneous injection of SER‐214 for 13 doses, and during the selected weeks plasma samples were taken for daily measurement of the released Rotigotine. The steady‐state release profile of Rotigotine in the toxicology studies, and the rescue of Parkinsonism in the MPTP‐model, suggests that this profile might also be attainable in humans. If so, then a single weekly subcutaneous injection of SER‐214, administered in a standard insulin syringe, should pro­ vide prompt control of motor fluctuations in patients with Parkinson’s disease who are responsive to a dopamine agonist. In the course of performing these IND‐enabling studies and others (to date, over two dozen studies were submitted to support the IND) we have determined

65

Polymers for Biomedicine 100 Plasma concentration (ng/mL)

66

10

1 week 1 week 5 week 9 week 12

0.1

0.01 0

1

2

3

4

5

6

7

Time (days) Note: Dose 7.0 mg/kg; male and female; n = 10 (mean ± SD)

Figure 3.11  Plasma concentration of Rotigotine following weekly SC injections in normal cynomolgus macaque monkeys; data shown for Weeks 1, 5, 9, and 12, The dose administered SC was 58 mg SER‐214 per kg (SER‐214 contains approximately 12% by weight attached Rotigotine, thus this equates to approximately 7 mg equivalents Rotigotine/kg body weight).

a number of very important properties of this polymer that will be summarized here: (1) the POZ polymer backbone does not undergo detectable chemical change during circulation in experimental animals or upon incubation with homogenates of select tissues, (2) the polymer and polymer conjugate are cleared almost exclusively by renal filtration, and this is a function of polymer size, (3)  whole body autoradiography and micro‐autoradiography of select tissues employing 14C‐labelled SER‐214 demonstrated the conjugate undergoes rapid renal filtration and is not taken up intracellularly by any tissue but the renal epi­ thelium in the kidney, and the minimal vacuolation at high doses resolves upon cessation of dosing, (4) attempts to induce antibodies to POZ have been unsuc­ cessful – POZ appears to be non‐immunogenic (in contrast to many other poly­ mers such as PEG and dextrans), and (5) polymers of POZ can significantly increase the solubility of some drugs (Rotigotine being one such example). POZ appears to be capable of providing steady‐state release pharmacokinet­ ics and pharmacodynamics for Rotigotine. Additional studies are underway at Serina to further characterize other small molecules attached to POZ.

3.6 ­Applications in Targeted Drug Delivery The clinical development of polymer therapeutics for oncology indications has  been less successful than anticipated when applied to small molecule ­oncolytics. Numerous platforms have been employed (PEG, HPMA, PLGA,

Poly(2-Oxazoline) Polymers in Drug Delivery

polydextrans), and candidate drugs have advanced steadily through the various phases of clinical development – but none have achieved approvals. Despite the widespread demonstration of the “EPR” effect (enhanced permeation and retention) for many of these polymer platforms [43,44], EPR by itself is proba­ bly not sufficient to guarantee a polymer therapeutic will succeed in the clinic. While the reasons for failure of these polymer oncolytics are undoubtedly complex, involving not only the platform technologies but the clinical indica­ tions in which they were advanced, we have taken a different approach with POZ‐oncolytics. We are advancing targeted POZ‐therapeutics that employ both small molecule and antibody targeting moieties. 3.6.1  Targeting POZ‐Therapeutics to the High Affinity Folate Receptor Alpha

Cancer is one of the most pressing clinical challenges in healthcare worldwide, and in the next 15 years is predicted to become the number one cause of death in the United States, surpassing heart disease and stroke [45]. There has never been a greater need for new approaches to develop drugs that offer not only a survival benefit – but the potential for an outright cure. We have been working on targeted POZ‐therapeutics for several years and several of our pipeline can­ didates are illustrated in Figure 3.12. Certain types of cancers express a unique receptor on their cell surface that is not expressed on the normal tissue from which that cancer arose (nor in other normal tissues); one such example is the high affinity folate receptor alpha [46]. This receptor has been well‐characterized and is an attractive candi­ date receptor for a POZ‐therapeutic approach – with estimates that as many as

H

N

N

N

N

N

N

N

N

H2O Biological pH Cytotoxic drug

Inert pendent group

Cleavable linker

Stable linker

Targeting group

Figure 3.12  General structure of a targeted POZ‐therapeutic: The polymer backbone is typically a 20 kDa POZ with multiple pendent groups (but as shown in Section 3.3 – this is programmable) to which is attached a cytotoxic linker:drug via click chemistry. The release of the attached drug is controlled by the nature of the linker attaching it to the polymer backbone. The targeting group can be either a small molecule or an antibody, antibody fragment or any other targeting ligand. In the following example the attached drug is irinotecan, a potent topoisomerase I inhibitor.

67

Polymers for Biomedicine

a million new cases of folate‐receptor cancers are diagnosed annually world­ wide [46]. Folic acid has been stably attached to the carboxyl end of the POZ polymer preserving the stereochemistry of the alpha carboxylate  –  which is essential for high affinity binding. We have attached several small candidate molecule oncolytics to folate‐targeted POZ polymers and have shown that these drug candidates can not only target the high affinity receptor (~Ki > kact,b, the products of degrada­ tion will be small fragments with MWs close to that of the original inimer. When truly branched polymers are formed, the products of degradation will be oligomers with lower DP than the starting material, but with MWs higher than that of the inimer.

339

Br R‴ R′ + R″

Y

Br3C

Functional group Br

CBr3

CBr3

Y

Y

R‴

R‴

Y

Br

Branching point R‴

R‴

R‴

Y

Y

Br

R‴

Br

CBr3

Y

Y

Br3C

Br

Br Br2

Br Br

Y

R‴ Y

Y

Br Br

Monomer ATRP

CBr2

Y

Y

Y

Br Br

Br

Br

Br

Br

Br

Br

Y

Br R‴ CBr3

Y Br

Br O = *

O O

O

* O

or *

O

S

S

O

* O

Scheme 11.25  Synthesis of highly branched polymers with hydrolysable (ester) or reductively degradable (disulfide) groups at the branching points by copolymerization of monovynyl monomers and crosslinkers in the presence of CBr4 and their use as macroinitiators for ATRP [177,178]. Source: Woodruff 2015 (178). Reproduced with permission of American Chemical Society.

X

X

X

X

i) kact,b >> kact,s

X kact,s

X

X

ii)

n3

X

X

X n4

iii) kact,s >> kact,b kdeact,b : large

n7

n5

X X n8 Degradation

X n6

X

X

m

m 2 Torr

O

O 8 O

O Ac

N H

O

O

n

N H

Ac

42

Scheme 12.8  Synthesis of 5‐ASA‐based PAE via melt‐condensation polymerization and polymer precursors.

To circumvent solubility issues and ensure that thermal rearrangement and/or polymer crosslinking did not occur, a modified synthetic method was employed to acquire 5‐ASA‐based PAEs [25]. Similar to the preceding method, the amine was still protected using an acyl chloride in pyridine, however, longer aliphatic acyl chlorides were utilized to enhance polymer solubility. The amidosalcylates were then coupled through their phenol functionality using pyridine and diacyl chlorides (sebacoyl and dodecanedioyl) and diacids subsequently polymerized using triphosgene with TEA. The resulting 5‐ASA‐based PAE not only exhibited higher Tg values but also possessed markedly improved solubility in common organic solvents, therefore enhancing PAE processibility. So far, solution polymerizations have been described using triphosgene as a coupling agent; however, milder reaction conditions (i.e., absence of base) may be necessary to avoid unwanted side reactions, or to polymerize heat‐ and acid‐sensitive monomers. Alkoxyacetylenes, specifically (trimethylsilyl) ethoxyacetylene (TMSEA), offer an alternative means of anhydride synthesis without the use of base, enabling simpler purification as the byproduct, ­trimethylsilylacetate, can be removed in vacuo [50]. Utilizing this knowledge, Qian and Mathiowitz investigated TMSEA for PAE synthesis [51]. Diacids (43) were first synthesized as previously described, through a pyridine‐mediated O‐acylation of Mandelic acid, an α‐hydroxy acid

Polymerized Drugs O

O O

HO

O

(CH)8

OH

O

O

Si(CH3)3

n C2H5O

+

43 C2H5O CHSi(CH3)3 H O

O O

n

O

44 C2H5O

O

O

(CH)8 O

O

O O n

O

O

O

O

CH2SiH2(CH3)3 O

n

46 + 45

O n(H3C)3Si

CH2

C

OC2H5

Scheme 12.9  Hypothesized TMSEA‐mediated solution polymerization of Mandelic acid.

with antibacterial properties, and sebacoyl chloride. 43 was then polymerized with TMSEA via an electrophilic addition‐elimination reaction (Scheme 12.9) in which Mandelic diacid (43) is hypothesized to first undergo a concerted electrophilic addition with TMSEA to acquire monoadduct (44). The increased electrophilic reactivity of 44 promotes a subsequent concerted electrophilic addition, acquiring diadduct (45) and promoting polymer chain growth. Following diadduct formation, trimethylsilylacetate is eliminated, completing the anhydride formation to acquire 46. Building upon the work established by Qian and Mathiowitz, Stebbins et  al. employed TMSEA in the synthesis of highly tunable mannitol‐based PAEs (Scheme 12.10a) [52]. To do so the primary alcohols of mannitol were

375

(a)

O OH OH OH

HO OH OH

HO

OH OH

TBDMS-Cl

OTBDMS

imidazole, DMF TBDMSO

R

HO

O

R

R OH

O

O O

O

R

O

O

TEA DCM

O

O

O

R 53

R

O

O

R

52

R

O

R

R

TBAF, AcOH THF

O OH

R

O

R

51

O O

O

R=

O

O

O O

O

HO O

TMSEA DCM O

O

50

O

O

R

OTBDMS O

48

O

O

TBDMSO

EDCl, DMAP, DCM

OH OH

47

R

49

O

R

O

O R

54

Scheme 12.10  (a) Synthesis of poly(tetraibuprofen mannitol succinate) PAE and PAE precursors. (b) Mechanism of TBDMS protection of mannitol primary alcohol through activated N‐t‐butyldimethylsiylimidazole silylating agent. An additional TBDMS protecting group will subsequently protect the second primary alcohol of mannitol via the same mechanism.

Polymerized Drugs

(b)

t-Bu Si Me

Cl t-Bu HN

Me

N

Si

N

Cl

N H

55

R

OH

t-Bu

Me

O

OH

R=

Si

H

56 OH

Me

Me

Me

OH

OH

Cl

H N

O N

R

58

Cl

t-Bu

H Si

H

Me

Me

O R

N N H

57

Scheme 12.10  (Continued)

first selectively protected using t‐butyldimethylsilyl chloride (TBDMS‐Cl) in the presence of imidazole following previously established methods [53]. In this mechanism nucleophilic imidazole first attacks TBDMS‐Cl, displacing chloride, to generate the very reactive N‐t‐butyldimethylsiylimidazole (56) silylating agent (Scheme 12.10b). Once produced, 58 undergoes nucleophilic substitution with mannitol to acquire a more robust silyl ether (48). The sec‑ ondary alcohols of 48 then undergo carbodiimide coupling with ibuprofen in the presence of 1‐ethyl‐3‐(3‐dimethylaminopropyl)carbodiimide (EDCI) and 4‐dimethylaminopyridine (DMAP) to acquire 50. The TBDMS protecting groups of 50 are then selectively deprotected using tetrabutylammonium fluoride (TBAF) and the resulting diol (51) chain‐extended using succinic anhydride in the presence of TEA to acquire tetraibuprofen mannitol suc‑ cinic acid (53). Once purified, 53 was polymerized using TMSEA to acquire relatively homogenous poly(tetraibuprofen mannitol succinate) with a Mw of 22.0 kDa and 1.5 PDI. TMSEA polymerization was chosen, as thermogravi‑ metric analysis (TGA) revealed that activated 53 (acetylated at carboxylic acids) began to decompose prior to its melting point, thus preventing the use of melt‐condensation polymerization.

377

378

Polymers for Biomedicine

12.3 ­Poly(Esters) Whereas polyanhydrides predominantly undergo surface erosion, enabling a near‐zero order release profile, PEs are primarily bulk eroding polymers with ester hydrolytic degradation occurring throughout the polymer system [54]. While bulk eroding polymers can exhibit burst releases in both physical and chemically incorporated systems, they are still some of the most extensively researched biodegradable biomaterials, offering greater stability than polyan‑ hydrides. Some of the most commonly utilized polyesters, such as poly (lactic acid), poly(glycolic acid), and poly(ε‐caprolactone), are synthesized via ring‑opening polymerizations of lactone monomers, often in the presence of ­catalysts. Although this synthetic methodology is effective, bioactive‐based polyesters would require synthesizing complex monomers; thus, polyesterifi‑ cations between a bioactive‐containing monomer and comonomer are more commonplace. Similar to polyanhydride synthesis, melt‐condensation polym‑ erizations offer higher Mw end‐product and simpler scale‐up; consequently, this is the principal method in bioactive‐based polyester synthesis. 12.3.1  Melt‐Condensation Polymerization

Synthesis and applications of SA‐based PAEs have been covered in the previ‑ ous sections. However, PE analogs of SA‐incorporated biodegradable polymers have also been synthesized and thoroughly investigated. Chandorkar et al. reported the synthesis of a novel cross‐linked PEs comprised of SA moieties within the polymeric backbone [55,56]. This SA polyester (SAP) (60) was syn‑ thesized utilizing melt condensation technique in two stages. First, SA diacid (6) was prepared using sebacoyl chloride as a linker molecule, analogously to the previously discussed methods. In the next step, the diacid was activated with acetic anhydride, followed by melt condensation polymerization in the presence of mannitol to acquire a cross‐linked, completely biodegradable PE (SAP, Scheme 12.11). SAP was then cured in a vacuum oven at 130 °C for ~24 h to obtain the cured SAP. Although the non‐cured SAP was soluble in tradi‑ tional organic solvent (e.g., chloroform, THF, DCM, acetone, dioxane, and ethanol) the curing process resulted in negligible solubility owing to extensive cross‐linking. Curing SAP was also found to decrease the SA content, 40–25% (w/w), and the SA release rate under physiological conditions, ~20% in approx‑ imately 5 days to ~3.5% over 4 months, when compared to non‐cured SAP. Polyesters, which are often synthesized from hydroxyacids or combinations of diacids and diols, require different synthetic reagents including alkyl‐inor‑ ganic catalysts such as those containing tin, titanium, or zinc. Among these, stannous octoate (SnOct2) has been predominantly used in the industry due to its low cost, low toxicity, and high efficiency [57,58]. Although zinc‐based cata‑ lysts are thought to be potentially less toxic than tin catalysts, the higher overall

O HO

O

HO OH

O +

O

Cl

2

8

O

Pyridine Cl

O

O

4

O

O O

Ac2O

O

O

THF

OH

8 O

O O

80°C

O O

6

8 O 59

180°C, high vacuum OH OH OH O

O O

O O

O

O

O O

O O

8 O

OH

O

OH

HO O

OH O

OH OH 48

Cross-linked polyester

60

Scheme 12.11  Synthesis of cross‐linked salicylic acid based polyester (SAP).

Mannitol

O

380

Polymers for Biomedicine

efficiency of the latter made them the catalyst of choice for many polymeriza‑ tions utilized in the biomedical and pharmaceutical applications [59]. Controlled and sustained release of NSAIDs, such as ibuprofen and naproxen, from a chemically incorporated polymeric matrix could be advantageous espe‑ cially considering their short half‐lives, severe gastrointestinal side effects at high doses, and low drug loading profiles within physically incorporated poly‑ mers [60]. To address these issues, novel biodegradable PEs comprised exclu‑ sively of biocompatible components (e.g., tartaric acid, 1,8‐octanediol) and ibuprofen or naproxen as a pendant group have been synthesized, containing 65–67% NSAID loading [8]. Scheme 12.12 illustrates the synthesis of these PEs in which tartaric acid was used as a backbone. Ibuprofen or naproxen, both of which contain a carboxylic acid group, was first coupled to each hydroxyl group of dibenzyl‐L‐tartrate via carbodiimide mediated esterification reaction. Benzyl protecting groups on the tartaric acid derivative were then selectively cleaved using Pd‐catalyzed hydrogenation, yielding NSAID‐containing tartaric acid diacid. This debenzylation method provided an easier work‐up and purification steps, enabling the higher yield synthesis of the desired products. Using stan‑ nous 2‐ethylhexanoate as the catalyst, 1,8‐octanediol and the synthesized diacid were subjected to melt‐condensation conditions. After the reactants melted, a monophasic mixture was formed which increased in viscosity as reaction pro‑ gressed. The chemical structures and purity of the synthesized compounds were confirmed by 1H, 13C NMR, and FTIR spectroscopies. Naproxen‐containing polyesters, having a more rigid structure, exhibited higher glass transition tem‑ peratures and slower bioactive release rates, when compared to the ibuprofen PEs. Cytocompatibility studies deemed the polymers nontoxic toward fibro‑ blasts and 1H NMR spectroscopy confirmed that the structure of the released NSAID was retained, suggesting preserved bioactivity. 12.3.2  Solution Polymerization

Whereas various catalysts and methods are commonly utilized in polyester melt‐condensation polymerization, few alternatives exist for solution polym‑ erization. Triphosgene, while commonly utilized in solution polyanhydride synthesis, is unsuitable in polyester synthesis owing to its inability to selectively couple hydroxyl and carboxylic acid functionalities. Thus, reaction conditions must be employed to specifically conjugate the aforementioned groups. One such means of polyesterification of bioactives via solution polymeriza‑ tion is through carbodiimide coupling, as demonstrated by Wattamwar et al. [61]. Here, trolox (67), an analogue of Vitamin E that is commonly ­utilized as  an antioxidant standard, was polymerized in the presence of N,N’‑dicyclohexylcarbodiimide (DCC) and DMAP to acquire oligomeric poly(trolox ester) (68) (Scheme  12.13). In this mechanism, the carboxylic acid of trolox first reacts with DCC, generating an O‐acyl urea (69), which

R

R

O OH O OH

R

+

BnO

O

OBn O

61

EDCI, DMAP

OH

DCM, r.t.

O

O

O

BnO O

O

62

O

Pd/C, H2

OBn

O

HO

DCM, r.t.

OH O

O

R

O

+

HO

O

R 64

63 R Ibuprofen

R=

O O

Stannous hexanoate 130°C

O

O

O O Naproxen

O R

Scheme 12.12  Synthesis of NSAID containing biodegradable polyesters.

O 6

O

n

2 Torr 100 rpm

OH 6 65

382

Polymers for Biomedicine

H

O

O

+

O

OH

R N C N R Carbodiimide

DMAP

O

O

O

O

n

Oligo (trolox) 68

Trolox 67

R N C N R Carbodiimide H

O

O O

O

O O

O

Trolox 67 n

DMAP

OH R N O C HN O R

O

O O

O

n

O-acyl urea 69

Poly(trolox) 70

O

N R N C OO HN R

O

O O

O

n

N-acyl urea 71

Scheme 12.13  Poly/oligo(trolox) synthesis via carbodiimide coupling and competing oxygen → nitrogen migration that can effectively hinder polyesterification.

subsequently reacts with DMAP forming an activated ester. The activated ester then readily reacts with the hydroxyl group of trolox, facilitating poly‑ ester formation (70). Following O‐acyl urea formation, an oxygen → nitrogen migration, acquiring an undesired side‐product N‐acyl urea (71), competes with ester activation via DMAP, capping the polymerization. To suppress the formation of N‐acyl ureas, p‐toluenesulfonic acid (PTSA) was added as acid catalysts have been shown to hinder this oxygen → nitrogen by lowering the reaction pH [62]. This also was found to increase polymer Mw, presumably due to less chain‐termination from N‐acyl urea formation resulting in short‐ chain oligomers. While trolox contains both hydroxyl and carboxylic acid functional groups, enabling self‐polymerization, many compounds lack this feature. As such, alternate methods are often utilized in the solution polymerization of bioactive containing polyesters. One such method applicable to dihydroxy‐bioactives,

Polymerized Drugs

(a) HO

OH

O

OH

H

72

O OH

pTSA

O

O

HO

Benzene

OH

O

Vanillin 73

74

(b) O Cl O 75

Cl +

HO

O

O

O OH DCM

O

O

O 74

O PVO

H+

O H

O

O

O

O

HO OH +

n 76 H2O2

OH OH

+ 2CO2

Scheme 12.14  Synthesis, characterization and hydrolysis of PVO (a) Synthesis of acetal‐ protected, diol‐containing vanillin prodrug, (b) Synthesis and subsequent degradation of PVO.

explored by Lee et al., utilizes a diol with a diacyl chloride in the presence a proton acceptor [63–65]. A unique feature of PEs synthesized by Lee is the presence of aromatic ­peroxalate esters, which have been shown to readily react with hydrogen peroxide (H2O2) [65], thus functioning as a free radical scavenger and imparting bioactivity on the polymer backbone. Antioxidants, vanillin and p‐hydroxybenzyl alcohol, were uti‑ lized in the synthesis of the aforementioned polyesters. As vanillin possesses a single phenol, it was first reacted with 2‐(hydroxymethyl)‐2‑methylpropane‐1,3‐ diol in the presence of PTSA to acquire an acetal‐protected­, diol‐containing vanil‑ lin prodrug (74, Scheme 12.14). The vanillin prodrug was subsequently dissolved in DCM with pyridine, to which oxalyl chloride was added drop‐wise at reduced temperatures to acquire poly(vanillin oxalate) PVO, an antioxidant‐based poly‑ mer possessing inherent bioactivity.

12.4 ­Green Chemistry Green chemistry aims to mitigate environmental, economic and health c­ oncerns associated with traditional chemistry through minimizing the use and genera‑ tion of hazardous waste and maximize efficiency en‐route to the final prod‑ uct [66,67]. Biodegradable polymer development is continuously increasing, and

383

384

Polymers for Biomedicine

resulting in high demand for various biomaterial applications. The principles of green chemistry can be implemented to biodegradable polymers to reduce the risks to human health during the process [67]. Non‐hazardous reactant mono‑ mers, safe solvents, and bio‐renewable enzymatic catalysts can be utilized. 12.4.1  One‐Pot Synthesis

In a continuing effort to increase efficiency, SA‐adipic PAEs were synthesized using a green chemistry approach. In Section 12.2.1, the optimized SA‐based diacid synthesis was discussed. In this approach, SA was directly coupled to a diacyl chloride through the acyl‐pyridinium intermediate to acquire SA diacid. In addition to reducing the number of synthesis steps, the overall yield was increased from 50 to 97%. SA‐based PAE synthesis was further improved by eliminating solvent and reducing raw material usage via a one‐pot synthetic method. The traditional and one‐pot approaches are shown in Figure 12.4. In the traditional method, SA‐adipic PAEs were synthesized in three steps; prepa‑ ration of polymer precursor, activation of polymer precursor and melt conden‑ sation polymerization. While the one‐pot method utilizes the same synthetic steps, isolation of the reaction intermediates is not performed. This method drastically decreased the overall production time of SA‐adipic PAEs, raw mate‑ rial and solvent usage without impacting polymer properties [68]. The one‐pot method can lower the manufacturing cost via short reaction time and high yield of the total process. Additionally, it minimizes the use of hazardous com‑ ponents which is beneficial for the environment as well as for engineers who are working in scale‐up plants. 12.4.2  Lipase‐Catalyzed Synthesis

Over 3000 enzymes have been identified and applied as catalysts over the past 30 years [69]. Enzyme‐catalyzed reactions have received much attention due to their high‐efficiency, non‐toxicity, environmental friendliness, and recyclabil‑ ity [70,71]. Among enzymes, lipase is the most commonly used biocatalyst, frequently employed in esterification, Michael addition, and transesterification reactions [70]. In recent years, lipase has been utilized in PE syntheses, com‑ monly with diol/diacid monomers [72,73]. Additionally, it has been applied to the synthesis of bioactive‐containing polymers by Wang et al., developing PE prodrugs of ketoprofen [74]. In Section 12.3.1, PEs comprised of tartaric acid, 1,8‐octancediol, and ibupro‑ fen were synthesized utilizing Tin (II) 2‐ethylhexanoate. As catalyst toxicity is a prevalent issue in biomaterials, Stebbins et al. have explored alternate, greener methods to synthesize ibuprofen containing PEs [9]. Using Novozym 435 (N435, lipase from Candida antarctica immobilized on acrylic resin), which is com‑ mon in PEs syntheses [75,76], PEs containing malic acid, ibuprofen, and ali‑ phatic diols were synthesized. In summary, this work focused on the green

(a) HO

(b) HO

O OH

Cl

HO

Cl

Pyridine

4

+

O

O O

THF, R.T

O

O

O

OH

Cl

OH

O 4

O

O

Cl 4

+ O

O

O

O

O Excess: r.t.

1. Pyridine 2. Acetic anhydride, 75°C 3. >2 Torr, 175°C

O

O O

O

O

O

180°C 2 mmHg

4

O

O

O

O

O O

O

O

O

O

O O 4

4

O

O

n

Figure 12.4  Comparison of traditional (a) and (b) one‐pot approach.

O

O O

O n

O O

O HO

OH O

BnBr, Na2CO3 DMF, 40°C

OH 77

O O

O O

OH

O +

OH

O

R

O n

N435 Ph2O 80–95 °C 2 Torr

O

HO

R

OH

HO

82

OH O

O O

83 81 a

R=

3

b

5

c

8

80

Pd/C, H2

O

O

O

O

2 mol% DMAP neat, 50°C

79

78 O

O O

O

Pivalic anhydride

Scheme 12.15  Synthesis of poly(ibuprofen‐L‐malate) and polymer precursors.

CPME

Polymerized Drugs

principles such as use of catalyst and renewable resources, solvent elimination, replacement of environmentally hazardous chemicals, and biodegradable prod‑ ucts to develop more environmentally friendly synthesis processes. The synthesis of poly(ibuprofen‐L‐malate) is shown in Scheme  12.15. Following previously published methods by Guo et al. [77], the carboxylic acids of malic acid were protected with benzyl bromide using sodium carbonate (Na2CO3) in anhydrous DMF to prevent unwanted side reactions and synthe‑ size 78. Subsequently, a solvent‐free esterification method using a catalytic amount of DMAP and pivalic anhydride, established by Sakakura et al. [78], was utilized to couple ibuprofen to the free alcohol of dibenzyl‐L‐malate. Following successful isolation 80 was deprotected via hydrogenolysis. In place of traditional solvents such as dichloromethane or tetrahydrofuran, less toxic and less volatile cyclopentylmethyl ether was chosen [79]. Ibuprofen‐L‑malic acid (81) was then polymerized with aliphatic diols (82) via N345‐catalyzed solution polymerization. Increasing the length of the aliphatic chain (Compound 83) displayed a slightly higher Mw and a decreased Tg of the polymer. Furthermore, the release profile of the polymers was evaluated; sus‑ tained, controlled ibuprofen release was achieved through the 30‐day study. The ibuprofen release rate was decreased with increasing aliphatic chain length of the diol; after 30 days, 42, 58, and 82% of total ibuprofen was released from octylene, pentylene, and propylene polymers, respectively.

12.5 ­Conclusion In this book chapter, novel, biodegradable, and biocompatible PAEs and PEs are reviewed. These systems have been developed to deliver bioactive materi‑ als, such as NSAIDs, antiseptics, opiods, antioxidants, and antimicrobials in a controlled and sustained manner. PAEs and PEs have been synthesized using different melt condensation and solution polymerization methodologies. Furthermore, polymer characterization techniques and release mechanism of these unique compounds are highlighted. Additionally, green polymer chemis‑ try approaches are discussed including one pot syntheses and lipase‐catalyzed polymerizations. As polymer‐based bioactive delivery systems are growing in numbers it is significant to investigate alternate synthetic methods for increased efficiency and reduced environmental impact.

­References 1 Pasut, G. Veronese, F.M. Prog. Polym. Sci. 32 (2007) 933–961. Duncan, R. Vicent, M.J. Adv. Drug Delivery Rev. 65 (2013) 60–70. 2 Allen, T.M., Cullis, P.R. Drug delivery systems; entering the mainstream, 3

Science 303 (2004) 1818–1822.

387

388

Polymers for Biomedicine

4 Wang, Y.S., Youngster, S., Grace, M., Bausch, J., Bordens, R., Wyss, D.F.

Adv. Drug Delivery Rev. 54 (2002) 547–570.

5 Levy, Y., Hershfield, M.S., Fernandez‐Mejia, C., Polmar, S.H., Scrudiery, D.,

Berger, M., Sorensen, R.U. J. Pediatr 113 (1988) 312–317.

6 Liu, S., Maheshwari, R., Kiick, K.L. Macromolecules 42 (2009) 3–13. 7 Zhang, Y., Chan, H.F., Leong, K.W. Adv. Drug Delivery Rev. 65 (2013)

104–120.

8 Rosario‐Melendez, R., Yu, W., Uhrich, K.E. Biomacromolecules 14 (2013)

3542–3548.

9 Stebbins, N.D. Yu, W., Uhrich, K.E. Macromol. Biosci. 15 (2015)1115–1124. 10 Goepferich, A., Tessmar, J. Adv. Drug Delivery Rev. 54 (2002) 911–931. 11 Goepferich, A. Biomaterials 17 (1996) 103–114. 12 Whitaker‐Brothers, K., Uhrich, K.E. J. Biomed. Mater. Res., Part A 76A (2006)

470–479.

13 Schmeltzer, R.C., Schmalenberg, K.E., Uhrich, K.E. Biomacromolecules 6

(2005) 359–367.

14 Anastasiou, T.J. Uhrich, K.E. Journal of Polymer Science: Part A: Polymer

Chemistry 41 (2003) 3667–3679.

15 Domb, A.J. Haffer, A.T.S. Biodegradable polymer blends for drug delivery,

US patent (1992), WO 9213567 A1 19920820.

16 Rosen, H.B., Chang, J., Wnek, G.E., Linhardt, R.J., Langer, R. Biomaterials 4

(1983) 131–133.

17 Dang, W., Daviau, T., Ying P., Zhao, Y., Nowotnik, D., Clow, C.S. et al. J Control 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32

Release 42 (1996) 83–92. Leong, K.W., Langer, R. Macromolecules 20 (1987) 705–712. Domb, A.J., Langer, R. J Polym Sci, Part A: Polymer Chem 25 (1987) 3373–3386. Erdmann, L. Uhrich, K.E. Biomaterials 21 (2000) 1941–1946. Mao, P., Liu, Z., Xie, M., Jiang, R., Liu, W., Wang, X., Meng, S., She, G. Mini‐Reviews in Medicinal Chemistry 14 (2014) 56–63. Rowland, M., Riegelman, S. J. Pharm. Sci. 57 (1968) 1313–1319. Schmeltzer R.C., Anastasiou, T.J., Uhrich K.E. Polym. Bull. 49 (2003) 441–448. Anastasiou T.J., Uhrich, K.E. J. Polym Sci: Part A: Polym Chem 41 (2003) 3667–3679. Kim, Y. Uhrich, K.E. J. Polym. Sci.: Part A: Polym Chem 48 (2010) 6003–6008. Carbone, A., Song, M., Uhrich, K.E. Biomacromolecules 9 (2008) 1604–1612. Schmeltzer, R.C., Uhrich, K.E., Polym. Bull. 57 (2006) 281–291. Campo, C.J., Anastasiou, T., Uhrich, K.E. Polym. Bull. 42 (1999) 61–68. Anastasiou, T. Uhrich K.E. Macromolecules 33 (2000) 6217–6221. Upton, R.N., Semple, T.J., Macintyre, P.E. Clin. Pharmacokinet. 33 (1997) 225–244. Mao, J., Price, D.D. Mayer, D.J. Pain 62 (1995) 259–274. Rosario‐Melendez, R., Harris, C.L., Delgado‐Rivera, R., Yu, L., Uhrich, K.E. J. Controlled Release 162 (2012) 538–544.

Polymerized Drugs

33 Domb, A.J., Ron, E., Langer, R. Macromolecules 21 (1988) 1925–1929. 34 Eckert, H., Forster, B., Triphosgene, a Crystalline Phosgene Substitute, Angew.

Chem. Int. Ed. 26 (1987) 894–895.

35 Valko, M., Leibfritz, M., Moncol, J., Cronin, M.T.D., Mazur, M., Telser, J. Int J

Biochem Cell Biol 39 (2007) 44–84.

36 Wang, Q., Gao, X., Gong, H., Lin, X., Saint‐Leger, D., Senee, J. J.Cosmetic

Science 62 (2011) 483–503.

37 Ouimet, M.A., Griffin, J., Carbone‐Howell, A.L., Wu, W.H., Stebbins, N.D., Di,

R., Uhrich, K.E. Biomacromolecules 14 (2013) 854–861.

38 Ouimet, M.A. Stebbins, N.D., Uhrich, K.E. Macromol Rapid Commun 34

(2013) 1231–1236.

39 Hu, W.X., Xia, C.N.,Wang, G.H., Zhou, W. J. Chem. Res. 9 (2006) 586–588. 40 Schmeltzer, R.C., Johnson, M., Griffin, J., Uhrich, K.E., J. Biomater. Sci., Polym.

Ed. 19 (2008) 1295–1306.

41 Ouimet, M. A., Faig, J. J., Yu, W., Uhrich, K. E., Biomacromolecules, 16 (2015),

2911–2919.

42 Prudencio, A., Faig, J.J., Song, M.J., Uhrich, K.E. Macromolec Biosci. 16 (2016)

214–222.

43 Fan, J., Jiang, H.L. Chen, D. Zhu, K.J. J. Appl. Polym. Sci. 100 (2006)

1214–1221.

44 Brewer, M.S. Food Science and Food Safety 10 (2011) 221–247. 45 Burt, S. Int J Food Microbiol 94 (2004) 223–253. 46 Carbone‐Howell, A.L., Stebbins, N.D., Uhrich, K.E. Biomacromolecules 15

(2014) 1889–1895.

47 Prudencio, A. Carbone, A.L., Griffin, J. Uhrich, K.E., Macromol Rapid

Commun 30 (2009) 1101–1108.

48 Cannarsi, M., Baiano, A., Sinigaglia, M., Ferrara, L., Baculo, R., Del Nobile,

M.A. Int. J. Food Science & Technology 43 (2008) 573–578.

49 Podolsky, D.K., New England Journal of Medicine 347 (2002) 417–129. 50 Kita, Y., Akai, S., Ajimura, N., Yoshigi, M.; Tsugoshi, T.; Yasuda, H.; Tamura, Y.,

J. Org. Chem. 51 (1986) 4150–4158.

51 Qian, H., Mathiowitz, E. Macromolecules 40 (2007) 7748–7751. 52 Stebbins, N.D., AYu, W., Uhrich, K. E. Biomacromolecules 16 (2015)

3632–3639.

53 Nishiyama, K, Yamada, A., Takahashi, M., Takeuchi, T., Kasai, K., Kobayashi,

S., et al. Chem. Pharm. Bull. 58 (2010) 495–500.

54 Uhrich, K.E., Cannizzaro, S.M., Langer, R., Shakesheff, K.M. Chemical

Reviews 99 (1999) 3181–3198.

55 Chandorkar, Y., Bhagat, R.K., Madras, G., Basu, B. Biomacromolecules 15

(2014) 863–875.

56 Chandorkar, Y., Bhaskar, N., Madras, G., Basu, B. Biomacromolecules 16

(2015) 636–649.

57 Storey, R., Sherman, J.W. Macromolecules 35 (2002) 1504–1512.

389

390

Polymers for Biomedicine

58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79

Pang, K., Kotek, R., Tonelli, A. Prog. Polym. Sci. 31 (2006) 1009–1037. Schwach, G., Coudane, J., Engel, R, Vert, M. Biomaterials 23 (2002) 993–1002. Brooks, P.M., Day, R.O. New England J Med 324 (1991) 1716–1725. Wattamwar, P.P. Mo, Y., Wan, R., Palli, R., Zhang, Q., Dziubla, T.D. Advanced Functional Materials 20 (2010) 147–154. Moore, J.S., Stupp, S. I., Macromolecules 23 (1990) 65–70. Kwon, J. Kim, J., Park, S., Khang, G., Kang, P.M., Lee, D. Biomacromolecules 14 (2013) 1618–1626. Park, H., Kim, S., Kim, S., Song, Y., Seung, K., Hong, D., et al., Biomacromolecules 11 (2010) 2103–2108. Kim, S., Park, H., Song, Y., Hong, D., Kim, O., Jo, E., et al. Biomaterials 32 (2011) 3021–3029. Anastas, P.T., Warner, J.C. Chemistry: Theory and Preactie, Oxford University Press, New York, 1998. Anastas, P.T., Eghbali, N. Chem. Soc. Rev. 39 (2010) 301–312. Faig, J.J., Smith, K., Uhrich, K.E. 248th ACS Meeting&Exposition, American Chemical Society, San Francisco, CA, 2014, pp. POLY‐489. Faber, K.M. Biotransformations in Organic Chemistry, New York: Springer‐ Verlag (2004). Gross, R.A., Kalra, B. Chem. Rev. 101 (2001) 2097–2124. Puskas, J.E., Seo, K.S., Sen, M. Y. European Polymer Journal 47 (2011) 524–534. Juais, D., Naves, A.F., Li, C., Gross, R.A., Catalani, L.H. Macromolecules 43 (2010) 10315–10319. Kato, M., Toshima, K., Matsumura, S., Biomacromolecules 10 (2009) 366–373. Wang, H.‐Y., Zhang, W.‐W., Wang, N., Li, C., Li, K., Yu, X.‐Q. Biomacromolecules 11 (2010) 3290–3293. Mahapatro, A., Kalra, B., Kumar, A. Gross, R.A. Biomacromolecules 4 (2003) 544–551. Azim, H., Dekhterman, A., Jiang, Z., Gross, R.A. Biomacromolecules 7 (2006) 3093–3097. Guo, W., Li, J., Fan, N., Wu, W., Zhou, P., Xia, C. Synth. Commun. 35 (2005) 145–153. Sakakura, A., Kawajiri, K., Ohkubo, T., Kosugi, Y., Ishihara, K. J. Am. Chem. Soc. 129 (2007) 14775–14779. Watanabe, K., Yamagiwa, N., Torisawa, Y. Organic Process Research & Development 11 (2007) 251–258.

391

13 Structural Design and Synthesis of Polymer Prodrugs Petr Chytil, Libor Kostka, and Tomáš Etrych Institute of Macromolecular Chemistry, Academy of Sciences of the Czech Republic, Prague, Czech Republic

13.1 ­Introduction Targeted therapy is a novel, highly innovative field of modern medicine, ­especially oncology, which has contributed immensely to improved outcomes for patients with diverse types of cancers during the last decade. Improving the therapeutic index of drugs is a major impulse for innovation in many thera­ peutic areas. Conventional drug formulations, such as tablets, capsules, pills, creams, liquids, aerosols, and injections involving a number of non‐specific administrations of a drug often have contraindications like inappropriate drug concentrations and excessive drug presence in healthy tissues. Frequent appli­ cations of high drug doses are often required because of the drugs’ unfavorable properties, such as short biological half‐life, excessive, or insufficient water‐ solubility, which restrict drug bioavailability. To overcome these drawbacks two main approaches have been investigated: (i) the development of new ­therapeutic substances with more specific effects and (ii) the improvement of conventional drugs by the use of various nano‐carrier systems. Apart from the development of novel anti‐cancer agents, new formulations of “classic” cytostatic drugs, so called drug delivery systems (DDS), including their ­encapsulation into nanoparticles, liposomes, or micelles, or covalent binding to polymer carriers appeared to be a very promising strategy. In contrast to  novel agents, the mechanism of the anti‐tumor activity of the drugs in DDS is not fundamentally different from the parent cytostatics, but is based on improved pharmacokinetic and pharmacodynamic properties, and signifi­ cantly lower toxic profiles. Recently, enormous efforts have been placed on the development of ­nanosized materials, such as liposomes, micelles, or nanoparticles, with the Polymers for Biomedicine: Synthesis, Characterization, and Applications, First Edition. Edited by Carmen Scholz. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

392

Polymers for Biomedicine

potential to serve as efficient diagnostic and/or therapeutic tool for severe ­diseases including cancer, and infectious and neurodegenerative disorders. Firstly, drug encapsulation during the self‐assembly of amphiphilic or hydro­ phobic copolymers in aqueous solutions is used for the drug incorporation. However, DDS with incorporated drugs face strong limitations, which may obstruct their further transfer to clinical tests and subsequently to the market: (i) the “burst effect (release),” during which a large fraction of adsorbed drug is spontaneously released after administration, can lead to severe toxicity in vivo; (ii) the encapsulation of poorly soluble drugs with a tendency to crystallization often requires the use of additional organic co‐solvents during nanomedicine preparation; (iii) the poor and insufficient drug loading, generally only a few percent, usually requires a relatively high concentration of polymer carrier to obtain a noticeable therapeutic effect, which can itself be toxic and damaging to patients. The mentioned drawbacks can be avoided using the prodrug approach, in which the drug is covalently linked to an appropriate polymer carrier (see Kopeček’s model, Figure  13.1). The inactive prodrug is further metabolized in vivo into an active drug or its metabolite. In the case of polymer prodrugs, this strategy improves drug solubility, prolongs circulation in blood, enhances pharmacokinetics and pharmacodynamics, and reduces adverse side effects. Especially the last feature is of major importance in many chemotherapy treat­ ment settings, as the therapy outcome can be impacted by potential damage to the immune system. The materials used for the construction of drug carriers must be nontoxic, biocompatible, non‐immunogenic and must be cleared from the body to prevent long‐term accumulation of the carrier in the organism. Over the last few decades, many polymer prodrug carrier systems were pro­ posed and tested as DDS. Many of these systems are based on water‐soluble polymer carriers bearing low‐molecular‐weight drugs; for example, cytostatic agents covalently bound by biodegradable spacers, which enable a controlled release of the active drug in tumor tissues or cells. Moreover, the new prodrug Drug Biodegradable spacer

Polymer carrier Targeting moiety

Figure 13.1  Schematic description of Kopeček‘s model of polymer prodrug based on the simplified Helmut Ringsdorf’s model. Source: Drobnik 1976 [1]. Reproduced with permission of John Wiley & Sons.

Structural Design and Synthesis of Polymer Prodrugs

formulations enable solubilization of otherwise water‐insoluble or poorly ­soluble drugs. In general, polymer‐prodrug systems should meet three basic criteria: 1) protect the drug from degradation during administration; 2) deliver the drug to the target tissue or cells; 3) enable controlled drug release due to exogenous (i.e., temperature) or endogenous (i.e., pH, enzymes) stimuli. In addition, the combination of therapeutic functionality with an imaging capability (so‐called theranostics) can facilitate visualization of targeted and treated tissue, and thus, enables the observation of the treatment progress. A relevant exemplifying of the prodrug strategy applied to polymers can be found in the field of cancer therapy, which certainly represents the most stud­ ied disease in drug delivery. Various anticancer agent–polymer conjugates have been designed for overcoming disadvantages of classical chemothera­ peutics. In this chapter, the design, structure, synthetic routes, and physico­ chemical and biological characterization of polymer prodrug systems is presented with special attention to the controlled drug release and elimina­ tion of ­polymer carriers used in prodrug therapy. This chapter will focus mainly on currently used water‐soluble polymer carriers, their synthesis, structure, and ability to deliver drugs to the final target.

13.2  Structural Aspects of Polymer Carriers While polymers of natural origin, such as polysaccharides and proteins, were initially used as carriers for polymer prodrugs [2,3] progress in polymer sci­ ence led to the synthesis of novel polymer carriers with tailor‐made structures and appropriate functional groups for the attachment of drugs or other biologi­ cally active compounds. Since it is equally important to deliver a drug to a target tissue and to eliminate the polymer carrier from the organism thereafter, the degradability of the polymer carrier is highly important. Three sub‐groups can be distinguished: polymers with a biodegradable backbone, polymers with biodegradable linkages between fundamental polymer blocks, and polymers that are not biodegradable. Unfortunately, many studies focus only on the delivery role of carriers and omit the necessity of the carrier elimination, which is discussed in detail in the last part of the chapter. The impact of the primary/secondary structure of a polymer carrier on drug delivery has been illustrated for poly(amino acid)s. Poly(α‐aminoacids); for example, poly(L‐Lysine), poly(L‐aspartic acid), poly(L‐glutamic acid) (PGA) and their derivatives are prepared by anionic ring‐opening polymerization of  respective N‐carboxyanhydrides (NCA), and they are enzymatically ­degradable [4,5]. For example, a conjugate of PGA with the anticancer drug

393

394

Polymers for Biomedicine

paclitaxel (PTX) is under clinical investigation (Phase II/III) [6]. In this case, PTX is bound by an ester bond, which is susceptible to lysosomal protease degradation leading to the release of the diglutamate derivative of PTX and subsequently also free PTX. Hudecz used the cis‐aconityl derivative of daunomycin (Dau) for its attach­ ment via an acid‐sensitive linkage to various structurally related synthetic branched polypeptides with a poly(L‐lysine) backbone [7]. The authors studied the relationship between the branched peptide carrier structure, biodistribu­ tion and cytotoxicity and found that changes in the primary/secondary struc­ ture of the carrier can alter the biodistribution profile and in vitro cytotoxic activity of the Dau–polymer conjugate. Detailed evaluation of biological prop­ erties of the–polymer‐Dau showed a decrease in cytotoxicity and an increase in therapeutic activity in the treatment of mice bearing L1210 lymphocytic leukemia [8]. In general, charged polymers, such as poly(aminoacid)s, have some disad­ vantages; undesired interaction with body compartments and accumulation in specific organs can occur, for example, liver and kidney. However, these limita­ tions can be exploited in some cases for the specific targeting of tissues. For example, the conjugate with PTX mentioned previously, which is tested in clinical trials, is intended especially for the treatment of ovarian cancer, because of the enhanced accumulation of the conjugate in the ovaries that is driven by the structure of the polymer carrier. Another structural aspect is biologically rooted and based on differences in healthy and diseased tissue. “Passive” targeting into solid tumors is one way to improve the anti‐tumor activity of polymer prodrug conjugates and it is ena­ bled by the enhanced permeability and retention (EPR) effect, which is caused by structural differences in healthy and malignant tissue [9]. To actually exploit the EPR effect, polymer carriers must have a prolonged blood circulation in order to achieve increased tumor uptake. Having a high molecular weight (HMW) prevents the fast elimination of the polymer carrier and its drug cargo from the organism by renal filtration. Seymour showed that the extent of pas­ sive polymer carrier accumulation in solid tumors depends primarily on its molecular weight [10]. However, this trend reverses at very high molecular weights due to the slower extravasation of these polymer systems as observed for example for star‐like poly(N‐(2‐hydroxypropyl)methacrylamide) (PHPMA) polymer with a molecular weight of about Mw = 1,000,000 g/mol [11]. Ideally, polymer drug carriers should be eliminated from the body after the drugs are released and delivered. While the molecular weight of polymer ­carriers with a biodegradable backbone is not limited, only non‐degradable polymers of molecular weights under a certain limit; approximately Mw = 50,000 g/mol for vinylic copolymers, are small enough to undergo renal filtration. Hence, the HMW polymer drug carriers exhibiting a significant EPR effect should contain biodegradable linkages between single non‐degradable

Structural Design and Synthesis of Polymer Prodrugs + Bifunctional reagent

Branched copolymer

Multivalent copolymer + Dendrimer

Semitelechelic copolymer

Grafted copolymer

Star-like copolymer + or

Telechelic or heterotelechelic (co)polymers

Multiblocked copolymer

Figure 13.2  Schematic description of synthesis of HMW polymer carriers suitable for passively targeted prodrug systems containing biodegradable linkages facilitating their elimination from the organism.

polymer chains with Mw below the limit of renal filtration in order to increase passive targeting and to allow the subsequent elimination of the carrier frag­ ments from the body (Figure 13.2). Alternatively, HMW supramolecular struc­ tures such as micelles formed by the self‐assembly of amphiphilic copolymers with molecular weights below the limit of renal threshold were proposed. Here, polymer carriers are removed as unimers by glomerular filtration. Water‐soluble polymer carriers based on N‐(2‐hydroxypropyl)methacryla­ mide (HPMA) copolymers have been profoundly studied because of their excellent in vitro and in vivo properties. The first synthesis and polymerization of HPMA was published in the 1970s [12]. This important polymer has since turned into a platform polymer that has been copolymerized to contain func­ tional groups suitable for the attachment of biologically active agents (drugs, enzymes, hormones) [13] and has also been used for the synthesis of HMW polymers.

395

396

Polymers for Biomedicine

The first proposed HMW structures were branched copolymers containing tetrapeptide Gly‐Phe‐Leu‐Gly (GFLG) spacers between linear PHPMA seg­ ments designed for enzymolysis by lysosomal enzymes in tumor cells [14,15]. They were prepared by copolymerization with the addition of the crosslink­ ing  agent N2,N5‐bis(N‐methacryloylglycyl‐DL‐phenylalanylleucyl) ornithine resulting in the formation of copolymers with Mw from 160,000 to 1,200,000 g/mol [14]. The anticancer drug doxorubicin (Dox) was attached to the enzy­ matically degradable GFLG spacer. Also, branched polymer carriers bearing hydrazide groups, which can bind to Dox, were formed by post‐polymerization crosslinking of the PHPMA bearing 4‐nitrophenyl glycyl‐DL‐phenylalanyl­ leucylglycinate spacers [15]. Prolonged blood circulation and tumor accumula­ tion of Dox was observed in vivo and enzymatic degradation of the HMW structures was verified in vitro by incubation in a buffer in the presence of cathepsin B. However, wide dispersities (Ð = 3–7) and poor reproducibility of the syntheses limited the further development of branched carriers. More advanced graft polymer carriers were prepared by the grafting of semitelechelic PHPMA onto multivalent PHPMA via spacers susceptible ­ to  enzymatic (GFLG oligopeptide) or reductive (S‐S bond) degradation (Figure 13.3) [16]. Semitelechelic PHPMA containing N‐hydroxysuccinimide (NHS) or thiazolidine‐2‐thione (TT) were grafted onto the copolymers bear­ ing GFLG spacers terminated by hydrazide or primary aminogroups distrib­ uted randomly along the polymer chain. Alternatively, semitelechelic PHPMA containing pyridyldisulfanyl end groups was attached onto copolymers with thiol groups formed by thiolation of polymer‐bound amine groups by 2‐imi­ nothiolane. Prolonged blood circulation and enhanced tumor uptake of the polymer conjugates with Dox bound by a pH‐sensitive hydrazone bond resulted in excellent anti‐tumor activity. The molecular weight of these copolymers was  usually about 90,000–120,000 g/mol, but again, dispersities were very high, Ð = 2.5–3. The most studied non‐degradable water soluble polymer used for the syn­ thesis of polymer‐drug conjugates is poly(ethylene glycol) (PEG) [17] and it has also been employed in generating HMW polymer carriers. Multi‐block PEG copolymers, Mw = 50,000 g/mol, were formed by linking PEG 2000 chains by an enzymatically degradable bis(E)K tripeptide linker in an interfacial polycon­ densation (dichloromethane‐water) of the bifunctional PEG bis(succinimidyl carbonate) with the oligopeptide‐based diamine linker protected by benzyl ester moieties [18]. Here, PEG and the linker were bound by a urethane bond and Dox was attached by an enzymatically degradable GFLG tetrapeptide spacer to a glutamyl residue. The multi‐block polymer‐Dox conjugate showed enhanced anti‐tumor activity in the treatment of mouse colorectal carcinoma C26. Subsequently, the peptide linker using a urethane bond was maintained, but Dox was bound by a pH‐sensitive hydrazone bond (Figure  13.4) [19]. Additional modifications were achieved by introducing hydrolytically

Structural Design and Synthesis of Polymer Prodrugs Synthesis of enzymatically degradable graft conjugate S S

Synthesis of reductively degradable graft conjugate O

O N

O

N HN

a

HO

PHPMA with terminal thiazolidine-2-thione (TT) groups +

x

O

HN

HO

5

O

NH

HN

N HN

2-Aminoethylpyridyldisulfid

O

a

HO

O

O

HN

b

5

O

NH

HN

O

O

O

PHPMA with terminal pyridyldisulfanyl groups +

O R1 NH

y

NH

NH

HN

O

S

b

5

O

HN

N

O

HN

S

z

HN

O

x

O

HN

HO

NH

HN

O

O

z

5

O

NH2

O R1 NH

y

HN

+



NH2Cl

O

O SH

Statistical PHPMA with primary aminogroups

Statistical PHPMA with sulfhydryl group

1. Conjugation reaction 2. Deprotection of hydrazide groups by trifluoroacetic acid 3. Conjugation with Dox.HCl

HN

O

HO O

x

O

HN

NH

O R1 NH

N

HN

O HO

R2

OH

HN

O

O

a

O

HN

HO

O

O

z

OH O

NH

R1 = NH

5

O OH

y

Enzymatically degradable graft conjugate:

O

NH2.HCl

O

OH

OH

R2 =

NH N

OH

OH O

O HO

Biodegradable graft PHPMA polymer conjugates with doxorubicin

O

NH2.HCl OH

NH

O

N Reductively degradable graft conjugate: O NH R1 = R2 =

O

O NH

GFLG oligopeptide spacer O

b

5

O

O +



NH2 Cl

S

S

NH N

Figure 13.3  Scheme of the synthesis of the HMW enzymatically or reductively degradable graft PHPMA conjugates with Dox bound by pH‐sensitive hydrazone bond.

397

398

Polymers for Biomedicine

O N

O

O

O

O 45

O

N

O

H2N

+

NH

NH2 O

O

O

Polycondensation Protected bis(E)K tripeptide O

OO

O O

NH 45

O

NH O

O

PEG bis(succinimidyl carbonate

O

O

OO

O

O

NH

NH

O

O NH n

O

O O

O HMW multi-block PEG 1. Hydrazinolysis 2. Dox.HCl NH2 NH

OO

O O

O

NH 45

O

OH

NH

NH O

O HN

O NH O HN

N

n

NH2

OH OH O

O HO

O O

NH2.HCl

HMW multi-block PEG conjugate with doxorubicin

OH

Figure 13.4  Scheme of the synthesis of the HMW enzymatically degradable multi‐block PEG conjugate with Dox bound by pH‐sensitive hydrazone bond.

degradable ester bonds (prepared by polycondensation of PEG bis‐succinate bearing terminal COOH groups activated to NHS esters with L‐lysine as a diamine linker [20]) and enzymatically and hydrolytically degradable bis(DP)K pentapeptides (prepared by polycondensation of PEG bis(succinimidyl ­carbonate) and the pentapeptide diamine linker [21]. With the development of controlled radical polymerization techniques, vari­ ous HPMA‐based telechelic or heterotelechelic polymers or copolymers were synthesized with narrow polydispersities. Biodegradable multi‐block PHPMA

Structural Design and Synthesis of Polymer Prodrugs

with enzymatically (GFLG tetrapeptide) degradable linkers were prepared by polyaddition reactions using thiol‐ene or azide‐alkyne click reactions [22–24]. A bifunctional RAFT agent with the GFLG tetrapeptide spacer was used for the preparation of telechelic PHPMA copolymers by RAFT polymerization initiated by 2,2′‐azobis‐(isobutyronitrile) (AIBN). After the removal of termi­ nal dithiobenzoate groups, free thiol groups were used in the polyaddition with bis‐maleimide, yielding a multiblock copolymer with an enzymatically degra­ dable HMW backbone [23]. Alternatively, a monofunctional RAFT agent con­ taining the GFLG tetrapeptide spacer terminated by a propargyl group was used for the synthesis of copolymers bearing alkyne chain ends. Heterotelechelic PHPMA containing both, alkyne and azide groups on opposite chain ends, was prepared by a post‐polymerization reaction with a diazido azoinitiator (Figure 13.5) [24]. Another bifunctional RAFT agent with two propargyl groups was used for the synthesis of telechelic PHPMA copolymers initiated by AIBN. The multi‐block copolymer was formed by the copper catalyzed polyaddition of α,ω‐dialkyne PHPMA and a diazido GFLG tetrapeptide linker [22]. The dis­ persity of the multi‐block PHPMA was wide due to the polyaddition reactions, although the precursor’s dispersity was quite low. Thus, the multi‐block copol­ ymers were fractionated in order to obtain HMW copolymers with narrow dispersities and used as carriers for Dox, PTX, and gemcitabine bound to the GFLG tetrapeptide spacer [25,26]. Star‐shaped dendrimer polymer structures, based again on PHPMA, are another approach to biodegradable HMW polymer carriers. Wang et  al. grafted  semitelechelic PHPMA terminated by NHS esters onto polyami­ noamide (PAMAM) dendrimers, and achieved molecular weights from 20,000 to 200,000 depending on the dendrimer generation (G2–G4 containing 16–64 surface amino groups, respectively) [27]. Dox was bound by a GFLG tetrapep­ tide spacer to the polymer precursor with the highest molecular weight. The drawback of the star‐like carrier was the absence of biodegradable ­linkages between the dendrimer core and polymer arms. The star‐like carrier system was further improved using enzymatically (GFLG tetrapeptide) or reductively (S‐S bond) degradable linkers [28]. While the previous semitel­ echelic polymer precursor (Mw = 5400 g/mol) used by Wang was prepared by radical chain‐transfer polymerization providing only short polymers, the use of functionalized azo‐initiators derived from 4,4′‐Azobis(4‐cyanovaleric acid) (ACVA) by Etrych et  al. prepared polymer precursors with Mw of about 25,000 g/mol or higher for the synthesis of star‐like PHPMA. Equally, thiol end groups on PHPMA, obtained in the reaction of TT‐terminated PHPMA with 2‐aminoethylpyridyldisulfide, followed by reduction with dithiothreitol were coupled to PAMAM dendrimers terminated by 2‐pyridyldisulfanyl groups, (Figure 13.6) [28]. The star polymer precursors were prepared by grafting the reactive sem­ itelechelic HPMA copolymer precursors onto the second or third generation

399

400

Polymers for Biomedicine 1. Polymerization HN

O

O

R NH

+

HO

OO

N

S

R

OH

S

N

AIBN, methanol, 50°C, 20 h O

O N F

HO

O NH

NH

2. Removal of dithiobenzoate groups O

N3

HO F

N

N

O

2

methanol, 70°C, 3 h O NH

NH OO

O

R

OH

N

O

HN

O

a

R NH

HO

N

b

N3

O

N O

O N F

HO

Heterotelechelic linear PHPMA conjugate with gemcitabine HO F Polyaddition

CuSO4/sodium ascorbate, H2O

O NH

NH N N N

O

O

O

R

OH

N

HN

O

HO

NH

R = NH O

O

O

R NH

b

N

O

N HO

HMW multi-block PHPMA conjugate with gemcitabine

a

O N F

O

HO F

O NH

NH O

Figure 13.5  Scheme of the synthesis of the HMW enzymatically degradable multi‐block PHPMA conjugate with gemcitabine bound by enzymatically degradable GFLG spacer.

n

Structural Design and Synthesis of Polymer Prodrugs

NH

O

N

N NH NH

2

N

S

O

PAMAM dendrimer (G2)

NH

O

NH O NH2

O

NH O

S O S NH

5

1. 2-Aminoethylpyridyldisulfid 2. Dithiothreitol

O

O

N

NH

N

N HN

O a

HN

O

HO SS

NH N O

NH

b

NH HN O O

PHPMA with terminal thiazolidine-2-thione (TT) groups

HS

O NH

S

O

O

N-Sukcinimidyl 3-(2-pyridyldisulfanyl)propionate

NH

N

HN

O

N

O

O a

HO

NH2

N S S HN

N N HN

O

NH N O

S

NH2

NH O

O NH O

O

S

H2N

O

N

NH O NH

b

5

NH HN

PHPMA with terminal thiol groups O

O

NH O NH

1. Conjugation reaction 2. deprotection of hydrazide groups by trifluoroacetic acid Modified PAMAM dendrimer with 2-pyridyldisulfanyl groups Star-like PHPMA carrier with free hydrazide groups O

HN

NH

O a

SS

2

HN

O b NO

NH

Dox.HCl

SS HN

HO O O OH

5

O

NH N OH OH

NH NH O

O NH

NH2.HCl OH

O

N

O O HO O O

N

S

O

NH

NH N O

NH

2

N

N O

SH O S NH

O SS

NH O O NH O

NH

N HN

O a

HN

HO O

NH O NH

O OH SSH

O

b

5

NH N OH OH

O O HO O Reductively degradable star-like PHPMA-Dox conjugate with drug bound by pH-sensitive hydrazone bond

O

NH2.HCl OH

Figure 13.6  Scheme of the synthesis of the HMW reductively degradable star‐like PHPMA conjugate with Dox bound by pH‐sensitive hydrazone bond.

401

402

Polymers for Biomedicine

PAMAM dendrimers containing 16 or 32 terminal amino groups and a diamin­ obutane or diaminohexane core. Non‐degradable star polymers ­containing Boc‐protected hydrazide groups were prepared by aminolysis of the TT end groups of the HPMA copolymers with amino groups of the PAMAM dendrimer in methanol. The enzymatically biodegradable star polymer was prepared by the reaction of the carboxyl end group of the GFLG‐OH terminal sequence of the linear polymer with amino groups of the PAMAM dendrimer in DMF using the carbodiimide coupling method. In both cases the polymer‐modified dendrimer was isolated by precipitation in ethyl acetate [11,32]. The molecular weight of the star‐like copolymers varied from 50,000 to 1,000,000 g/mol, depending on the dendrimer generation. The acceptably low dispersity (usually about Ð = 1.6–2.1) resulted from precursors prepared by RAFT polymerization and made fractionation unnecessary [29]. However, about 10% of the unbound linear precursor always remained, but did not ­substantially influence the in vivo anticancer activity of the Dox‐conjugates. In vivo evaluation of star‐like Dox‐conjugates showed prolonged blood cir­ culation, enhanced tumor accumulation and excellent anti‐tumor activity even at very low doses for various syngeneic tumor models (Figure  13.7) [11,30]. The crucial role of the molecular weight of the carrier was demonstrated by showing the impact on the circulation time, accumulation in tumors, tumor‐to blood and tumor‐to muscle ratio, and elimination via urine [31]. Enhanced therapeutic efficacy of star conjugates bearing other drugs than Dox; for instance, pirarubicin [33] and docetaxel [32] was also shown for the treatment of various syngeneic and xenograft tumors.

13.3 ­Synthetic Routes Macromolecular chemistry offers a range of polymerization techniques suita­ ble for the preparation of hydrophilic random multivalent, semitelechelic or telechelic polymers [34]. Conventional free radical and controlled/living radi­ cal polymerization (CRP) methods are typically used for the synthesis of PHPMA, the polymer considered here. 13.3.1  Radical Polymerization

A radical solution polymerization initiated by functionalized initiators ­represents a relatively simple way of preparing random multivalent and/or semitelechelic hydrophilic polymers. However, it is important to note that the functionality, molecular weight and dispersity of the polymers prepared in this way is limited by the mechanism of termination of chain growth (dispropor­ tionation vs recombination) and chain transfer reactions. Naturally, the termi­ nation process can be influenced to a certain extent by several factors, like

Structural Design and Synthesis of Polymer Prodrugs

(a) 10000

Tumour growth (mm3)

8000 6000 4000 2000 1500 1000 500 0 0

10

20

30

40

50

60

70

Time (days)

(b) 100

Surviving mice (%)

80 60 40 20 0 0

10

20

30

40

50

60

70

80

Time (days)

Figure 13.7  In vivo effect of polymer–Dox conjugates with pH‐triggered activation of drug on the growth of T cell lymphoma EL‐4 (a) and survival of mice (b) at a dose 1 × 15 mg Dox (eq.)/kg (—star‐like conjugate, Mw = 260,000 g/mol, Ð = 1.9, 10.2 wt.% Dox; —— grafted conjugate, Mw = 130,000 g/mol, Ð = 3.5, 9.6 wt.% Dox; ‐ ‐ ‐ linear conjugate, Mw = 35,000 g/ mol, Ð = 1.9, 9.9 wt.% Dox; ‐ ‐ ‐ untreated controls). Tumor growth: mean tumor volume is plotted; V = a*b2/2, where a = longer diameter, b = shorter diameter. Arrow marks administration of the treatment.

polymerization temperature, type of the solvent, initiator or monomer ­concentration, and so on. The original synthesis of PHPMA bearing reactive ONp is based on the ­radical precipitation copolymerization of HPMA with respective comonomers

403

404

Polymers for Biomedicine

in acetone initiated with AIBN [35]. By changing the copolymer composition, polymer precursors with varying amounts of reactive groups randomly distrib­ uted along polymer chains have been synthesized and employed in polymer– drug conjugate preparations in the 1980s and 1990s [36]. Molecular weights of the copolymers were limited to 20,000 to 35,000 g/mol due to chain‐transfer reactions and steric hindrances of the polymer radicals [35]. The dispersity of the copolymers is rather low, about 1.6–2.0, but their molecular weight cannot be easily controlled. Homogenous polymerizations carried out in organic sol­ vents (dimethylsulfoxide, N,N‐dimethylformamide) afford higher molecular weights and a slightly better control over the reaction. Unfortunately, dispersi­ ties are higher and the content of reactive groups decreased due to hydrolysis during the polymerization, leading to polymers with poorly defined structures. Later on, PHPMA with TT reactive groups have been developed for the pur­ pose of improving the precursor properties [37]. These copolymers can be prepared by solution polymerization, their molecular weight can be easily con­ trolled and the specific reactivity of TT groups makes it possible to perform aminolytic conjugation reactions, both in organic and aqueous solutions. All the reactive PHPMA precursors are multivalent with 10–15 reactive groups randomly distributed along the polymer chain. Two methods are available for the synthesis of semitelechelic polymers: radical polymerization or copolym­ erization of HPMA in the presence of chain‐transfer agents [38] mainly thiol group‐containing acids, or radical copolymerization using bifunctional azo‐ initiators containing TT groups [16]. The terminal TT group can be further modified, for instance, with maleimido [39] or 2‐pyridyldisulfanyl [40] groups. Such semitelechelic PHPMA were used for the conjugation with proteins and glycoproteins and for the synthesis of branched, grafted and star‐like HMW polymer–Dox conjugates (Figure 13.6). Molecular weights of polymers prepared by chain‐transfer polymerization range from 3,000 to 15,000 g/mol, on the other hand, copolymerization using bi‐functional azo‐initiators achieve higher molecular weights (10,000– 100,000 g/mol), but the dispersity is higher as well. Polymers prepared by chain‐transfer polymerization require thorough purification as polymer chains without reactive end group are always present. 13.3.2  Controlled Polymerization Technique

Low dispersity is one of the most important requirements for polymer drug conjugates designed for in vivo applications, therefore, controlled polymeriza­ tions of HPMA have been investigated. While ATRP of HPMA in bulk or solution led only to low conversions when common ATRP ligands were ­ employed [41], RAFT polymerizations in acetate buffer yielded HPMA poly­ mers with molecular weights from 15,000 to 97,000 g/mol and a dispersity below 1.1 [42]. The introduction of functional groups was achieved by RAFT

Structural Design and Synthesis of Polymer Prodrugs

copolymerizations of HPMA and N‐(methacryloyloxy) succinimide [43] or pentafluorophenyl methacrylate [44]. The molecular weight of the copolymers was between 3000–135,000 g/mol with a dispersity of 1.1–1.3. Limiting factors for CRP methods are often their incompatibility with polymerizing water soluble monomers and the use of toxic additives (solvent, initiator, catalyst, chain transfer agents, etc.). However, the following syntheses meet the stringent requirements for the production of safe and efficient drug delivery systems. The selection of the appropriate RAFT agents for individual monomers and reaction conditions is most crucial [45]. Typically heterobi­ functional dithioesters, trithiocarbonates, or xanthates (with the general for­ mula ZC(=S)SR) are used as chain transfer agents that reversibly deactivate propagating radicals to reduce the effective concentration of active chain radi­ cals and the probability of mutual chain termination. Xu et al. activated the carboxyl group of a dithiobenzoate‐based RAFT agent with TT to control the polymerization of HPMA [46]. The dispersity and molecular weights of PHPMA materials prepared were consistent with a con­ trolled polymerization mechanism. The TT end group remained intact throughout the polymerization leading to a TT group terminated PHPMA, which was then exploited to attach the PHPMA to a dendrimer [29,46] or to conjugate it to lysozyme [47]. Barz et  al. synthesized a RAFT agent and a diazoinitiator, both containing a pentafluorophenyl (PFP) activated ester. Subsequent RAFT polymerizations of several methacrylates resulted in homopolymers and diblock copolymers with an excellent control over the molecular weight distribution [48]. These polymers are suitable for further modifications with amino group‐bearing molecules, like drugs or antibodies. Boyer et  al. synthesized poly(N‐isopropylacrylamide) using 3‐(benzylsulfa­ nylthiocarbonylsulfanyl)‐propionic acid as a RAFT agent and used AIBN as initiator [49]. The aminolysis of the Z polymer end‐groups in the presence of  2,2′‐dithiopyridyl disulfide yielded 2‐pyridyl disulfanyl groups that are ­available for subsequent reactions with different thiol group‐modified biologi­ cally active compounds using thiol‐disulfide exchange chemistry. Chytil et al. ­demonstrated the RAFT polymerization of HPMA and 1‐(tert.‐butoxycar­ bonyl)‐2‐(6‐methacrylamido hexanoyl)hydrazine initiated by AIBN in the presence of 4‐cyano‐4‐thiobenzoylsulfanyl‐pentanoic acid as chain transfer agent [50]. The dithiobenzoate end‐group was removed by reduction with sodium borohydride followed by an in situ reaction with N‐(2‐aminoethyl) maleimide trifluoroacetate. A semitelechelic copolymer bearing a TT reactive end‐group was subsequently prepared by the reaction of the homobifunctional TT reagent 3,3′‐disulfanediylbis[1‐(2‐thioxothiazolidin‐3‐yl)propan‐1‐one] with the amino group‐containing polymer. The syntheses of telechelic or heterotelechelic copolymers are a prerequisite for the successful preparation of advanced drug delivery systems. For example, RAFT agents containing enzymatically degradable linkers [23,24] or azide

405

406

Polymers for Biomedicine O

S O

S

HN

HN HO

O

S S

+

O

N

S

S

S

N N HN

N

N

N O

O

S

S

N

N

Dithiobenzoate groups O

NH

HN

O

O

O

O N

NH

N3

N

2 DMSO, 70°C, 4 h

O

S

NH

N N HN TT

b

HO

O

S

O

HN

2

tert.-butyl alcohol/DMSO, 70°C, 16 h

NH HN

O a

S

O a

O

HN

b

N

N3

O Azide groups

HO

O

NH

HN O

O

Boc-protected hydrazide groups

Figure 13.8  Scheme of the synthesis of heterotelechelic PHPMA containing reactive terminal groups; i.e., TT for aminolytic reactions and azide groups for azide‐alkyne click reactions, and tert‐butyloxycarbonyl (Boc) group‐protected hydrazide groups distributed along the polymer chain enabling attachment of drug by hydrazone bond after deprotection.

groups [22] were prepared for the synthesis of HMW multi‐block copolymers. The copolymer bearing a TT and an azido group on opposite polymer chain  ends [29] (Figure  13.8) was used for the preparation of the star‐like copolymers.

13.4 ­Drug Coupling Strategies In order to achieve adequate biological effects the active drug must be released from the polymer carrier only in the target tissue or cell. This is especially important in cancer treatments as cancer drugs are highly toxic and should remain bound to the carrier during transport through the blood stream. Most polymer prodrug conjugates exploit a spacer that is either enzymatically degradable or stimuli‐sensitive and enables the tumor‐specific drug release. In general, polymer drug conjugates can be synthesized either directly by the polymerization of monomers containing a drug or by post‐polymerization

Structural Design and Synthesis of Polymer Prodrugs

drug attachment. The direct polymerization enables precise control over the polymer composition but not all drugs survive polymerization conditions and also copolymerization parameters are not always favorable. Unused spacers are  always left in the post‐polymerization drug attachment method (see Figure 13.10 later). The majority of studied polymer‐drug conjugates contain spacers, which are degradable by lysosomal enzymes, thus ensuring the drug is released only inside tumor cells. Peptide, glycoside and phosphate bonds are susceptible to cleavage by lysosomal enzymes, but α‐aminoacids or oligopeptides are most commonly used [1]. After studying the effects of various oligopeptide struc­ tures in PHPMA drug conjugates on the drug release and stability in aqueous solutions, the GFLG spacer was selected due to its stability in blood and degra­ dability in the presence of lysosomal enzymes (41.6% of released Dox after 46 h incubation with lysosomal enzyme cathepsin B) (Figure 13.9) [51]. The supe­ rior anti‐tumor activity of PHPMA polymer conjugates was first shown for anticancer agents; for example, puromycin, Dau, Dox, using the GFLG tetra­ peptide spacer and later various other drugs; for example, methotrexate, ami­ noellipticine, geldamycin, campthothecin, PTX, and so on, were introduced using the same oligopeptide spacer [52,53]. Several conjugates with anticancer agents bound by GFLG are now in clinical trials (PHPMA conjugates with Dox (PK1, PK2), camptothecin, PTX, or platinates) [54,55]. Usually, polymer carriers bearing oligopeptides terminated by ester or amide reactive groups, for instance, ONp or TT group, were used for the attachment of drugs containing primary amino groups, for example, anthracyclines. The drugs without available amino groups were modified before their attachment to the carrier, for example, methotrexate, 5‐fluorouracil, platinates, ellipticine, or carboxyl‐bearing polymer precursors were utilized for the attachment of drugs containing hydroxyl group, for example, camptothecin, by forming an ester bond. Besides lysosomal enzymes other specific enzymes can be exploited for con­ trolled drug release. Spacers containing an aromatic azo bond can be cleaved by an azo‐reductase in the colon in the treatment of colon diseases [56]. The alternatives to enzymatically degradable spacers are pH‐sensitive hydro­ lysable spacers that degrade at pH 5–6. Compared to enzymatically degradable spacers pH‐sensitive spacers are advantageous because of their independence of certain enzyme levels, and their synthesis is often easier and less expensive. However, small amounts of drug can be released into the blood stream at physiological pH as they are slowly hydrolyzed even at neutral pH. Since ­residence times in the blood stream are on the order of minutes to hours, pH‐ sensitive spacers can be selected if their hydrolysis rate at mild acidic pH is much higher than at neutral pH. Several spacers; for example, hydrazone, cis‐aconityl, trityl bonds, and hydrolyzable oligopeptide have been utilized as pH‐sensitive linkages [59] (Figure 13.9).

407

Polymers for Biomedicine

(a) O

O a

HN HO

HN

b

HN

O

O a

O

O

O HN

O

O

O

O

OH O

hydrazone bond

O NH2.HCl

O

OH

O OH O

OH O

1

O HO

OH

HO

HO

(b)

O

HO

HO O

N OH

OH

O OH O OH O

NH OH

OH

cis-aconityl spacer

NH

b

R

HN

GFLG spacer

O

O

O a

HO O

O

HN

HN

NH

NH

HN

b

R

HO

O

3 O

R represents aminoacid or oligopeptide spacer; e.g., G, GG, GFLG, 4-aminobenzoic acid, 6-aminohexanoic acid.

2

100 90 80 Dox release (%)

408

70 60 50 40 30 20 10 0 0

10

20

30

40

50

Time (h)

Figure 13.9  Schematic structure of the PHPMA‐Dox conjugates differing in the structure of biodegradable spacer: enzymatically degradable glycyl‐DL‐phenylalanyl‐L‐leucyl‐glycyl spacer (1) and pH‐sensitive cis‐aconityl (2) and hydrazone (3) spacers (a). The drug release from the conjugates incubated at 37°C in phosphate buffers at pH 7.4 (▲‐ ‐ ‐ 2; ●‐ ‐ ‐ 3) and pH 5.0 (∇— 1 ▲— 2; ●— 3) (b). (The conjugate 1 was incubated in the presence of 0.5 μM cathepsin B. The conjugates 2 and 3 contained glycyl‐glycyl spacer as R. Source: Duncan 2010 and Ulbrich 2003 [57,58].

The hydrazone bond is of interest because of its relative stability at neutral pH and its increasing hydrolysis rate with decreasing pH. A hydrazone bond is formed between the C13 keto group of Dox and the hydrazide group along the polymer carrier [15,19]. Alternatively, Dox can be introduced directly by the polymerization of HPMA with a monomer containing drug (Figures  13.10a

Structural Design and Synthesis of Polymer Prodrugs

(a) O HN

O HN

Polymerization

+

HO

5

O O

OH

HN

O

O x

HN

HO

N

O

OH

OH

O OH OH

OH O

NH2.HCl

O

OH

OH

O

O HO

O

NH N

OH O

5

O

NH

y

O

NH2.HCl

O NH2.HCl

O

OH

OH

Doxorubicin

OH

(b)

O

O HO

O

O HO

PHPMA-doxorubicin conjugate O HN

O HN

Polymerization

+

5

O

HO

HN

HO

NH

O

NH2

(c)

O

O x

HN

O

Drug conjugation

y

5 NH NH2

O

O

O

O

Drug conjugation OH

O

Drug derivatization

O

O

NH

O O

HO

O

O

O O

HN

O x

HO

Paclitaxel levulinate

OH O

O

O

NH

NH N

OH

O

O

OH O

y

5

O

O O

O HN

O

O

OH

O O

O

HO

O

O

O O

O

NH

O O

HO

O

Paclitaxel

O

O

O O

O

PHPMA-paclitaxel conjugate

Figure 13.10  Scheme of the synthesis of linear PHPMA conjugates by polymerization of polymerizable Dox derivative (a) or by post‐polymerization reaction with Dox.HCl (b) or a PTX oxoderivative (c).

and b) [60]. The attachment of Dox via a hydrazone bond allows for ­significantly higher drug loading than that described previously for the lysoso­ motropic prodrug system. Linear PHPMA with a Dox loading up to 24 wt% maintained full solubility of the drug conjugates, whereas loadings higher than 8 wt% Dox bound via amide bonds to the GFLG spacer lead to aggregation and

409

410

Polymers for Biomedicine

precipitation of the conjugate in aqueous solutions. Hydrazide precursors are readily synthesizable over a wide range of molecular weights [57,60] and hydrazone conjugates of Dox showed a better in vivo anti‐tumor effect com­ pared to GFLG conjugates. Drugs containing hydroxyl group; for example, dexamethasone, PTX or docetaxel, or amino groups; for example, 5‐fluorouracil, 9‐aminocamptoth­ ecin, reversin 121, are suitable for derivatization after introducing ketogroups (Figure  13.10c). The structure of the used oxoacid spacer is crucial for the final therapeutic outcome of a given prodrug because: (i) it influences the drug release rate and (ii) the derivatization should change the biological activity of the drug. Since the oxoderivative of the drug, not the drug itself, is  released it must be verified that the biological activity; for example, ­c ytoxicity, of the active agent is maintained. However, the degradation of the derivative to the free drug can be expected due to the presence of lysosomal enzymes or esterases. The in vivo fate of released model drugs was studied by non‐invasive multi­ spectral optical imaging. Dual fluorescent HPMA conjugates, containing a near infrared fluorescent dye coupled via a stable hydrazide bond functioning as the carrier label and another far red dye modelling the drug bound to the carrier via an oxoacid spacer with different structures were investigated (Figure 13.11a) [61]. It was shown that the conjugate with the slower releasing spacer (about 1–2 % of the drug was released at pH 7.4 within 5 h; 12% the drug was released at pH 5.0 within 5 h, Figure 13.11b) circulated longer in blood and its tumor accumulation was also much higher (Figure 13.11b). On the other hand, the model drug that was released quickly (10% of the drug was released at pH 7.4 within 5 h; 90% of the drug was released at pH 5.0 within 5 h) could not reach the tumor effectively. The effect was independent of the carrier structure. Cis‐aconityl acid is another pH‐sensitive spacer and Dox release from the conjugates was pH‐dependent, but compared to the hydrazone‐containing spacer much slower, over the studied pH range. The linkage was completely stable at neutral pH modeling the blood stream. The drug release rate at lower pH was also significantly lower compared with the Dox‐containing hydrazone drug conjugate. Less than 40% of drug was released within 48 h of incubation (Figure  13.11b). PHPMA‐Dox conjugates exhibited significant cytotoxicity against human ovarian carcinoma and mouse EL‐4 lymphoma, but the in vivo therapeutic efficacy on mouse EL‐4 lymphoma was significantly lower com­ pared to hydrazone‐containing PHPMA‐Dox conjugates. The extremely low release rate led to unsatisfactory results [58,62]. Another type of stimuli‐sensitive spacers consists of reductively sensitive disulfide linkages. They are stable during blood circulation and in extracellular fluids, and degrade rapidly in the reductive environment present in intracellu­ lar compartments such as the cytoplasm and the cell nucleus [63]. Some

Structural Design and Synthesis of Polymer Prodrugs

(a) OPB

IPB

PYR

0.123

0.217

0.154

p.i. 0.10

1d 0.10

2d 0.10

3d

0.05

0.05

4d 5d 0.00

0.00

0.00

(b) 100

Model drug release (%)

90 80 70 60 50 40 30 20 10 0

0

5

10

15

20

25

Time (h)

Figure 13.11  Drug model (fluorescent dye DY‐676) distribution in human colon carcinoma‐ bearing nude mice after injection of star‐like PHPMA containing different spacers – 4‐(2‐ oxopropyl)benzoyl (OPB), 4‐isopropyl‐4‐oxobutanoyl (IPB), and 4‐oxo‐4‐(2‐pyridyl)butanoyl (PYR) spacers. Source: Chytil 2013 [61]. Reproduced with permission of Elsevier. (a) Release of drug model from star‐like PHPMA incubated in phosphate buffered saline at pH 5.0 (●— OPB; ▲— IPB; ∇— PYR) and pH 7.4 (●‐ ‐ ‐ OPB; ▲‐ ‐ ‐ IPB; ∇‐ ‐ ‐ PYR.) at 37°C (b).

411

412

Polymers for Biomedicine

water‐soluble polymer‐drug conjugates with a reductive release mechanism were described; for example, photosensitizer mesochlorin e6, methotrexate [64,65]. In recent years, a number of polymer‐drug conjugates containing self‐immo­ lative spacers were proposed. Here, the drug is released after the fragmentation of the spacer is caused by a triggering action; for example, enzymolysis, pH‐ sensitive hydrolysis, and so on [66]. One of the most successful self‐immolative spacer is p‐aminobenzyloxycarbonyl forming an acid labile imine bond that hydrolyses in the acidic environment of tumor tissue or cells. The advantages of self‐immolative spacers lie in the possibility for precise control of the degra­ dation rate and the drug is released in its original form, not as derivative.

13.5 ­Elimination of Carriers All carriers should be eliminated from the body to avoid their long‐term accu­ mulation in the organism after the biologically active cargo is delivered. The balance between elimination of polymer drugs from the blood stream by the kidneys, liver and other organs and extravasation from the blood vasculature into the tumor influences strongly the effectiveness of the drug delivery system [67]. Glomerular kidney permeability decreases with increasing hydrodynamic radius and molecular weight and is also influenced significantly by polymer charge. The clearance of negatively charged macromolecules is restricted, while the clearance of positively charged macromolecules with the same molecular weight is enhanced [68–70]. The renal clearance of linear flexible macromolecules, such as dextran and poly(vinylpyrrolidone), is up to 10 times greater than that of proteins with an equivalent hydrodynamic radius [68]. Macromolecular branching leads to decreased renal clearance and significantly prolongs blood circulation time [71]. Hence, branched, grafted, star‐like or multiblock PHPMA conjugates or multiblock PEG conjugates showed prolonged blood circulation and enhanced tumor accumulation and activity against various solid tumor models. Polymer chains of molecular weights of 2000 g/mol for PEG and below 50,000 g/mol for PHPMA were connected by biodegradable spacers to each other or to a den­ drimer core. Enzymatically, reductively, or hydrolytically degradable spacers designed for the intracellular degradation of the high‐molecular‐weight poly­ mer carriers to excretable products were proposed. It was shown that grafted, diblock, or multi‐block polymer carriers containing S‐S bonds were degraded within several hours to polymer fragments with molecular weights below the limit of renal filtration in a pH 6.0 buffer containing 3 mM of glutathione, which modeled the cytosolic environment [40]. Similarly, HMW conjugates containing GFLG oligopeptide spacers were degraded within 24 h for grafted and 72 h for star‐like conjugates in a pH 6.0 buffer containing 0.5 μM cathepsin B that represented lysosomal enzymes [16,28]. Probably, the steric hindrance

Structural Design and Synthesis of Polymer Prodrugs

prevented easy access of enzymes to the GFLG oligopeptide substrate, ­especially in the case of highly robust and dense star‐like conjugates. Multi‐block PHPMA conjugates containing GFLG oligopeptide spacers were degraded within 24 h in 5.6 μM cathepsin B [22]. Only about 10% of the starting PEG 2000 was detected after 48 in cathepsin B (0.4 μM) for PEG multi‐blocks linked by GFLG spacers. However, the molecular weight of the multi‐block polymer changed dramati­ cally, indicating exo‐ and endo‐type degradation of the polymer carrier [18]. The multi‐block PEG conjugates linked by the enzymatically and hydrolytically degradable bis(DP)K pentapeptide were degraded by a similar manner. Only about 20% of mPEG was detected after hydrolysis of a model diblock copolymer in buffer pH 5.0, but about 50% of mPEG was detected after enzymolysis of the diblock copolymer in cathepsin B (0.8 μM) [21]. The biodegradability of HMW polymer conjugates was also shown in vitro on cell cultures [11,40]. Biodegradability tests of fluorescently labeled polymer carriers bearing the dye DY‐615 in suspensions of EL4 T‐cell lymphoma cells showed that the rate of degradation was much faster for reductively degradable star‐like conjugates (close to completion within 24 h of incubation) than for enzymatically degradable star‐like conjugates (several days). See Figure 13.12 for gel permeation chromatograms at 615 nm showing the time‐dependence of the degradation of reductively degradable star‐like conjugates. This finding 600 500

A (550 nm)

400 300 200 100 0 10

15

20

25

30

35

40

45

50

Retention time (min)

Figure 13.12  Gel permeation chromatograms of reductively degradable star‐like PHPMA fluorescently labeled by DY‐615 (—) and its degradation products after 6 h‐ (—), 12 h‐ (‐ ‐ ‐), and 24 h‐incubation (∙∙∙∙∙) with EL4 T‐cells. Fluorescently labeled linear PHPMA (‐ ‐ ‐) was used as control. Source: Etrych 2011 [11]. Reproduced with permission of Elsevier.

413

414

Polymers for Biomedicine

was likely due to the differences in the accessibility for the small glutathione molecule and the bulky enzyme cathepsin B. Various PHPMA‐Dox conjugates with different molecular weights, dispersi­ ties and molecular architectures (flexible linear vs less flexible star‐like struc­ tures) were investigated in vivo for their blood clearance, renal elimination, tumor and liver accumulation, and survival of mice bearing EL 4 T‐cell lym­ phoma [31]. The molecular weights, determined by gel permeation chromatog­ raphy using static light scattering detection, of linear polymer‐Dox conjugates excreted in the urine of mice by renal filtration were up to 70,000 g/mol, while the highest molecular weight of the star conjugates excreted in urine was ~50,000 g/mol. Based on these findings it was hypothesized that polymers will be eliminated by glomerular filtration up to a limit of about 50,000 g/mol and longer linear polymer chains pass through the renal filtration by a slower pro­ cess (worm‐like motion of polymer chains), which is interdicted for the more rigid star‐like polymers. The amount of polymer‐Dox conjugate eliminated by renal filtration decreased with increasing molecular weight, and linear conju­ gates with comparable molecular weights to the star conjugates were elimi­ nated more quickly. Polymers were not excreted as the whole polymer fraction, but rather as narrow polymer fractions with molecular weights gradually increasing with time following injection. Polymer‐Dox conjugates with higher molecular weights exhibited longer blood circulation times and higher tumor accumulations, which resulted in significantly improved anti‐tumor activities of the HMW conjugates in vivo. These findings were confirmed in vivo using the dual fluorescent polymer drug conjugates described previously [72]. In contrast to other in vivo studies, the authors were able to track the in vivo fate for several weeks and to observe the distribution of the polymer and a conjugated model drug simultaneously by multispectral optical imaging. For that purpose, HPMA copolymers differ­ ing in polymer architecture, linear and star‐like, were labeled with two fluores­ cent probes with different emission properties. The relative biodistribution in the body between the 30,000 g/mol linear and 200,000 g/mol star‐like polymers did not differ significantly, but the star‐like polymer circulated much longer. Although the polymers were non‐degradable, even the star‐like polymer was partly eliminated within three months. As the HPMA copolymers were not biodegradable, the authors assumed a major influence of hepatobiliary elimi­ nation, especially for the star‐like polymer, which was confirmed by fluores­ cent signals coming from the intestine.

13.6 ­Conclusion The development of new synthetic routes based on controlled polymerization techniques, opens new horizons for more sophisticated prodrug‐based drug delivery systems. Various stimuli‐sensitive spacers enabling controlled drug

Structural Design and Synthesis of Polymer Prodrugs

release and controlled biodegradability of the polymers were designed and described. The design of tailor‐made polymer therapeutics and the tissue‐ or cell‐ specific drug delivery are promising for future prodrug drug delivery systems.

­Acknowledgement This work was supported by the Ministry of Education, Youth and Sports of CR within the National Sustainability Program I, Project POLYMAT LO1507.

­References 1 Drobník, J., Kopeček, J., Labský, J., Rejmanová, P., Exner, J., Saudek, V., Kálal, J.:

Enzymatic cleavage of side chains of synthetic water‐soluble polymers. Makromolekulare Chemie 177, 2833–2848. 1976. 2 Putnam, D., Kopeček, J.: Polymer conjugates with anticancer activity. Biopolymers 122, 55–123. 1995. 3 Haag, R., Kratz, F.: Polymer therapeutics: Concepts and applications. Angewandte Chemie, International Edition in English 45, 1198–1215. 2006. 4 Schacht, E.H., Vansteenkiste, S., Seymour, L.W., Kostelnik, R.J.: Macromolecular carriers for drug targeting. In: The Practice of Medicinal Chemistry. Academic Press, New York, 1996, pp. 718–736. 5 Li, C.: Poly(L‐glutamic acid) – anticancer drug conjugates. Advanced Drug delivery Reviews 54, 695–713. 2002. 6 Duncan, R., Vicent, M.J.: Polymer therapeutics‐prospects for 21st century: The end of the beginning. Advanced Drug delivery Reviews 65, 60–70. 2013. 7 Hudecz, F., Clegg, J.A., Kajtar, J., Embleton, M.J., Szekerke, M., Baldwin, R.W.: Synthesis, conformation, biodistribution, and invitro cytotoxicity of daunomycin branched polypeptide conjugates. Bioconjugate Chemistry 3, 49–57. 1992. 8 Gaal, D., Hudecz, F.: Low toxicity and high antitumour activity of daunomycin by conjugation to an immunopotential amphoteric branched polypeptide. European Journal of Cancer 34, 155–161. 1998. 9 Maeda, H., Matsumura, Y.: EPR effect based drug design and clinical outlook for enhanced cancer chemotherapy. Adv Drug Deliv Rev 63, 129–130. 2011. 10 Seymour, L.W., Miyamoto, Y., Maeda, H., Brereton, M., Strohalm, J., Ulbrich, K., Duncan, R.: Influence of molecular‐weight on passive tumor accumulation of a soluble macromolecular drug carrier. European Journal of Cancer 31A, 766–770. 1995. 11 Etrych, T., Kovář, L., Strohalm, J., Chytil, P., Říhová, B., Ulbrich, K.: Biodegradable star HPMA polymer‐drug conjugates: Biodegradability, distribution and anti‐tumor efficacy. Journal of Controlled Release 154, 241–248. 2011.

415

416

Polymers for Biomedicine

12 Kopeček, J., Bažilová, H.: Poly[N‐(2‐hydroxypropyl)methacrylamide]—I.

13 14

15

16

17

18

19

20

21

22

23

24

Radical polymerization and copolymerization. European Polymer Journal 9, 7–14. 1973. Kopeček, J., Kopečková, P.: HPMA copolymers: Origins, early developments, present, and future. Advanced Drug Delivery Reviews 62, 122–149. 2010. Dvořák, M., Kopečková, P., Kopeček, J.: High‐molecular weight HPMA copolymer‐adriamycin conjugates. Journal of Controlled Release 60, 321–332. 1999. Etrych, T., Jelínková, M., Říhová, B., Ulbrich, K.: New HPMA copolymers containing doxorubicin bound via pH‐sensitive linkage: synthesis and preliminary in vitro and in vivo biological properties. Journal of Controlled Release 73, 89–102. 2001. Etrych, T., Chytil, P., Mrkvan, T., Šírová, M., Říhová, B., Ulbrich, K.: Conjugates of doxorubicin with graft HPMA copolymers for passive tumor targeting. Journal of Controlled Release 132, 184–192. 2008. Pasut, G., Veronese, F.M.: PEG conjugates in clinical development or use as anticancer agents: An overview. Advanced Drug Delivery Reviews 61, 1177– 1188. 2009. Pechar, M., Ulbrich, K., Šubr, V., Seymour, L.W., Schacht, E.H.: Poly(ethylene glycol) multiblock copolymer as a carrier of anti‐cancer drug doxorubicin. Bioconjugate Chemistry 11, 131–139. 2000. Pechar, M., Braunová, A., Ulbrich, K., Jelínková, M., Říhová, B.: Poly(Ethylene glycol) ‐ Doxorubicin conjugates with pH‐controlled activation. Journal of Bioactive and Compatible Polymers 20, 319–341. 2005. Raunová, A., Pechar, M., Ulbrich, K.: Degradation behavior of poly(ethylene glycol) diblock and multiblock polymers with hydrolytically degradable ester linkages. Collection of Czechoslovak Chemical Communications 69, 1643– 1656. 2004. Pechar, M., Braunová, A., Ulbrich, K.: Poly(ethylene glycol)‐based polymer carrier of doxorubicin degradable by both enzymatic and chemical hydrolyses. Collection of Czechoslovak Chemical Communications 70, 327–338. 2005. Luo, K., Yang, J., KopečKová, P., KopečEk, J.I.: Biodegradable multiblock poly[N‐(2‐hydroxypropyl)methacrylamide] via reversible addition − fragmentation chain transfer polymerization and click chemistry. Macromolecules 44, 2481–2488. 2011. Pan, H.Z., Yang, J.Y., Kopečková, P., Kopeček, J.: Backbone degradable multiblock N‐(2‐hydroxypropyl)methacrylamide copolymer conjugates via reversible addition‐fragmentation chain transfer polymerization and thiol‐ene coupling reaction. Biomacromolecules 12, 247–252. 2011. Yang, J., Luo, K., Pan, H., Kopečková, P., Kopeček, J.: Synthesis of biodegradable multiblock copolymers by click coupling of RAFT‐generated heterotelechelic polyHPMA conjugates. Reactive & Functional Polymers 71, 294–302. 2011.

Structural Design and Synthesis of Polymer Prodrugs

25 Kopeček, J.: Polymer–drug conjugates: Origins, progress to date and future

directions. Advanced Drug delivery Reviews 65, 49–59. 2013.

26 Yang, J., Kopeček, J.: Macromolecular therapeutics. Journal of Controlled

Release 190, 288–303. 2014.

27 Wang, D., Kopečková, P., Minko, T., Nanayakkara, V., Kopeček, J.: Synthesis of

28

29

30

31

32

33

34

35

36 37

starlike N‐(2‐hydroxypropyl)methacrylamide copolymers: Potential drug carriers. Biomacromolecules 1, 313–319. 2000. Etrych, T., Strohalm, J., Chytil, P., Černoch, P., Starovoytova, L., Pechar, M., Ulbrich, K.: Biodegradable star HPMA polymer conjugates of doxorubicin for passive tumor targeting. European Journal of Pharmaceutical Sciences 42, 527–539. 2011. Chytil, P., Koziolova, E., Janouskova, O., Kostka, L., Ulbrich, K., Etrych, T.: Synthesis and properties of star HPMA copolymer nanocarriers synthesised by RAFT polymerisation designed for selective anticancer drug delivery and imaging. Macromolecular Bioscience 15, 839–850. 2015. Etrych, T., Strohalm, J., Chytil, P., Říhová, B., Ulbrich, K.: Novel star HPMA‐ based polymer conjugates for passive targeting to solid tumors. Journal of Drug Targeting 19, 874–889. 2011c. Etrych, T., Šubr, V., Strohalm, J., Šírová, M., Říhová, B., Ulbrich, K.: HPMA copolymer‐doxorubicin conjugates: The effects of molecular weight and architecture on biodistribution and in vivo activity. Journal of Controlled Release 164, 346–354. 2012. Etrych, T., Strohalm, J., Sirova, M., Tomalova, B., Rossmann, P., Rihova, B., et al.: High‐molecular weight star conjugates containing docetaxel with high anti‐tumor activity and low systemic toxicity in vivo. Polym. Chem. 6, 160–170. 2015. Nakamura, H., Koziolova, E., Etrych, T., Chytil, P., Fang, J., Ulbrich, K., Maeda, H.: Comparison between linear and star‐like HPMA conjugated pirarubicin (THP) in pharmacokinetics and antitumor activity in tumor bearing mice. European Journal of Pharmaceutics and Biopharmaceutics 90, 90–96. 2015. Soyez, H., Schacht, E., Vanderkerken, S.: The crucial role of spacer groups in macromolecular prodrug design. Advanced Drug Delivery Reviews 21, 81–106. 1996. Strohalm, J., Kopeček, J.: Poly[N‐(2‐hydroxypropyl)methacrylamide]. IV. Heterogeneous polymerization. Angewandte Makromolekulare Chemie 70, 109–118. 1978. Duncan, R.: Designing polymer conjugates as lysosomotropic nanomedicines. Biochemical Society Transactions 35, 56–60. 2007. Šubr, V., Ulbrich, K.: Synthesis and properties of new N‐(2‐hydroxypropyl)‐ methacrylamide copolymers containing thiazolidine‐2‐thione reactive groups. Reactive & Functional Polymers 66, 1525–1538. 2006.

417

418

Polymers for Biomedicine

38 Oupicky, D., Ulbrich, K., Rihova, B.: Conjugates of semitelechelic poly[N‐(2‐

39

40

41 42

43

44

45

46

47

48

49

hydroxypropyl) methacrylamide] with enzymes for protein delivery. Journal of Bioactive and Compatible Polymers 14, 213–231. 1999. Etrych, T., Mrkvan, T., Říhová, B., Ulbrich, K.: Star‐shaped immunoglobulin‐ containing HPMA‐based conjugates with doxorubicin for cancer therapy. Journal of Controlled Release 122, 31–38. 2007. Etrych, T., Kovář, L., Šubr, V., Braunová, A., Pechar, M., Chytil, P., et al.: High‐molecular‐weight polymers containing biodegradable disulfide bonds: synthesis and in vitro verification of intracellular degradation. Journal of Bioactive and Compatible Polymers 25, 5–26. 2010. Teodorescu, M., Matyjaszewski, K.: Atom transfer radical polymerization of (meth)acrylamides. Macromolecules 32, 4826–4831. 1999. Scales, C.W., Vasilieva, Y.A., Convertine, A.J., Lowe, A.B., Mccormick, C.L.: Direct, controlled synthesis of the nonimmunogenic, hydrophilic polymer, poly(N‐(2‐hydroxypropyl)methacrylamide) via RAFT in aqueous media. Biomacromolecules 6, 1846–1850. 2005. Yanjarappa, M.J., Gujraty, K.V., Joshi, A., Saraph, A., Kane, R.S.: Synthesis of copolymers containing an active ester of methacrylic acid by RAFT: Controlled molecular weight scaffolds for biofunctionalization. Biomacromolecules 7, 1665–1670. 2006. Herth, M.M., Barz, M., Moderegger, D., Allmeroth, M., Jahn, M., Thews, O., et al.: Radioactive labeling of defined HPMA‐based polymeric structures using [F‐18]FETos for in vivo imaging by positron emission tomography. Biomacromolecules 10, 1697–1703. 2009. Šubr, V., Kostka, L., Strohalm, J., Etrych, T., Ulbrich, K.: Synthesis of well‐ defined semitelechelic poly[N‐(2‐hydroxypropyl)methacrylamide] polymers with functional group at the α‐end of the polymer chain by RAFT polymerization. Macromolecules 46, 2100–2108. 2013. Xu, J.T., Boyer, C., Bulmus, V., Davis, T.P.: Synthesis of Dendritic Carbohydrate End‐Functional Polymers via RAFT: Versatile Multi‐Functional Precursors for Bioconjugations. Journal of Polymer Science Part A‐Polymer Chemistry 47, 4302–4313. 2009. Tao, L., Liu, J.Q., Xu, J.T., Davis, T.P.: Synthesis and bioactivity of poly(HPMA)‐lysozyme conjugates: the use of novel thiazolidine‐2‐thione coupling chemistry. Org. Biomol. Chem. 7, 3481–3485. 2009. Barz, M., Tarantola, M., Fischer, K., Schmidt, M., Luxenhofer, R., Janshoff, A., et al.: From defined reactive diblock copolymers to functional HPMA‐based self‐assembled nanoaggregates. Biomacromolecules 9, 3114–3118. 2008. Boyer, C., Granville, A., Davis, T.P., Bulmus, V.: Modification of RAFT‐ polymers via thiol‐ene reactions: A general route to functional polymers and new architectures. Journal of Polymer Science Part A: Polymer Chemistry 47, 3773–3794. 2009.

Structural Design and Synthesis of Polymer Prodrugs

50 Chytil, P., Etrych, T., Kříž, J., Šubr, V., Ulbrich, K.: N‐(2‐Hydroxypropyl)

51

52 53 54 55

56

57

58

59 60

61

62

methacrylamide‐based polymer conjugates with pH‐controlled activation of doxorubicin for cell‐specific or passive tumour targeting. Synthesis by RAFT polymerisation and physicochemical characterisation. European Journal of Pharmaceutical Sciences 41, 473–482. 2010. Ulbrich, K., Pechar, M., Strohalm, J., Šubr, V., Říhová, B.: Synthesis of biodegradable polymers for controlled drug release. Annals of the New York Academy of Sciences 831, 47–52. 1997. Hoffman, A.S.: The origins and evolution of “controlled” drug delivery systems. Journal of Controlled Release 132, 153–163. 2008. Kopecek, J., Kopeckova, P.: HPMA copolymers: origins, early developments, present, and future. Advanced Drug Delivery Reviews 62, 122–149. 2010. Vicent, M.J., Duncan, R.: Polymer conjugates: nanosized medicines for treating cancer. Trends in Biotechnology 24, 39–47. 2006. Duncan, R., Vicent, M.J.: Do HPMA copolymer conjugates have a future as clinically useful nanomedicines? A critical overview of current status and future opportunities. Adv Drug Deliv Rev 62, 272–282. 2010. Kopeckova, P., Rathi, R., Takada, S., Rihova, B., Berenson, M.M., Kopecek, J.: Bioadhesive N‐(2‐Hydroxypropyl) methacrylamide copolymers for colon‐ specific drug‐delivery. Journal of Controlled Release 28, 211–222. 1994. Etrych, T., Chytil, P., Jelínková, M., Říhová, B., Ulbrich, K.: Synthesis of HPMA copolymers containing doxorubicin bound via a hydrazone linkage. Effect of spacer on drug release and in vitro cytotoxicity. Macromolecular Bioscience 2, 43–52. 2002. Ulbrich, K., Etrych, T., Chytil, P., Jelıń ková, M., Řı́hová, B.: HPMA copolymers with pH‐controlled release of doxorubicin: In vitro cytotoxicity and in vivo antitumor activity. Journal of Controlled Release 87, 33–47. 2003. Ulbrich, K., Subr, V.: Polymeric anticancer drugs with pH‐controlled activation. Adv Drug Deliv Rev 56, 1023–1050. 2004. Etrych, T., Mrkvan, T., Chytil, P., Koňák, Č., Říhová, B., Ulbrich, K.: N‐(2‐ hydroxypropyl)methacrylamide‐based polymer conjugates with pH‐controlled activation of doxorubicin. I. New synthesis, physicochemical characterization and preliminary biological evaluation. Journal of Applied Polymer Science 109, 3050–3061. 2008. Chytil, P., Hoffmann, S., Schindler, L., Kostka, L., Ulbrich, K., Caysa, H., et al.: Dual fluorescent HPMA copolymers for passive tumor targeting with pH‐ sensitive drug release II: Impact of release rate on biodistribution. Journal of Controlled Release 172, 504–512. 2013. Choi, W.M., Kopečková, P., Minko, T., Kopeček, J.: Synthesis of HPMA copolymer containing adriamycin bound via an acid‐labile spacer and its activity toward human ovarian carcinoma cells. Journal of Bioactive and Compatible Polymers 14, 447–456. 1999.

419

420

Polymers for Biomedicine

63 Meng, F., Hennink, W.E., Zhong, Z.: Reduction‐sensitive polymers

64

65

66

67

68 69

70

71

72

and bioconjugates for biomedical applications. Biomaterials 30, 2180–2198. 2009. Shen, W.C., Ryser, H.J.P., Lamanna, L.: Disulfide spacer between methotrexate and poly(D‐Lysine) – a probe for exploring the reductive process in endocytosis. Journal of Biological Chemistry 260, 905–908. 1985. Shiah, J.G., Sun, Y., Kopečková, P., Peterson, C.M., Straight, R.C., Kopeček, J.: Combination chemotherapy and photodynamic therapy of targetable N‐(2‐ hydroxypropyl)methacrylamide copolymer‐doxorubicin/mesochlorin e(6)‐OV‐TL 16 antibody immunoconjugates. Journal of Controlled Release 74, 249–253. 2001. Erez, R., Segal, E., Miller, K., Satchi‐Fainaro, R., Shabat, D.: Enhanced cytotoxicity of a polymer–drug conjugate with triple payload of paclitaxel. Bioorganic & Medicinal Chemistry 17, 4327–4335. 2009. Fox, M.E., Szoka, F.C., Frechet, J.M.: Soluble polymer carriers for the treatment of cancer: the importance of molecular architecture. Accounts of Chemical Research 42, 1141–1151. 2009. Venkatachalam, M.A., Rennke, H.G.: Structural and molecular‐basis of glomerular‐filtration. Circ Res 43, 337–347. 1978. Asgeirsson, D., Venturoli, D., Fries, E., Rippe, B., Rippe, C.: Glomerular sieving of three neutral polysaccharides, polyethylene oxide and bikunin in rat. Effects of molecular size and conformation. Acta Physiol 191, 237–246. 2007. Rennke, H.G., Venkatachalam, M.A.: Glomerular‐permeability of macromolecules – effect of molecular‐configuration on the fractional clearance of uncharged dextran and neutral horseradish‐peroxidase in the rat. Journal of Clinical Investigation 63, 713–717. 1979. Nasongkla, N., Chen, B., MaCaraeg, N., Fox, M.E., Frechet, J.M.J., Szoka, F.C.: Dependence of pharmacokinetics and biodistribution on polymer architecture: effect of cyclic versus linear polymers. Journal of the American Chemical Society 131, 3842‐ 2009. Hoffmann, S., Vystrčilová, L., Ulbrich, K., Etrych, T., Caysa, H., Mueller, T., Mäder, K.: Dual fluorescent HPMA copolymers for passive tumor targeting with pH‐sensitive drug release: synthesis and characterization of distribution and tumor accumulation in mice by noninvasive multispectral optical imaging. Biomacromolecules 13, 652–663. 2012.

421

Part VI Biocompatibilization of Surfaces

423

14 Polymeric Ultrathin Films for Surface Modifications Henning Menzel Institut für Technische Chemie, Technische Universität Braunschweig, Braunschweig, Germany

14.1 ­Methods for Preparation of Ultrathin Polymer Films 14.1.1  Self‐Assembled Monolayers

Although not polymeric, self‐assembled monolayers are of great importance for the preparation of polymeric coatings. They are ultrathin films and serve as anchor layers for polymers or starting points for grafting from polymeriza­ tions. Self‐assembled monolayers are spontaneously formed by the immersion of a substrate in a solution of an active surfactant [1,2] (see Figure 14.1). 14.1.1.1  Thiols on Gold

Thiols and disulfides form well defined and highly ordered monolayers on gold, silver, and copper, yielding e.g. gold(I)thiolates on the surface [1–6]. The  mechanism of the process is not completely understood [1], however, the kinetics of the SAM formation indicates that there is a reversibility in the binding, which allows ordering within the SAM by changes in the binding sites [1, 7, 8]. 14.1.1.2  Silanes on Hydroxylated Surfaces

Chlorosilane‐ or alkoxy groups are hydrolyzed by trace amounts of water ­present on the surface, for example, produced by condensation of two Si‐OH‐ groups at the surface. Actually, Tripp and Hair found that there are no ­monolayers formed when no water is present [9]. The silanol groups can then  condense with other Si‐OH‐groups either present at the surface or on other alkyl silanes. In the latter case, a two‐dimensional polysiloxane network Polymers for Biomedicine: Synthesis, Characterization, and Applications, First Edition. Edited by Carmen Scholz. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

424

Polymers for Biomedicine

Functional group Van der Waals interaction

Alkyl chain

Physi- or chemisorption at the surface Substrate

Head group interacting with the surface

Figure 14.1  Schematic of a self‐assembled monolayer and the molecules necessary for that.

is created with some bonds to the substrate surface [2, 9–11]. The network has enough flexibility so that the alkyl chains form an ordered layer, however, the extent of the order depends on the length of the alkyl chain and the ­temperature [12]. 14.1.1.3  Alkyl Phosphonates on Metal Oxides

Alkyl phosphonates and alkyl phosphonic acids form monolayers on metal oxides like TiO2, TaO2, ZrO2, Al2O3, and others [13–19]. The binding mode involves two or three M‐O‐P bonds and typically also includes a P = O bond, which is lost as double bond upon formation of the SAM [20]. In the case of alkyl phosphonic acid the reaction can be understood as an acid base reac­ tion, while for alkyl phosphonates a hydrolysis of the phosphonic acid ester by the surface bound hydroxyl groups is suggested [20]. The phosphonate monolayers show a significantly higher hydrolytic stability compared to silane monolayers [21]. The mechanism and kinetics resemble more the silane monolayers than the thiol on gold SAM, that is, there is an island growth mechanism [9]. The binding seems to be more or less irreversible and therefore ordering processes, which rely on changes of the binding sites of molecules, are less likely. Therefore, SAMs of alkyl phosphonates are somewhat less ordered compared to thiols on gold. 14.1.1.4  Catechol on Metals, Metal Oxides, or Polymers

Mussel adhesive proteins are remarkable materials that display an extraordi­ nary capability to adhere to many different substrates. They are rich in 3,4‐ dihydroxy‐L‐phenylalanine (DOPA) and the amino acid lysine. It has been shown that the 1,2‐dihydroxybenzene (catechol) group together with primary amino groups is responsible for the adhesion [22]. Dopamine (4‐(2‐amino­ ethyl)benzene‐1,2‐diol) combines the two groups in one molecule and can form stable layers on substrates just by immersing the substrates in an aqueous ­solution of dopamine at pH = 8.5. Under these conditions dopamine autopoly­ merizes to poly(dopamine), which adheres as thin layer on the substrate [23].

Polymeric Ultrathin Films for Surface Modifications

The method can be used on metals like Cu or stainless steel but also noble metals (Au, Ag, Pt, and Pd), oxides (TiO2, amorphous and crystalline SiO2, Al2O3, and synthetic polymers like polystyrene (PS), polyethylene (PE), poly­ carbonate (PC), polyethylene terephthalate (PET), polytetrafluoroethylene, (PTFE), and polydimethylsiloxane (PDMS). Moreover, poly(dopamine) can be further modified by metallization or via attaching additional organic adlayers [23,24], for example, it can be modified so that it bears initiators for a grafting from polymerization (vide infra) [25]. The reactions of the catechols as found in DOPA and dopamine comprise the surface adhesion of the catechol moiety but also oxidation, radical formation, and consecutive reactions (Figure 14.2) [22]. The adhesion is mainly caused by the catechol functionality, while the oxidized o‐quinone functionality is primarily responsible for crosslinking [26]. The mechanism of the adhesion depends mainly on the material used as sub­ strate. Calculations indicate that a dissociative, bidentate structure bridging two Ti‐Atoms (Figure 14.3) is most favorable [27]. Similar interactions can be envisioned for other surfaces bearing hydroxyl groups (Glass, SiO2, or metal oxides), however, for binding to polymers the reactivity of the oxidized and/or radical species (Figure 14.2) might be more important. Copolymers of DOPA and L‐lysine have been prepared and show good adhe­ sion [22,29]. Kuang et al. have developed a tripeptide (Figure 14.4), which can

O

O

O Fe

NH

O (a) HO

(g)

O M

NH

O

O

NH NH2 (e) Oxidation

Imine (b) formation

O

O

O

Surface HO adhesion

O

N

3

Metal chelation

O

O

NH DOPA

NH

(d) Radical generation

DOPA o-quinone NH2

HO NH

O O

NH

HO

(f) Coupling

HO

O

.O NH

Michael adduct (c) formation

O HO NH

Biaryl

N

O

Figure 14.2  Possible hypothetical adhesion, oxidation and crosslinking reaction pathways for peptidyl DOPA and DOPA ortho‐quinone residues. Source: Deming 1999 [22]. Reproduced with permission of Elsevier.

425

426

Polymers for Biomedicine Molecular

Dissociative

R R

O

OH

H O O

O

OH

O

H

Ti

R

Ti

O

O

O

O Ti

R

O O

O O Ti

O

Monodentate

Ti O

O O O Ti

O

Ti O

O

O

O Ti

Bidentate

Figure 14.3  Possible catechol‐TiO2‐interactions. The bridging, dissociative bidentate structure of two Ti‐atoms on the right side is the most favorable according to theoretical calculations [27,28]. Source: Data from Malisova 2010 [28].

Figure 14.4  Tripeptide conjugated with a tertiary bromide, able to polymserize at high pH on metal oxide, metallic, and polymeric substrates to form a thin ATRP initiator layer. Source: Kuang 2012 [30]. Reproduced with permission of American Chemical Society.

NH2

H N

Br O

O N H

H N

O NH2

O OH

OH

OH OH

be used to form a SAM‐like layer bearing an initiator for a grafting from polymerization via ATRP (vide infra) [30]. Self‐assembled monolayers are useful systems to introduce functionalities in  order to tailor the surface of materials for biomedical applications [17, 31–34]. However, SAMs typically have a limited stability and are difficult to  functionalize. Thin polymeric films can have significant advantages over SAMs. 14.1.2  Grafting From

Methods to create thin polymer films should allow to adjust the layer thickness over a wide range and to introduce functional groups. Grafting from polymeri­ zations build a polymer layer atop a surface by a polymerization starting from

Polymeric Ultrathin Films for Surface Modifications

Grafting from

Grafting onto

Surface

Figure 14.5  Schematic of a self‐assembled monolayer and the molecules necessary for that. CN

Me OH

O

Cl Si Me

Me O Si

O CN O

Me

Me O Si Me

Me

O

Me

N

N

N

Me

N

Me

Me

Base

CN

Me

Styrene/toluene

CN

CN O O

Me Immobilized polystyrene

Figure 14.6  Synthesis of monolayers of polystyrene terminally attached to SiOx surfaces by using a self‐assembled monolayer of AIBN like azo compound with a chlorosilane headgroup. Source: Prucker 1998 [35]. Reproduced with permission of American Chemical Society.

a surface‐bound initiator (Figure 14.5). The initiator is typically installed at the surface via a self‐assembled monolayer. Grafting from polymerizations by free radical polymerization were carried out by introducing azogroup based initiators to glass and silicon surfaces. The azogroups decompose and form radicals that then can polymerize vinyl mono­ mers, for example, styrene (Figure 14.6) [35–37]. The advantage of this method is that a high density of grafted polymer chains can be achieved, since the chains are formed at the surface and only small monomer molecules have to diffuse to the surface [35]. At high grafting densities the polymer chains are stretched, which results in very special film properties [38]. For example, a PS

427

428

Polymers for Biomedicine

film placed on a PS brush prepared by grafting from will show dewetting, although the surface bound polymer chains and the chains of the free polymer are chemically identical. However, a mixing of the bound and non‐bound PS molecules at the interface would require a further stretching of the PS mole­ cules, which is entropically very unfavorable [39]. The grafting from method can be used to prepare films for biomedical applications. Rühe et al. studied different methacrylate and acrylamide coatings prepared by grafting from for their adhesion and neurite outgrowth properties [40]. Surface radicals can also be generated by a plasma treatment of the material [41]. Thin polycaprolactone (PCL) films were subjected to an Ar‐plasma ­followed by the grafting of 2‐aminoethyl methacrylate (AEMA) under UV‐ irradiation. The poly(AEMA) film was subsequently used to covalently immobilize gelatin on the surface (see Figure  14.7). Osteosarcoma cells ­ OH O Ar PCL

OH O

O2

Plasma Exposure

OH O

O

O

PCL

Plasma treatment

PCL

Graft polymerization

PCL

Protein covalent immobilization

OH O

PCL

O

O AEMA solution, degassed UV exposure

Gelatin/EDC RT, 4h PCL Gelatin dip coating

PCL

Protein physisorption

Figure 14.7  A schematic representation of the general surface modification strategy presented in ref. 42, consisting of four steps. (1) Plasma treatment, (2) AEMA‐graft polymerization and (3) coating with gelatin by physisorption or by (4) covalent immobilization. Source: Desmet 2010 [42]. Reproduced with permission of John Wiley & Sons.

n

Polymeric Ultrathin Films for Surface Modifications

H N NH2

NH•

NH2 O

Si O O Si

OH OH OH

O

O

O

Ce4+

O

Si O

O

NH

O

NH

O

Si O

O

O

Figure 14.8  Strategy for grafting from by redox polymerization of N‐isopropylacrylamide. Source: Wang 2005 [43]. Reproduced with permission of Elsevier.

­ emonstrated better cell adhesion and cell‐viability on the modified surfaces, d compared to the pure PCL films [42]. Wang et  al. reported a surface initiated redox polymerization [43]. The amino groups of a self‐assembled monolayer of 3‐aminopropyltriethoxysilane (APTES) on a glass surface and Ce4+ ions as redox initiating system were used for surface initiated polymerizations of N‐isopropylacrylamide (see Figure 14.8). 14.1.3  Grafting Onto

Grafting from methods allow for the best control over grafting density, layer thickness, and other parameters, however, they typically include multistep syntheses. In many cases the high level of control is not necessary for the application. Grafting onto methods are typically less laborious and for many applications sufficient in terms of control over the structure of the resulting layer. Grafting onto can be accomplished by two general methods. (i) Employing an anchor layer at the surface, which holds functional groups able to bind a polymer and (ii) using functional polymers which are able to bind to the surfaces. 14.1.3.1  Anchor Layers

Employing anchor layers for the grafting onto has the advantage that polymers can be grafted, which do not react with the surface by themselves. Photochemical grafting methods provide an almost universal approach to bind different poly­ mers to a surface and they are fast and easy to conduct. Additionally, photo­ chemical grafting can be carried out with spatial control to obtain laterally structured surfaces. Among the various different photochemically active groups benzophenone (Figure 14.9) and arylazide are typically used. 14.1.3.1.1  Benzophenone  Upon irradiation with UV‐light benzophenone forms a biradical. This biradical can attack polymers with at least one CH‐ Group and bind them (Figure  14.9) covalently. This method is extremely

429

Polymers for Biomedicine

CH2

CH2

O

HC O

O C

H O

O

O

O Si

O Si

O Si

Substrate

Substrate

Substrate



Figure 14.9  Schematic of the photochemical grafting onto using a benzophenone derivative. Source: Prucker 1999 [44]. Reproduced with permission of American Chemical Society. Figure 14.10  Influence of the chain dimensions, i.e. the bulk radii of gyration, Rg, on the maximum film thickness for photochemically attached polystyrene layers on benzophenone monolayers. Source: Prucker 1999 [44] Reproduced with permission of American Chemical Society.

15

dmax [nm]

430

10

5

0 0

10

20

30

40

50

Rg(bulk) [nm]

versatile and has been used in different concepts for the preparation of ultrathin polymers layers. Prucker et  al. investigated the principles of this method; for example the dependence of the layer thickness on the irradiation time or thickness of the  spin cast layer. They found a linear relation between the polymer chain dimensions (i.e., radius of gyration of the polymer coil) and the layer thickness (Figure 14.10) [44].

Polymeric Ultrathin Films for Surface Modifications

Besides silane based anchor layers [44] phosphonic acid based layers have  also been developed, which readily enable the formation of stable self‐ assembled monolayers on metallic and ceramic surfaces as they are used for implants [45,46]. The potential of this method for biomedical application has been exemplified by Murata et al. showing that cell attachment can be tailored by grafting different preformed polymers to a surface [47]. The method was used by Adden et al. for an intensive screening of different polymers for the adhesion of osteoblast cells [46]. The benzophenone layers cannot only bind preformed polymers, but the radicals formed by the photoreaction can also initiate a polymerization. Thus, benzophenone containing SAMs can also be used for grafting from reactions [48]. 14.1.3.1.2  Arylazide  Arylazide can also be used for grafting polymers onto

surfaces [49,50]. The arylazide groups decomposes under UV‐light irradiation into a nitrene and nitrogen gas. The highly reactive nitrene undergoes ­several reactions as schematically shown in Figure 14.11 and is therefore very universal [51]. Thus, by using azide photochemistry almost every polymer can be grafted onto a surface, and light induced preparation of micro‐structured ­patterns is also possible [52].

14.1.3.1.3  Other Methods  Besides the photochemical grafting onto many other

reactions can be used to tether a preformed polymer to a surface, however, the reactions are typically much more specific and less general. Additionally, they can be much more laborious. An example of a more specialized method

Nitrene :N

N3

Ring expansion

O

O

HN

R–H

HN

Substrate

N2

O Si Substrate

N

NH R R

H N R

R–H Reactive hydrogen

hν O Si

R–NH2

N

N

R

R

R

NH2 N H N

R NH R

Figure 14.11  Schematic of the different possible reaction taking place after the initial photoreaction of aryl azides, R can be a polymer or a side chain of a polymer Source: Lábbe 1969 [51]. Reproduced with permission of American Chemical Society.

431

432

Polymers for Biomedicine

is  the  transformation of a hydroxy‐terminated poly(vinylmethylsiloxane‐ co‐dimethylsiloxane) into a macro chain transfer agent (macro‐CTA) and subsequent RAFT‐polymerization of 2‐methacryloyloxyethyl phosphoryl­ choline  (MPC). Thereby, a triblock copolymer was obtained. This polymer was grafted onto hydrogen‐terminated polysiloxane, which is installed at the surface as “anchor layer,” via the vinyl‐groups along the polysiloxane middle block (Figure 14.12). The grafting of the MPC blocks on the silicone improved wettability and lubrication [53]. 14.1.3.2  Direct Grafting onto a Surface Using Polymer Bound Functional Groups

A direct grafting of polymers to a surface is possible if the polymers bear ­functional groups capable of forming stable bonds with the substrate surface. Chlorosilanes and alkyoxysilanes can react with hydroxyl groups as found on glass or the natural silicon oxide layer of silicon. Therefore, polymers bearing trimethoxysilyl groups can form layers on glass and silicon without the need of  an anchor layer. Huang et  al. prepared block and random copolymers of 3‐trimethoxysilyl propyl methacrylate and dimethylamino ethyl methacrylate (Figure 14.13), as antimicrobial coatings [54]. The trimethoxysilane group is rather sensitive to moisture and the siloxane bond can be hydrolyzed under physiological conditions [21]. Furthermore, although metals like titanium, which are often used as implants, have some hydroxyl groups at the surface due to the natural oxide layers, a pretreatment is necessary to increase their number and obtain homogeneous silane films. Phosphonic acid groups are advantageous as head groups in SAMs on metal oxide layers [21,33] and they have also been used in polymeric systems [55–57]. A suggested mechanism includes the hydrolysis of the phosphonic ester and a reaction of the resulting phosphonic acid group with the surface [16,55]. The grafting onto using such reactive polymers is typically very easy to perform. The polymer is dissolved and coated via common techniques like spray or spin‐coating. The polymer binds to the surface  –  in the case of the phosphonates supported by an annealing step – and the non‐bound polymer is removed in a washing step (Figure 14.14) [55,56]. As already pointed out, catechol groups have been used in bioinspired ­adhesives and can be used for coatings on many substrates [23,24]. An example for a copolymer with catechol anchoring groups is presented in Figure 14.15. The quaternary ammonium side chains endow the coatings with a potent ­antibacterial activity (vide infra), the methoxyethyl side chains are introduced to tune the hydrophobic/hydrophilic balance, and the catechol groups promote immobilization of the polymers [58]. For organic substrates, for example, polymers, the coating requires com­ pletely different groups for the attachment of thin films. For grafting from

O

HO

Si O Si O

Si O l

S

O

Si

OH

m

CN OH

S O

S

CN

O

O

S

Si O Si O l

O

O

CN S O

–O

n O O P O O N+

O O

Si

O

m

O

N+

S

NC O

S O

macro-CTA O– O P O O

O

S

Si O

AiBN

Si O Si O

Si O l

Si m

O

S

NC O O

S n O

O

O O P O– O N+

Figure 14.12  Synthetic route to prepare a Triblock copolymer with a middle block bearing vinylsilane groups which can be used to bind the block‐copolymer to hydrogen terminated polysiloxane. Source: Iwasaki 2007 [53]. Reproduced with permission of Elsevier.

O

stat O

O O

N

stat O

O

Si O O

O

O O

O O

O

stat O

P O O

N

O

O O P

Si O O

OH OH OH

stat O

O

OH OH OH

OH O

O

Figure 14.13  Polymers with trimethoxysilane groups for grafting onto glass and silicon dioxide [54] and polymers with phosphonate groups for grafting onto titanium substrates. Source: Lorenz 2011 [56]. Reproduced with permission of John Wiley & Sons.

e.g. ellipsometry and contact angle measurements Coating of the Titanium with polymer solution

Stored in an oven at 120 °C for 17 h

Washing

Figure 14.14  Coating scheme for polymers with phosphonate groups for grafting onto titanium substrates.

O

OO

OO

N+

stat NH

O

N+

HO OH O

O

OO

O

O

OHOH OH

sta t NH

O OH O

Figure 14.15  Polymers with catechol groups for grafting‐onto on glass. Source: Han 2011 [58]. Reproduced with permission of American Chemical Society.

Polymeric Ultrathin Films for Surface Modifications

n N

O

stat

m O

O

n HN

O

stat

m O

H N

S

NH

O

N3

O N3

Figure 14.16  Polymers with benzophenone [47] or arylazide groups [61] for a photo‐ chemically induced grafting from. Source: Murata 2004 [47] and Ito 1997 [61].

approaches the already mentioned photo‐chemically active benzophenone [47,59] and arylazide groups [60,61] are very versatile, examples for copolymer attachments are depicted in Figure 14.16. While electrostatic interactions are typically too weak for low molecular weight compounds to result in stable films, polymers can indeed form coatings by electrostatic interactions with the substrate. The key are polyions as they carry a multitude of ionic groups. Even if some of the weak electrostatic bonds are broken, the polymer is stat held on the surface. Another strong driving force for n m the adsorption and therefore also contributing to the O O O O film stability is the entropy. Upon adsorption of the polymer to a charged surface, small counter ions are p O released from the substrate surface and from the NH3+ polymer chain. The gain in entropy due the freely p = 8 or 45 diffusing counter ions is much higher than the  entropy loss due the polymer adsorption [62]. Figure 14.17  Copolymer that forms coatings by Therefore, simple polycations (or to a lesser extent electrostatic interaction polyanions, because s­ ubstrate surfaces are typically on negatively charged negatively charged) can be used for the coating substrates and makes the ­ of  substrates. Ionov et  al. have  suggested to use a substrate protein polycationic copolymer ­ composed of aminoethyl resistant. Source: Lonov 2010 [63]. Reproduced methacrylate hydrochloride and oligoethyleneglycol with permission of methacrylate to render a s­urface protein resistant American Chemical Society. (Figure 14.17) [63]. 14.1.4  Layer‐by‐Layer Deposition

The adsorption of a polyion onto a substrate can reverse the surface charge. Subsequently an oppositely charged polyion can be adsorbed to the ­substrate.

435

436

Polymers for Biomedicine

In this way, complex films can be build layer by layer (LbL). This technique has been introduced by Decher et al. [64] and has developed into a versatile tool to prepare nanoscopic assemblies [65]. Not only simple polyions can be used for the LbL‐technique, but also proteins and other bio‐related polymers can be incorporated [65–68].

14.2 ­Examples in Medical Applications In medical applications, the tailoring of surface properties is of particular interest. Many materials used in biomedical applications have been chosen because of their bulk properties. Metals for endoprotheses or dental implants are chosen for their mechanical strength; silicone for cochlear implants has been selected for its softness. In most cases the bulk properties are decisive for the application, but the surface properties have to be taken into account when the implant tissue interaction is considered. Although most implant materials are more or less compatible with the body, they are in most cases bio‐inert. The modification of the surface with a polymer can either improve the biocompat­ ibility or even make the material bioactive. 14.2.1  Improvement of Cell Adhesion

In many cases the aim of an implant modification is to improve the cell adhe­ sion. An early example has been reported by Rühe et al. they used the grafting from method (Figure 14.6) to grow different polymers on glass surfaces and to test them for survival and growth of neurons [40]. Another example for a coat­ ing prepared by a grafting from approach was described by Uchida et al., who coated PS substrates with temperature sensitive poly(N‐isopropyl acrylamide) (polyNIPAM). The grafting from was carried out by spreading a solution of the NIPAM monomer on the PS dish and irradiate it with an electron beam. The irradiation creates radicals at the surface, which start the polymerization. When the lower critical solution temperature of the polyNIPAM is reached the layer changes its hydrophilicity significantly [69]. The layer is very hydrophilic at low temperature and cells do not adhere, however, at 37° the layer becomes much more hydrophobic and turns into a very good cell support. Therefore, cells can be seeded onto a polyNIPAM‐layer at 37°C, grow and form a continu­ ous sheet of cells. However, as soon as the temperature is lowered the cells detach (Figure  14.18) [70,71], a mechanism that has been developed into a very successful approach to grow and harvest cell sheets, for example, for the treatment of heart lesions [72]. Many different polymers can be readily screened using the grafting onto method with benzophenone bearing SAMs as anchor layer. Chang et al. used the method to find polymers that can alter the properties of glutaraldehyde

Polymeric Ultrathin Films for Surface Modifications Extracellular matrices Basement membrane proteins

Cell-cell junction proteins Hydrophobic > 32°C Cell culture

poly(N-isopropylacrylamide)

Lowering culture temperature

Cell culture plate Hydrophilic < 32°C Cell sheet harvest

Figure 14.18  Schematic illustration of cell sheet engineering using temperature sensitive polyNIPAM layers. Source: Matsuura 2014 [72]. Reproduced with permission of Elsevier.

modified porcine heart valves in such a way that vital endothelial cells can grow on the implant surfaces [73]. A similar investigation was carried out by Adden et al. [46] that focused on finding polymer coatings compatible with osteogenic precursor cells. The ease of the method enabled the authors to test 25 very dif­ ferent polymers. The investigation allowed also to check whether the cell growth correlates with physical properties of the polymer layers. For  example, cell growth was plotted as a function of the advancing contact angle of the respec­ tive polymer layers (Figure 14.19). It was found that there is no cell adhesion below a contact angle of 40°. At higher contact angles, some  polymers show good adhesion, while others do not support the cell growth at all. Therefore, a contact angle higher than 40° was a necessary but not a sufficient condition for  a high cell growth score. This evidenced that a single physical parameter cannot predict the behavior of cells on the polymer film [46]. 14.2.2  Binding Growth Factors

In the examples presented so far, the interactions between the polymer and the cells are governed by the chemical and physical properties of the polymer coatings themselves. However, the coatings themselves can be peptides or proteins. The RGD‐peptide sequence (Arg‐Gly‐Asp) is the most common binding motif and cells can bind to it via receptors at the cell surface, the so called integrins. The RGD‐sequence can therefore be used to mediate cell binding to surfaces. A aminopropylsilane monolayer was modified by Porté‐Durrieu et al. with 3‐(maleimido)propionic acid N‐hydroxysuccinimide ester. The maleim­ ide group was subsequently used to bind a peptide via a cysteine residue using the thiol‐ene‐reaction (Figure  14.20). The resulting surfaces present a cyclic RGD peptide and promote osteoplast adhesion [74].

437

1

PMAA

Growth score

PVP

2

PS

PVdeP

3

PEG

PAA PVAcAm

4 PHEAA PVmAcAm

0

PViBE

PHEMA PSSde

PDMAA

20

PVBE PVAc

PMTA

PNVP1/2

PVBP

PAcSt

PEtOx

PPAMA

40

PHySt

60

80

100

θAdv [°]

Figure 14.19  Plot of the growth score for mesenchymal progenitor stem cells (C3H10T1/2) on different polymer layers bound to glass substrates using the grafting onto method with benzophenone bearing SAMs versus the advancing contact angle of the respective polymer layer. The data indicate that a contact angle higher than 45° is a necessary but not a sufficient condition for a high cell growth score. Source: Adden 2007 [46]. Reproduced with permission of Taylor & Francis.

O

HN

O

S

O

O N

O

NH2

NH

Si O O O

O

HN

O

Si O O O

O

O

O

O

O

O SH

O NH

Si O O O

Figure 14.20  Schematic of the activation of an APTES SAM with 3‐(maleimido)propionic acid N‐hydroxysuccinimide ester and subsequent binding of a cyclic peptide with an SH‐group via a thio‐ene‐reaction. Source: Porté‐Durrieu 2004 [74]. Reproduced with permission of Elsevier.

Polymeric Ultrathin Films for Surface Modifications

Signaling proteins, also called growth factors or cytokines, are proteins that control growth and differentiation of cells. If attached to a surface they render it bioactive. Bone morphogenic protein 2 (BMP2) is a growth factor that effects the differentiation of mesenchymal cells into osteoblasts and therefore leads to enhanced local bone growth [75–77]. Early studies of surface immobilized BMP2 showed an improved metal bone contact and proved that bound BMP2 is more efficient compared to soluble BMP‐2 doses [78,79]. BMP2 was initially attached to surfaces by physical adsorption on pretreated Titanium [78,80] or by covalent attachment to, for example, silk fibroin films [79]. A covalent attachment method was developed based on self‐assembled monolayers of alkyl phosphonates that carry carboxyl or hydroxyl moieties as functional end groups. Subsequently, these groups were activated by carbonyl diimidazole for hydroxyl groups and N‐hydroxy succinimide for carboxyl groups (Figure 14.21), respectively. The activated SAMs were then able to bind proteins via amino groups, such as, for example, lysine residues [31]. According to ELISA tests 6  ng/cm2 of surface‐ bound BMP are sufficient to show biological activity [unpublished results]1. While this small amount of BMP2 showed biological activity, the effects might be too weak to effectively improve osseointegration of implants. A ­possible way to attach more possible binding sites on a surface is the use of polymer ­interlayers. Shi et  al. have immobilized dextran on a poly(dopamine) layer. HO O

O N O

HO O

Self assembly

O HO P OH

OH OH TiO2

O

Activation

THF, 120°C, 48 h

P

protein

O O

TiO2

O

Binding the

NHS + EDC

O

HN

O

O

P

O O

O

TiO2

P

O O

TiO2

Figure 14.21  Self‐assembly of 12‐carboxydodecyl phosphonic acid, activation as N‐hydroxysuccimide ester and reaction with amino groups for binding a protein. Source: Adden 2006 [31]. Reproduced with permission of American Chemical Society. 1  The experimental details for the ELISA and the cells test can be found in ref. [56].

439

440

Polymers for Biomedicine

First, dextran was modified by oxidation resulting in aldehyde groups along the polymer chain. These aldehyde groups can react with the amino groups of the poly(dopamine) anchor. Subsequent reduction of the Schiff bases results in a stable attachment [81]. Carboxymethylchitosan was bound to a poly(dopamine) layer via amide bond formation between the amino groups of the poly(dopamine) and the carboxyl groups. BMP2 was subsequently bound via the carboxylic acid functions of the carboxymethylchitosan [82]. Titanium substrates modi­ fied in this way promoted osteoblast spreading, alkaline ­phosphatase activity, and calcium mineral deposition [81,82]. Adden et  al. developed a self‐binding copolymer of (4‐vinylbenzyl)phos­ phonic acid diethylester (VBP) and N‐acryloxysuccinimide (NHS). In this ­copolymer, the VBP moiety allows for grafting onto titanium surfaces in a ­simple coat‐dry‐wash procedure (Figure  14.14). The NHS group is able to react with primary amino groups of proteins, for example, lysine. The VBP‐ NHS‐copolymers were shown to be indeed reactive against primary amino groups [55]. However, the amount of BMP2 bound to these interlayers is small [83]. Therefore, the functional group for the binding of the protein was replaced by glycidyl methacrylate (GMA) [56]. Epoxide groups as present in the glycidyl moiety react readily with nucleophiles such as primary amines, hydroxyl, and sulfhydryl groups and the linkages are very stable [84]. During the film formation process the GMA moieties can undergo some hydrolysis and transesterification, which result in slightly crosslinked films (see Figure 14.22). The film thickness of 60–80 nm is significantly higher than for non‐crosslinked films. Furthermore, the films have a higher hydrophilicity and are believed to form surface bound hydrogels [56]. The increased film thickness and the hydrogel character of the film allowed a much higher loading with BMP2. Via ELISA tests a loading of 100–120 ng/cm2 has been estimated, that is a tenfold increase over the VBP‐NHS‐copolymers [56]. The ELISA detects the BMP2 molecules via selective antibody‐antigen interac­ tions. Therefore, it can be assumed, the detected BMP2 is correctly folded and not denaturized, however, an ELISA does not give any information about the biologi­ cal activity. Lorenz et al. used a luciferase assay (BREluc test) for testing the bio­ logical activity. Compared to the ELISA test, only about 4–8% of the bound BMP2 is biologically active. The authors attributed this to a limited accessibility of the BMP2. While the proteins, like BMP2 or antibodies used in the ELISA, can penetrate the copolymer layer to some extent, the cells used in the BREluc test are too large and retained at the surface. Therefore, only those BMP2 molecules, which are directly present at the surface and accessible in the correct orientation can interact with the cell surface receptors and induce biological activity [56]. 14.2.3  Antimicrobial Coatings

Implant related infections are a severe problem in medicine as bacteria can form biofilms on non‐natural implant surfaces. Within these biofilms, the ­bacteria are embedded in an organic matrix of extracellular polysaccharides

t ou g ith in W slink s cro

Titanium

Titanium

Drying and binding

After washing

Titanium

Titanium

Titanium

cro W ss ith lin kin g

Figure 14.22  Schematic presentation of the process of film production: films directly after coating and for a crosslinked and a non‐crosslinked polymer film. Source: Lorenz 2011 [56]. Reproduced with permission of John Wiley & Sons.

442

Polymers for Biomedicine

and are protected from systemic antibiotics. Such an infection and biofilm formation typically cannot be treated conservatively, but removal of the ­ implant becomes necessary. A modification of the implant surface to make it antibiotic without compromising the cell adhesion is therefore highly desirable. First examples of antibiotic polymer coatings have been published already in 1972 [85,86]. The polymer layer is created by hydrolysis and condensation of a triethoxysilane equipped with a quaternary ammonium group bearing a long alkyl chain. The method is very similar to the formation of a silane SAM, but here the silane is intentionally hydrolyzed. This hydrolysis yields highly reac­ tive silanol groups, which condense forming not only contacts with the surface but mostly a two‐dimensional polymer network. Thus, a polysiloxane network is formed, which has some chemical bonds with the surface (Figure 14.23). The coating is clearly antibacterial and antifungal but was not expected to be cell compatible, although this aspect was not investigated [85,86]. Some excellent reviews about the further development of cationic polymers as antibacterial coating are available [87,88] and their basic ideas shall be exem­ plified here. Kurt et al. have implemented a polymer surface modifier concept

N+

N+

N+

3 H2O O Si O O

–3 CH3OH

HO Si OH OH OHOH OH

N+

–3 H2O Si

O Si O

O OH O

Figure 14.23  Hydrolysis and condensation of 3‐(trimethoxysilyl)‐propyldimethyloctadecyl ammonium chloride with surfaces containing reactive functional groups. Source: Adapted from Isquith 1972 [85].

Polymeric Ultrathin Films for Surface Modifications

that relies on specially designed copolymers, which are blended with the polymer to be modified and show a thermodynamically driven surface ­ ­enrichment [89] (Figure 14.24). As an implementation of this concept a polyu­ rethane was synthesized, in which the soft block has a surface‐active moiety, a  trifluoromethyl group (A in Figure  14.24) and an antibacterial long chain alkyl ammonium group (B in Figure 14.24). In a polymer blend this polyure­ thane surface modifier is enriched at the surface and presents the antibacterial alkyl ammonium groups [89]. Synthetic polypeptides (i.e., homo‐, statistical, and block copolymers of one or a few amino acids) can have antibacterial properties. Poly‐L‐lysine as a cati­ onic polymer is well known for its antibacterial properties, [90,91] but it is not well compatible with cells. Some et  al. have used poly‐L‐lysine adsorbed on graphene oxide to produce composites that indeed show antibacterial activity and good cell compatibility [92]. A covalent attachment promises a better control of the antibacterial polymer layer on top of a surface compared to a simple adsorption process. Tiller et al. bound cationic poly(N‐alkyl vinylpyridinium) to a self‐assembled monolayer of aminopropyltriethoxy silane, (see Figure 14.25) [93], which showed a strong antibacterial effect. However, the cell compatibility was not checked in this study, but can be expected to be low. The same group also used a slightly dif­ ferent method to attach polyvinylpyridinium chains at a surface. A SAM of aminopropyltriethoxy silane provides amino groups at the surface, which were reacted with acryloyl chloride, resulting in a SAM with acrylamide moieties at the surface. Polymerization of vinyl pyridine at the surface resulted in binding of the polymer chains by incorporation (copolymerization) of the acrylamide moieties (“grafting through”) [94]. The poly(vinylpyridine) layer was subse­ quently quaternized with alkyl bromide of different alkyl chain lengths, and the influence of the alkyl chain length on the antimicrobial effectivity was studied. The results indicate, that short and medium length alkyl chains (C3–C6) are more effective then long alkyl chains (C10–C16) [94]. This effect might be due to the different structure of the alkyl chains at the surface. Sum frequency generation vibrational spectroscopy on poly(N‐alkylvinylpyridinium) gave ­evidence that the longer alkyl chains promote the disorientation of the alkyl groups [95]. The mechanism of the antibacterial effect on the surface coatings is still a matter of debate. Almost all antibacterial polymer surface modifications are cationic and include some hydrophobic units. These structural properties are the same as found in antimicrobial peptides (AMP), which have been used as natural role model for the design of antibacterial polymers [96–99]. The common characteristics of many AMPs are cationic and hydrophobic groups clustered in domains, resulting in an “amphiphatic” nature [100]. The cationic regions of the AMPs are believed to interact with anionic lipids on bacterial cell membranes. This interaction destroys the integrity of the bacterial cell

443

Polymer surface modifier A B

Bulk polymer C12H25 N+ A=

O

O N H

=B

F3C

O

H N

O O

H N

O O

O O 0,89

O 0,11 O

H N

O

H N

O O

O

N H

Figure 14.24  Polymer surface modifier (PSM) concept where A and B are complementary side chains that generate an amphiphilic, biomimetic soft block, and the implementation as polyurethane with a soft block having trifluoromethyl and dimethylbutyldodecyl ammonium as A and B side chains. Source: Kurt 2007 [89]. Reproduced with permission of American Chemical Society.

Polymeric Ultrathin Films for Surface Modifications

Br

Br– N+

N Br Br

N+

Br –

N

N+

Br

Br –

n

NH2

Si O O O

NH Si O O O

NH Si O O O

N+ Br –

NH Si O O O

Figure 14.25  Schematic of the derivatization of a surface with cationic poly(n‐hexyl vinylpyridinium) polymers via an aminopropyltriethoxysilane monolayer. Source: Tiller 2002 [93]. Reproduced with permission of John Wiley & Sons.

membrane, for which several mechanisms have been devised (Figure  14.26) [101]: (i) the formation of pores by integration of the AMP via the hydrophobic regions (“Barrel–Stave” model), (ii) the disintegration of the membrane in the so called “carpet” model [102], (iii) the aggregation of anionic lipids, which results in a reorganization of the membrane and therefore in defects [103], and (iv) an ion exchange between the positive charges of the AMPs and the structurally critical mobile cations (Ca2+and Mg2+) at the bacterial membranes [104]. The first two mechanisms require that the hydrophobic regions of the AMPs or, in the case of antimicrobial polymers, the quaternary polymer should be able to penetrate the cell walls causing damage to the membrane [94]. Moreover, there are reports that give evidence for an alkyl chain insertion, but on the other hand, there are also indications that the exchange of cations, or the aggregation of lipids due to electrostatic interactions are the prominent mechanisms. For example the antibacterial activity of poly(alkyloxazoline) telechelics with one quaternary N,N‐dimethyldodecyl ammonium (DDA) end group in solu­ tion was found to be greatly controlled by the other end group (satellite group). The proposed mechanism for the influence of the satellite group suggests an aggregation of the hydrophobic end groups in unimolecular micelles, which results in an insertion of the alkyl chains in the cell membrane in close proxim­ ity, which in turn causes a stronger disturbance of the cell membrane integrity (Figure 14.27) [105]. In the case of the cation exchange mechanism it is expected that the charge density plays an important role. Kügler et al. have prepared surfaces equipped with poly(N‐alkylvinylpyridinium) layers by two different methods. They used a grafting from approach according to Biesalski et al. [106] to create a surface

445

Antimicrobial peptid/polymer „Barrel-Stave“- model Aggregation of anionic lipids

Mg2+

Bacterial cell membrane „Carpet“- model

Mg2+

Mg2+

Mg2+

Exchange of structure stabilizing cations

Figure 14.26  Schematic of some mechanism proposed for the attack of antimicrobial peptides or polymers on bacterial membranes.

Polymeric Ultrathin Films for Surface Modifications

(a)

(b)

+-N-

+-N-

+-N-

-N+

Figure 14.27  Schematic illustration of the proposed actions of telechelic antimicrobial polymers with satellite function (alkyl–PMOX–DDA) binding to a phospholipid membrane. (a) no aggregation of the end groups and insertion in different positions of the membrane; (b) aggregation of the end groups and insertion in close proximity. Source: Waschinski 2005 [105]. Reproduced with permission of John Wiley & Sons.

with a high density of cationic charges and the method of Tiller et  al. [94], which results in a lower charge density. Thereby, the charge density within the organic layer was varied between 1012 and 1016 positive charges per cm2. Inoculation experiments with different bacteria strains indicate that bacterial death occurs above a threshold value for the surface charge density, which depends on the bacterial type and the growth state of the bacteria [104]. A similar investigation by Huang et al. [54] used block copolymers and random copolymers from 2‐(dimethylamino)ethyl methacrylate and 3‐(trimethoxysi­ lyl)propyl methacrylate and grafted them onto glass substrates. Depending on the block length or random distribution, respectively, the resulting polymer layers show different charge densities at the surface. Antimicrobial activity of the surfaces against E. coli increased with the density of available quaternary amino groups [54]. Both, the “insertion” and the two “electrostatic interactions” hypotheses can explain the mechanism of antimicrobial polymers on surfaces. However, fac­ tors such as the effect of the chain structure, different concentrations and

447

448

Polymers for Biomedicine

molecular weights as well as the different structure in solution and at a surface have to be taken into account; actual structure–property relations are not fully understood. Therefore, in the search for new and effective antibacterial coatings typically a screening is necessary. A very efficient high throughput screening was developed based on acrylates with different side chains. Several different acrylates were mixed in different compositions (>500 different ­mixtures) and printed as 300 µm dots and photo‐polymerized [107]. This first‐ generation microarray was incubated with fluorescent bacteria, to investigate which of the coatings can prevent bacterial adhesion. Antimicrobial coatings were further developed by Zou et al. who prepared coatings using a grafting onto approach with a benzophenone SAM as anchor layer (Figure  14.9) to bind a potent antimicrobial polymer on the basis of poly(oxonorbornene) having cationic amino groups. The polymer at the sur­ face was crosslinked via a thiol‐ene reaction with a star shaped polyethylene glycol bearing thiol endgroups. Onto this network a layer of a zwitterionic poly(oxynorbonene) was bound via some of the amino groups. The latter layer was installed to prevent protein adsorption which could hamper the effect of the antimicrobial polymer [108]. Moreover, coatings were deposited in an easier less laborious way; the ­copolymerization of N‐hexylvinylpyridinium and phosphonate groups yielded self‐binding antimicrobial copolymers (Figure 14.28a) [57]. Depending on the exact composition, these copolymers show selectivity: Bacteria are effectively hindered to adhere to the surface, while cells do adhere [57]. Another self‐binding antibacterial copolymer was prepared by reacting poly(vinylbenyzl chloride) with N,N‐dimethylaminopropyltrimethoxy silane and N,N‐dimethylbutyl amine (Figure 14.24b) [109]. The copolymer enabled a simple, inexpensive and widely applicable preparation of bactericidal surfaces starting from hydroxylated materials (glass, cellulose, etc.). However, these copolymers are not selective, and toxic to cells. A similar approach was published by Shamby et  al. who quaternized poly(vinylpyridine) with bromopropyltrimethoxysilane and an alkyl bromide to obtain a copolymer which can bind to hydroxylated surfaces (Figure 14.28c) [110]. If the quaternization is not complete, pyridine groups remain in the polymer, which can be used to complex silver ions. Reduction of the silver ions in the film yields silver nanoparticles, which also have antibacterial effects. Thus, a dual action antibacterial material is obtained. The cationic polymer kills bacteria on contact, while the silver ions leaching from the nanoparticles kill bacteria in close proximity [111]. Catechol groups can also be used to create self‐binding polymers. Han et al. prepared a terpolymer of dopamine methacrylamide for binding, methoxyethyl acrylate to adjust the hydrophilic/hydrophobic balance and dimethylaminoe­ thyl methacrylate quaternized with dodecyl bromide as antibacterial groups (Figure 14.15). The copolymer binds to the surface via catechol groups and the

Polymeric Ultrathin Films for Surface Modifications

(a)

O

(b) N

stat x O

+

N

n

y

stat x

O Si O O

N+ – Br

O P O O

N+ Cl–

Cl

y

N+

Cl–

O Si O O

(c) Br n N

Br

stat x

+ O Si O O

N

stat y

y N+ Br–

N+

O

Br–

O Si O

Figure 14.28  Self‐binding antibacterial copolymers (a) prepared by copolymerization of DMMEP and VP and subsequent quaternization with hexyl bromide. Source: Pfaffenroth 2011 [57]. Reproduced with permission of John Wiley & Sons. (b) By quaternization of poly(vinylbenzylchloride). Source: Bouloussa 2008 [109]. Reproduced with permission of Royal Society of Chemistry. And (c) by quaternization of poly(vinylpyridine) Source: Sambhy 2008 [110]. Reproduced with permission of American Chemical Society.

dodecyl groups are oriented away from the surface, as evidenced by sum ­frequency generation spectroscopy [58]. The coatings show very good antibac­ terial effects; however, cell adherence was not tested. Carboxymethylated chitosan was grafted onto titanium surfaces pretreated with a poly(dopamine) layer. The surfaces modified in this way were not only able to bind BMP2, but also showed antibacterial properties, while promoting attachment, alkaline phosphatase activity, and calcium mineral deposition of both osteoblast and human bone marrow‐derived mesenchymal stem cells [82]. A very general problem of antibacterial coatings on the basis of thin poly­ mer films is the adsorption of proteins that can mask the active coating [112,113]. Therefore, the antibacterial effect should be favorably combined with an antifouling/releasing function of the coating. Such a system can be obtained by installing cationic N,N‐dimethyl‐2‐morpholinone (CB‐ring) at the surface, by ATRP of the corresponding methacrylate (Figure  14.29)

449

450

Polymers for Biomedicine Repell Release Kill O N+ O

O

N+

Hydrolysis

OH

Regeneration O

O

O O

O O

SI-ATRP

O Dry surface

O–

N+

O Wet surface

Figure 14.29  A smart polymer coating repeatedly switches between the attacking function (CB‐Ring, to kill bacteria under dry conditions) and defending function (CB‐OH, to release and resist bacteria under wet conditions). CB‐Ring can be hydrolyzed to CB‐OH in neutral or basic aqueous solutions and can be regenerated by dipping CB‐OH in acidic media. Source: Cao 2012 [115]. Reproduced with Permission of John Wiley & Sons.

initiated by a thiol SAM on gold. The CB‐ring is slowly hydrolyzed into the corresponding zwitterionic carboxy betaine (CB‐OH) [114]. While the CB‐ ring kills bacteria on contact, the zwitterionic form is very hydrophilic and represents an antifouling surface. Therefore, any dead bacteria or bacterial proteins are removed from the surface, when it is in the zwitterionic state. The  original CB‐ring structure can be regenerated by acid treatment [115]. Lienkamp et  al. combined antifouling and antibacterial functionality within a  polymer coating in order to prevent masking of the coating by adsorbed proteins and dead cells [108].

14.3 ­Summary Ultrathin films can be used for several biomedical applications. They can be prepared by different methods. In many cases, self‐assembled monolayers are  starting points or anchor layers for binding the active components or a  polymer layer, for example, by a photochemical process (grafting onto). Self‐assembled monolayers are important as initiators for surface initiated ­polymerizations (grafting from). Several examples for grafting onto and graft­ ing from approaches have been presented and their advantages and disadvan­ tages were discussed. Several examples of biomedical applications of ultrathin films for improved cell adhesion, cytokine delivery, and as antibacterial ­coatings have been presented.

Polymeric Ultrathin Films for Surface Modifications

­References 1 A. Ulman, An Introduction to Ultrathin Organic Films – From Langmuir-

Blodgett to Self-Assembly Academic Press, San Diego (1991).

2 A. Ulman, Chem. Rev. 96, 1533 (1996). 3 M. D. Porter, T. B. Bright, D. L. Allara, C. E. D. Chidsey, J. Am. Chem. Soc. 109,

3559 (1987).

4 R. G. Nuzzo, D. L. Allara, J. Am. Chem. Soc. 105, 4481 (1983). 5 H. Sellers, A. Ulman, Y. Shnidman, J. E. Eilers, J. Am. Chem. Soc. 115,

9389 (1993).

6 C. A. Widrig, C. Chung, M. D. Porter, J. Electroanal. Chem. 310, 335–339

(1991).

7 K. A. Peterlinz, R. Georgiadis, Langmuir 12, 4731–4740 (1996). 8 N. Camilone III, Langmuir 20, 1199–1206 (2004). 9 C. P. Tripp, M. L. Hair, Langmuir 11, 1215 (1995). 10 A. Ulman, Adv. Mater. 2, 573 (1990). 11 D. L. Allara, R. G. Nuzzo, Langmuir 1, 45–52 (1985). 12 J. B. Brzoska, I. B. Azouz, F. Rondelez, Langmuir 10, 4367 (1994). 13 J. T. Woodward, A. Ulman, D. K. Schwarz, Langmuir 12, 3626 (1996). 14 E. S. Gawalt, M. J. Avaltoni, N. Koch, J. Schwartz, Langmuir 17, 5736–5738 (2001). 15 W. Gao, L. Dickinson, C. Grozinger, F. G. Morin, L. Reven, Langmuir 12,

6429 (1996).

16 R. Frantz, J.‐O. Durand, M. Granier, G. F. Lanneau, Tetrahedron Lett. 45,

2935 (2004).

17 C. Viornery, Y. Chevolot, D. Léonard, O. Aronsson, P. Péchy, H. J. Mathieu, P.

Descouts, M. Grätzel, Langmuir 18, 2582 (2002).

18 R. Helmy, A. Y. Fadeev, Langmuir 18, 8924 (2002). 19 G. Zorn, I. Gotman, E. Y. Gutmanas, R. Adadi, C. N. Sukenik, Chem. Mater. 20 21 22 23 24 25 26 27 28 29 30

17, 4218–4226 (2005). G. Guerrero, P. H. Mutin, A. Vioux, Chem. Mater. 13, 4367 (2001). B. M. Silvermann, K. A. Wieghaus, J. Schwartz, Langmuir 21, 225 (2005). T. J. Deming, Curr. Opin. Chem. Biol. 3, 100–105 (1999). H. Lee, S. M. Dellatore, W. M. Miller, P. B. Messersmith, Science 318, 426–430 (2007). H. Lee, B. P. Lee, P. B. Messersmith, Nature 448, 338–341 (2007). X. Fan, L. Lin, J. L. Dalsin, P. B. Messersmith, J. Am. Chem. Soc. 127, 15843–15847 (2005). M. Yu, J. Hwang, T. J. Deming, J. Am. Chem. Soc. 121, 5825–5826 (1999). P. C. Redfern, P. Zapol, L. A. Curtiss, M. C. Thurnauer, J. Phys. Chem. B 107, 11419–11427 (2003). B. Malisova, Dissertation, ETH Zürich, 2010. M. Yu, T. J. Deming, Macromolecules 31, 4739–4745 (1998). J. Kuang, P. B. Messersmith, Langmuir 28, 7258–7266 (2012).

451

452

Polymers for Biomedicine

31 N. Adden, L. J. Gamble, D. G. Castner, A. Hoffmann, G. Gross, H. Menzel,

Langmuir 22, 8197–8204 (2006).

32 S. Pallu, C. Bourget, R. Bareille, C. Labrugere, M. Dard, A. Sewing, et al.,

Biomaterials 26, 6932 (2005).

33 J. Schwartz, M. J. Avaltoni, M. P. Danahy, B. M. Silvermann, E. L. Hanson,

J. E. Schwarzbauer,et al., Mater. Sci. Eng. C 23, 395 (2003).

34 E. S. Gawalt, M. J. Avaltoni, M. P. Danahy, B. M. Silvermann, E. L. Hanson,

K. S. Midwood, et al., Langmuir 19, 200 (2003).

35 O. Prucker, J. Rühe, Langmuir 14, 6893 (1998). 36 O. Prucker, M. Schimmel, G. Tovar, W. Knoll, J. Rühe, Adv. Mater. 14,

1073–1077 (1998).

37 R. Konradi, J. Rühe, Langmuir 22, 8571–8575 (2006). 38 J. Rühe, in Polymer Brushes, edited by R. Advincula, W. J. Brittain, K. C. Caster

et al. (Wiley‐VCH, Weinheim, 2004), pp. 1–31.

39 G. Reiter, P. Auroy, L. Auvray, Macromolecules 29, 2150–2157 (1996). 40 J. Rühe, R. Yano, J. S. Lee, P. Köberle, W. Knoll, A. Offenhäuser, J. Biomater.

Sci. Polymer Edn. 10, 859 (1999).

41 T. Desmet, R. Morent, N. De Geyter, C. Leys, E. Schacht, P. Dubruel,

Biomacromolecules 10, 2351–2378 (2009).

42 T. Desmet, T. Billiet, E. Berneel, R. Cornelissen, D. Schaubroeck, E. Schacht, P.

Dubruel, P Macromol. Biosci. 10, 1484–1494 (2010).

43 Y. P. Wang, K. Yuan, Q. L. Li, L. P. Wang, S. J. Gu, X. W. Pei, Mater. Lett. 59,

1736–1740 (2005).

44 O. Prucker, C. A. Naumann, J. Rühe, W. Knoll, C. W. Franck, J. Am. Chem. Soc.

121, 8766 (1999).

45 J. Pahnke, J. Rühe, Macromol. Rapid Commun. 25, 1396 (2004). 46 N. Adden, A. Hoffmann, G. Gross, H. Windhagen, F. Thorey, H. Menzel,

J. Biomater. Sci. Polymer Edn. 18, 303–316 (2007).

7 H. Murata, B. J. Chang, O. Prucker, M. Dahm, J. Rühe, Surf. Sci. 570, 111 (2004). 4 48 T. Goda, R. Matsuno, T. Konno, M. Takai, K. Ishihara, Coll. Surf. B:

Biointerfaces 63, 64–72 (2008). T. Miyasaki, Y. Machawa, K. Koyama, Thin Solid Films 180, 73 (1989). M. A. Bartlett, M. Yan, Adv. Mater. 13, 1449 (2001). G. L´Abbe, D Chem. Rev. 69, 345–363 (1969). D. J. Pritchard, H. Morgan, J. M. Cooper, Angew. Chem. Int. Ed. 34, 91 (1995). Y. Iwasaki, M. Takamiya, R. Iwata, S. Yusa, K. Akiyoshi, Coll. Surf. B 57, 226–236 (2007). 54 J. Huang, R. R. Koepsel, H. Murata, H. Wu, S. B. Lee, T. Kowalewski, et al., Langmuir 24, 6785–6975 (2008). 5 N. Adden, L. J. Gamble, D. G. Castner, A. Hoffmann, G. Gross, H. Menzel, 5 Biomacromolecules 7, 2552 ‐2559 (2006). 6 C. Lorenz, A. Hoffmann, G. Gross, H. Windhagen, P. Dellinger, K. Möhwald, 5 et al., Macromol. Biosci. 11, 234–244 (2011). 49 50 51 52 53

Polymeric Ultrathin Films for Surface Modifications

57 C. Pfaffenroth, A. Winkel, W. Dempwolf, L. J. Gamble, D. G. Castner, M.

Stiesch, H. Menzel, Macromol. Biosci. 11, 1515–1525 (2011).

58 H. Han, J. Wu, C. A. Avery, M. Mizutani, X. Jiang, M. Kamigaito, et al., 59 60 61 62

63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81

Langmuir 27, 4010–4019 (2011). R. Toomey, D. Freidank, J. Rühe, Macromolecules 37, 882 (2004). N. Gomez, Y. Lu, S. Chen, C. E. Schmidt, Biomaterials 28, 271–284 (2007). Y. Ito, G. Chen, Y. Guan, Y. Imanishi, Langmuir 13, 2756 (1997). V. Kabanov, in Multilayer Thin Films ‐ Sequential assembly of Nanocomposite Materials, edited by G. Decher and J. B. Schlenoff (Wiley‐VCH, Weinheim, 2003), pp. 47–86. L. Ionov, A. Synytska, E. Kaul, S. Diez, Biomacromolecules 11, 233–237 (2010). G. Decher, J. D. Hong, J. Schmitt, Thin Solid Films 210–211, 831 (1992). G. Decher, J. B. Schlenoff, Multilayer Thin Films ‐ Sequential assembly of Nanocomposite Materials. (Wiley‐VCH, Weinheim, 2003). T. Crouzier, K. Ren, C. Nicolas, C. Roy, C. Picart, Small 5, 598–608 (2009). A. N. Zelikin, ACS Nano 4, 2494–2509 (2010). G. Decher, in Multilayer Thin Films, edited by G. Decher and J. B. Schlenoff (Wiley‐VCH, Weinheim, 2003), pp. 1–46. M. Heskins, J. E. Guillet, J. Macromol. Sci., Chem. 2, 1441 (1968). N. Yamada, T. Okano, H. Sakai, F. Karikusa, Y. Sawazaki, Y. Sakurai, Makromol. Chem., Rapid Commun. 11, 571–576 (1990). K. Uchida, K. Sakai, O. H. Kwon, E. Ito, T. Aoyagi, A. Kikuchi, et al., Macromol. Rapid Commun. 21, 169 (2000). K. Matsuura, R. Utoh, K. Nagase, T. Okano, J. Controlled Release 190, 228–239 (2014). B. J. Chang, O. Prucker, E. Groh, A. Wallrath, M. Dahm, J. Rühe, Coll. Surf. A 198, 519 (2002). M. C. Porté‐Durrieu, F. Guillemot, S. Pallu, C. Labrugere, B. Brouillaud, R. Bareille, et al., Biomaterials 25, 4837 (2004). K. Bessho, T. Tagawa, M. Murata, Clinical Orthopaedics Related Res. 268, 226 (1991). J. M. Wozney, in Cellular and Molecular Biology of Bone, edited by M. Noda (Academic Press, New York, 1993), pp. 3–22. A. Hoffmann, H. A. Weich, G. Gross, G. Hillmann, Appl. Microbiol. Biotechnol. 57, 294–308 (2001). G. Voggenreiter, K. Hartl, S. Assenmacher, M. Chatzinikolaidou, H. M. Rumpf, H. P. Jennissen, Mat. Wiss. Werkstofftech. 32, 942–948 (2001). V. Karageorgiou, L. Meinel, S. Hofmann, A. Malhota, V. Volloch, D. Kaplan, J. Biomed. Mater. Res. 71A, 528 (2004). H. P. Jennissen, T. Zumbrink, M. Chatzinikolaidou, J. Steppuhn, Mat. Wiss. Werkstofftech. 30, 838–845 (1999). Z. Shi, K. G. Neoh, E. T. Kang, C. K. Poh, W. Wang, Tissue Eng. A 15, 417–426 (2009).

453

454

Polymers for Biomedicine

82 Z. Shi, K. G. Neoh, E. T. Kang, C. K. Poh, W. Wang, Biomacromolecules 10,

1603–1611 (2009).

83 H. Menzel, A. Hoffmann, G. Gross, W. Dempwolf, F. Thorey, H. Windhagen,

et al., BIOMaterialien 9, 130 (2008).

84 G. T. Hermanson, Bioconjugate Techniques. (Academic Press, San Diego,

1996).

85 A. J. Isquith, E. A. Abbot, P. A. Walters, Appl. Microbiol. 24, 859–863 (1972). 86 A. J. Isquith, C. J. McCollum, Appl. Environ. Microbiol. 36, 700–704 (1978). 87 L. Ferrreira, A. Zumbuehl, J. Mater. Chem. 19, 7796–7806 (2009). 88 J. C. Tiller, Adv. Polym. Sci. 240, 193–217 (2011). 89 P. Kurt, L. Wood, D. E. Ohmann, K. J. Wynne, Langmuir 23, 4719–4723 (2007). 90 S. Shima, H. Matsuoka, T. Iwamoto, H. Sakai, J. Antibiot. 37, 1449–1455 (1984). 91 T. J. Deming, Prog. Polym. Sci. 32, 858–875 (2007). 92 S. Some, S.‐M. Ho, P. Dua, E. Hwang, Y. H. Shin, H. J. Yoo, et al., ACS Nano

6, 7151–7161 (2012).

93 J. C. Tiller, B. H. Lee, K. Lewis, A. M. Klibanov, Biotech. Bioeng. 79, 465–471

(2002).

94 J. C. Tiller, C. J. Liao, K. Lewis, A. M. Klibanov, Proc. Nat. Acad. Sci. USA 98,

5981 (2001).

95 A. M. Oliveira, P. B. Miranda, D. F. S. Petri, J. Phys. Chem. C 116, 18284–

18291 (2012).

96 G. N. Tew, D. Liu, B. Chen, R. J. Doerksen, J. Kaplan, P. J. Carroll, et al. Proc.

Nat. Acad. Sci. USA 99, 5110 (2002).

97 L. Arnt, K. Nüsslein, G. N. Tew, J. Polym. Sci. A., Polym. Chem. Ed. 42,

3860 (2004).

98 G. J. Gabriel, A. E. Madkour, J. M. Dabkowski, C. F. Nelson, K. Nüsslein,

G. N. Tew, Biomacromolecules 9, 2980–2983 (2008).

99 K. Lienkamp, A. E. Madkour, G. N. Tew, Adv. Polym. Sci. 251, 141–173 100 101 102 103 104 105 106 107 108 109 110

(2013). M. Zasloff, Nature 415, 389–395 (2002). L. T. Nguyen, E. F. Haney, H. J. Vogel, Trends Biotechnol. 9, 464–472 (2011). Y. Shai, Biopolymers 66, 236–248 (2002). R. M. Epand, R. F. Epand, Biochim. Biophys. Acta 1788, 289–294 (2009). R. Kügler, O. Bouloussa, F. Rondelez, Microbiology 151, 1341–1348 (2005). C. J. Waschinski, V. Herdes, F. Schueler, J. C. Tiller, I Macromol. Biosci. 5, 149 (2005). M. Biesalski, J. Rühe, Macromolecules 32, 2309 (1999). A. L. Hook, C.‐Y. Chang, J. Yang, J. Luckett, A. Cockayne, S. Atkinson, et al., Nature Biotechn. 30, 868–875 (2012). P. Zou, W. Hartleb, K. Lienkamp, J. Mater. Chem. 22, 19579–19589 (2012). O. Bouloussa, F. Rondelez, V. Semety, Chem. Commun. 10.1039/b716026g, 951–953 (2008). V. Sambhy, B. R. Peterson, A. Sen, Langmuir 24, 7549–7558 (2008).

Polymeric Ultrathin Films for Surface Modifications

111 V. Sambhy, M. M. MacBride, B. R. Peterson, A. Sen, J. Am. Chem. Soc. 128,

9798–9808 (2006).

112 H. Murata, R. R. Koepsel, K. Matyjaszewski, A. J. Russell, Biomaterials 28,

4870–4879 (2007).

113 R. Müller, A. Eidt, K.‐A. Hiller, V. Katzur, M. Subat, H. Schweikl, et al.,

Biomaterials 30, 4921–4929 (2009).

114 Z. Cao, N. Brault, H. Xue, A. Keefe, S. Jiang, Angew. Chem. Int. Ed. 50,

6102–6104 (2011).

115 Z. Cao, L. Mi, J. Mendiola, J.‐R. Ella‐Menye, L. Zhang, H. Xue, S. Jiang,

Angew. Chem. Int. Ed. 51, 2602–2605 (2012).

455

457

15 Surface Functionalization of Biomaterials by Poly(2‐oxazoline)s Giulia Morgese and Edmondo M. Benetti Laboratory for Surface Science and Technology, Department of Materials, ETH Zürich, Zürich, Switzerland

15.1 ­Introduction Surface modification of biomaterials by grafting functional polymers has recently attracted increasing interest, most of all in the development of implants, scaffolds for tissue engineering, drug‐delivery, bio‐imaging systems, and biosensors [1–4]. In all these applications, controlling the interaction between the exposed surface of the biomaterial and the surrounding biological medium has become of paramount importance to determine the performance of the entire device. The formation of dense assemblies of surface‐grafted hydrophilic polymers hinders any unspecific interaction with proteins, ­bacteria  or cells [5,6], and further allows the subsequent introduction of ­chemical functions or biomolecules that can trigger a specific and independent bio‐response [7]. This includes, the immobilization of peptides or proteins at the polymer film [8–13], which can stimulate stem cells to adhere, migrate or differentiate towards a particular tissue type [14,15]. Alternatively, in the case of antimicrobial polymer films, several biocides can be coupled to the polymer layer in order to provide not only an antifouling character but also bio‐active properties [16–19]. Antifouling polymer surfaces can be produced by either “grafting‐to” or “grafting‐from” methods (Figure  15.1). In the former case, polymer chains featuring reactive functions towards the target surface are deposited by ­ wet  deposition processes, to form polymer monolayers (Figure  15.1a) [20]. By contrast, grafting‐from techniques encompass a surface‐confined polym­ erization process, which generates grafted polymer layers from an initiator‐ functionalized surface (Figure 15.1b) [21,22]. Both these methods yield dense and stretched polymer grafts assuming a “brush” morphology [23,24]. Polymers for Biomedicine: Synthesis, Characterization, and Applications, First Edition. Edited by Carmen Scholz. © 2017 John Wiley & Sons, Inc. Published 2017 by John Wiley & Sons, Inc.

458

Polymers for Biomedicine

(a)

(b)

Figure 15.1  Surface‐functionalization by grafting‐to (a) and grafting‐from (b) approaches.

The formation of hydrated polymer brush layers, represents a prerequisite to reach full inertness towards biological contamination. This is due to the interfacial entropic and enthalpic barriers induced by a swollen and dense brush, which hinder surface interaction with proteins and larger biological objects. Polymers used for the surface modification of biomaterials have been largely based on poly(ethylene glycol) (PEG) derivatives. However, a n ­ umber of draw­ backs were encountered due to PEG’s susceptibility to ­oxidation and enzy­ matic degradation when immersed in physiological media, which leads to the formation of toxic compounds, including p­eroxides and ­aldehydes [2,25–28]. Thus efforts have been directed on finding alternative polymers possessing similar biocompatibility and p ­ erformance to PEGs. Among these, poly‐2‐ alkyl‐2‐­oxazolines (PAOXAs) have recently emerged as suitable substitutes [29–33]. Their tertiary amide‐bearing backbone stabilizes them in biological media against e­ nzymatic activity, and in addition, they showed low toxicity, high  ­ biocompatibility, and  higher  stability compared to  PEG analogues [31,34–38]. The most ­hydrophilic poly‐2‐methyl‐2‐oxazoline (PMOXA) and

Surface Functionalization by Poly(2-oxazoline)s

poly‐2‐ethyl‐2‐­oxazoline (PEtOXA) were applied as surface modifiers, to pro­ duce grafted polymer assemblies showing excellent biopassive character and outstanding c­hemical stability [39,40]. PMOXA and PEtOXA adsorbates are synthesized by cationic ring‐opening polymerization (CROP), which is a versatile and controlled polymerization process allowing the synthesis of monodispersed polymers and the introduc­ tion of chain‐end functions by using chemically tailored initiator/terminator species (Figure  15.2) [41,42]. Mono‐ and hetero‐bifunctional polymers can be  easily synthesized by CROP, and either applied directly as adsorbates or used as building blocks for the preparation of surface‐reactive copolymer ­species [42]. PAOXA films have been successfully applied on a variety of ­biomaterial surfaces, including organics and inorganics. In this chapter, a concise but comprehensive prospectus on these surface modification methods will be provided. Special emphasis will be placed on the chemical tailoring of PAOXAs and how it addresses the functionalization of  diverse surfaces. Additionally, the performance of PAOXA assemblies as biopassive interfaces will be highlighted, emphasizing the potential these ­polymers have as the next‐generation of antifouling coatings.

15.2 ­Grafting‐To of End‐Functional Polyoxazolines The precise modulation of the chemical structure of the initiator and the ter­ minator species during the CROP process enables the synthesis of chain‐end functionalized PAOXAs [42]. Surface‐reactive groups can be introduced at the chain‐ends of the polymers through single or multiple synthetic steps. Some examples of end‐functionalized PMOXA and PEtOXA surface‐modifiers, capable of assembling through grafting‐to methods on a target surface are reported in Figure 15.3. Silane and catechol‐terminated PAOXA homopolymers were reacted to  ­ silicon oxide and metal oxide surfaces by wet deposition processes, ­forming  densely grafted and highly water swellable films [43–45]. Alkoxysilane‐­ terminated PEtOXA were applied on silicon oxide to produce hydrophilic polymer grafts forming a brush assembly (Figure  15.3a) [43,45]. Following a similar approach, catechol‐terminated PMOXA adsorbates assembled on the surface of ZnO nanocrystals (NCs), due to the chelating ability of the dopa­ mine derivatives used as anchors (Figure  15.3b and Figure  15.4) [44]. The hydrophilicity of PMOXA chains, their chemical robustness and the stability of the catechol‐metal oxide interaction [46–49], allowed for the formation of dense and highly swollen polymer shells. These provided an outstanding col­ loidal stability to the NCs, which could be kept efficiently dispersed in aqueous media for up to 6 months (Figure 15.4).

459

460

Polymers for Biomedicine

(a) O n

Poly(ethylene glycol), PEG O N n

Poly-2(methyl-2 oxazoline), PMOXA

O N n

Poly-2(ethyl-2 oxazoline), PEtOXA

(b) CROP process

Initiation R1 –X + N

R1

O

N +O – X

R2

R2

Propagation R1

N+ O – X R2

+ nN

O

R1 O

R2

N R2

O

N+ O – X n R2

R1 O

R2 N

N

n

X

R2

Termination R1 O

N

N

n

R2

O – + H–R3 X

R1

R3 n+1

N

R2 O

R2

Figure 15.2  (a) The chemical structures of PEG, PMOXA and PEtOXA. (b) The CROP process, featuring initiation by electrophiles, propagation by opening of the oxazolinium ring and isomerization, and termination by a suitable nucleophile. The synthesis of α,ω‐heterobifunctional PAOXA presenting R1 and R3 chain‐end groups is especially highlighted.

Surface Functionalization by Poly(2-oxazoline)s

(a)

O

R O O Si O

N

N H

n

R = –CH3 or –C2H5 n = 15 or 30

(b)

O

H N

HO NO2

HO

N

O

N n

n = 25 or 50

(c)

S RAFT

S

O

m

S

OH S

N

O

m = 20 or 40 n = 6 or 12

O

N

N

n-1

m

O

Ammonolysis

O

O CROP

SH

m

O

O

O N

n

n

Figure 15.3  Silane‐, catechol‐, and thiol‐bearing PAOXA adsorbates for the functionalization of silicon oxide, metal oxide, and gold surfaces, respectively.

(a)

(b)

(c)

(d) 30 nm

Zno

6 months

H O N N

O

O N

O

1 μm N O

Figure 15.4  (a) Functionalization of ZnO NCs by PMOXA‐nitrodopamine adsorbates. Atomic force microscopy (AFM) tapping‐mode micrograph (b) and transmission electron micrograph (c) displaying PMOXA‐stabilized ZnO NCs (d) ZnO NCs/PMOXA aqueous suspensions after 6 months of storage, no signs of aggregation or precipitation are observed. Source: Morgese 2015 [44]. Reproduced with permission of American Chemical Society.

461

462

Polymers for Biomedicine

Bottle brush polymers consisting of a poly(methacrylic acid) (PMAA) back­ bone, short PMOXA side chains (PMAA‐g‐PMOXA) and a thiol function at the PMAA chain‐end, were applied to gold surfaces (Figure 15.3c) [50]. These polymeric adsorbates were synthesized by reversible addition chain‐transfer polymerization (RAFT), to form the endgroup‐functionalized PMAA, which was subsequently used as macro‐terminator agent for the CROP of 2‐methyl‐2‐ oxazoline (MOXA), via the deprotonated carboxylic acid groups. SH‐PMAA‐g‐ PMOXA efficiently formed self‐assembled monolayers (SAMs) on gold substrates and led to a remarkable reduction of surface contamination by dif­ ferent proteins [50]. Although direct grafting‐to of PAOXA species bearing a surface‐reactive function represents a relatively easy approach to functionalize biomaterials and nanoparticles, the formation of polymer assemblies with a sufficiently high grafting density to fully prevent the adhesion of proteins has never been demonstrated to date. This is presumably due to the self‐limiting character of  the grafting‐to process. In particular, during polymer chemisorption, the formation of ultra‐dense, and thus fully biopassive layers is inhibited by the  steric hindrance between already chemisorbed chains and others ­approaching to the surface. The fabrication of dense and biopassive PAOXA grafts requires the use of block‐copolymers or graft‐copolymers including comonomers forming seg­ ments that enhance the reactivity of the entire adsorbate towards the target substrate. This strategy is often accompanied by the use of “grafting promot­ ers,” previously immobilized on the inorganic surface, which can favor the chemisorption of the polymer adsorbates by providing multiple anchoring points for the reactive segments (Figure 15.4). Grafting promoters can be thin films of highly functionalizable polymers or functional SAMs, both of them exposing reactive groups that can quantitatively and readily combine with the PAOXA‐based adsorbates. Following a similar approach, Zhang et  al. first synthesized star‐shaped poly(ethylenimine) (PEI)‐g‐PMOXA adsorbates, which were subsequently reacted with surface‐immobilized polydopamine (PDA) layers capable of mul­ tiple binding to the amino functions on PEI (Figure 15.5) [51], thus forming PMOXA brush interfaces. Density and coverage of the brushes can be easily controlled by modulating the PMOXA grafting density along the star copoly­ mers, and the length of the PMOXA segments. Optimization of these param­ eters allowed the fabrication of PMOXA brushes displaying full biopassivity towards the adsorption of different proteins (Figure 15.6). Following an alternative approach, copolymerization of 2‐isopropyl‐2‐­ oxazoline (iPrOXA) with 2‐(2‐methoxycarbonylethyl)‐2‐oxazoline (esterOXA) to yield PiPrOXA‐b‐PesterOXA allowed the preparation of copolymers with functional PAOXA segments featuring carboxylic acid moieties as side chains [52]. These were efficiently grafted to spin‐coated poly(glycidyl methacrylate)

(a) O

O

1. MeOTf 2. Na2CO3/H2O

N

n

MOXA

H2N

O

OCN

O N

N

O

NCO

HMDI

OH

CHCl3

N

reflux

n1

PMOXA-NCO

NH2

N

n2

PEI core O

NH2

Hyperbranched PEI CHCl3

NCO

O

PMOXA-OH H N

H N

O

nN

O N n

N

O

H N

O

N H

O

reflux

N H

PEl-g-PMOXA

H N n1

PMOXA arm

NH2

N

n2

NH2

(b)

Platelet

Protein

Cell

Mixture

Linear PMOXA/PDA film

H2N

OH OH

Su

PDA

bs

HO

tra

te

NH2

Star PMOXA/PDA film

HO

Figure 15.5  (a) Synthesis of linear PMOXA‐OH and multi‐arm star‐copolymer PEI‐g‐PMOXA. (b) Fabrication of PMOXA/ PDA films exploiting PDA as grafting promoter layer. Source: Zhang 2015 [51]. Reproduced with permission of Royal Society of Chemistry.

Polymers for Biomedicine 700 Fibrinogen γ -Globulin

600 Protein adsorption (ng/cm2)

HSA Lysozyme Myoglobin

500 400 300 200 100

PE

I–

PM

O XA

kD a 25

(5 K) –O g w (7 H ith 0 ou ) – t d PM op O am XA PE I– in (5K e g( ) w 70 ith ) – do PM pa O m XA in ( e 5K )

I PE

h /2 PD A

re

Au

0

Ba

464

Figure 15.6  Adsorption of five different proteins on differently functionalized Au surfaces as measured by surface plasmon resonance (SPR) spectroscopy. Source: Zhang 2015 [51]. Reproduced with permission of Royal Society of Chemistry.

(PGMA) films, via the reaction between the epoxy groups along the PGMA and the carboxylate functions of the PAOXA copolymers (Figure  15.7), and finally ­produced PiPrOXA brushes stretching out at the interface in the high density regime. The application of glycidyl groups to ease the reactivity of PAOXA adsorbates towards the surface was also reported by Bai et al., who proposed the copolymerization of glycidyl methacrylate (GMA) and PMOXA‐ methacrylate (PMOXA‐MA) to produce random copolymers PMOXA‐MA‐r‐ PGMA by free radical polymerization [53]. These bottle‐brush adsorbates thus possessed a surface‐reactive polymethacrylate backbone exposing epoxy groups alternated to longer PMOXA side‐chains. Subsequent wet deposition onto silicon oxide surfaces, followed by thermal treatment, enabled the reac­ tion between the silanol functions at the substrate and the epoxy‐bearing copolymer backbones, and led to the immobilization of the copolymer (Figure 15.8). The reactivity towards the silicon oxide surface and the composition of the  subsequently formed coatings were tuned by varying the comonomer

Surface Functionalization by Poly(2-oxazoline)s O N HOOC O N

N

N

O

O

O

N

O

O

O

O

OH

4

100

O

O

PiPrOXA-b-PesterOXA x-2

PGMA

Figure 15.7  PiPrOXA‐b‐PesterOXA copolymers were applied for the functionalization of PGMA‐coated surfaces to yield dense PiPrOXA brush interfaces. Source: Agarwal 2012 [52]. Reproduced with permission of American Chemical Society.

O

1) Spin-casting x

O

O

y

O O

2) Δ

O

O n

x

O

N

O

O N

n

O

y

HO

x/y = 3, 1 or 0.33 O

OH

OH

O

Si

Si

Si

O

O

O

O

O

O

Figure 15.8  PMOXA‐MA‐r‐PGMA bottle‐brush copolymers are capable of reacting with the silicon oxide surface through the reaction between the epoxy groups of the GMA units and the silanol functions at the substrate. Source: Bai 2014 [53]. Reproduced with permission of Royal Society of Chemistry.

composition. Higher relative concentrations of GMA produced thicker ­copolymer films, due to the higher reactivity of the copolymer backbones towards the silanol groups on the substrate. However, the antifouling proper­ ties of the copolymer films were determined by the relative PMOXA‐MA ­content. Hence, assemblies of PMOXA‐MA‐r‐PGMA with a PMOXA:GMA

465

466

Polymers for Biomedicine

ratio of 3:1 showed the lowest film thickness among the copolymers studied, but efficiently prevented the adsorption of proteins and the adhesion of Escherichia coli (E. coli) (Figure 15.9). Functional films promoting the grafting of PAOXA adsorbates also included poly‐(4‐azidomethylstyrene) (PS‐N3) layers, which were used in the surface grafting of alkyne‐terminated PMOXA via Cu‐catalyzed Huisgen cycloaddi­ tion (click) [54], and dense assemblies of tetraether lipids (TEL), which enabled multiple binding of amino‐bearing PEtOXA random copolymers via cyanuric chloride reactive spacers (Figure 15.10) [55]. In both these cases, antifouling PAOXA brushes hindering the adsorption of proteins, cells, and bacteria were successfully fabricated.

15.3 ­Photochemical Surface Grafting of Polyoxazolines Surface grafting of PAOXAs can be accomplished by applying photosensitive layers producing highly reactive species to covalently bind the polymer adsorbates. Examples include benzophenone [56–58] and perfluorophenyl azide (PFPA) [59–61] derivatives, which can be deposited on diverse inorganic  and polymer surfaces. These molecules form highly reactive ­ ­carbon radicals and nitrene species, respectively, upon UV irradiation, which react with PAOXAs without any specific coupling function (Figure  15.11 and  15.12). Both benzophenone and PFPA bind to the CH groups along the  PAOXA chains, forming monolayers of polymer grafts with high ­efficiency [57]. Although benzophenone‐mediated grafting was easily applied for the func­ tionalization of biomaterials, the formation of densely grafted, biopassive PEtOXA brushes has not yet been demonstrated. In contrast, the application of PFPA monolayers both on gold and silicon oxide surfaces allowed the fabrica­ tion of PEtOXA brushes that efficiently hindered surface contamination by bovine serum albumin (BSA) and other biomolecules (Figure 15.12) [61]. This was attained by photografting high molar mass PEtOXA, which yielded a higher surface concentration of hydrophilic EtOXA units when compared to shorter polymer grafts. The broad applicability of this method was demonstrated by incorporating PFPA functions into polymers (Figure 15.12) that could be deposited on sur­ faces and applied as grafting promoters. The subsequent UV‐assisted immobi­ lization of PEtOXA with different molar masses produced brush layers that effectively resisted the adsorption of proteins [61], marine bacteria, and zoo­ spores from algae [62].

(i) 50 45 40

Thickness (nm)

35 30 25 20 15 10 5 0 PMOXA-r3/1-GMA

PMOXA-r1/1-GMA

PMOXA-r1/3-GMA

(ii) (a)

(b)

(c)

(d)

Figure 15.9  PMOXA‐MA‐r‐PGMA copolymers presenting higher content of GMA comonomer formed thicker films on silicon oxide surfaces (i). Nevertheless, the best antifouling and antimicrobial properties were displayed by copolymers presenting the highest concentration of PMOXA segments. (ii) Optical micrographs of stained E. Coli adhering on (a) the pristine, unfunctionalized silicon oxide, (b) PMOXA‐MA(1)‐r‐PGMA(3), (c) PMOXA(1)‐MA‐r‐PGMA(1), and (d) PMOXA(1)‐MA‐r‐PGMA(0.33). Source: Bai 2014 [53]. Reproduced with permission of Royal Society of Chemistry.

(a)

HO

N n

O

N N N

N3 HO

N

n

O n = 25 or 50

x -1

(b) O O

N

N

N n

NH2

NH

O

NH

N

N

n

n

CI

N

N CI N N

O

O

n = 25 - 55

Tetraether lipids (TEL)

Figure 15.10  Surface functionalization by PAOXA adsorbates exploiting click reactions (a) between alkyne‐PMOXA and PS‐N3, Source: Agarwal 2012 [52]. Reproduced with permission of American Chemical Society. (b) Conjugation between PEtOXA‐NH2 and cyanuric chloride‐bearing TEL assemblies. Source: Tauhardt 2014 [55]. Reproduced with permission of Royal Society of Chemistry.

z N

O N

1) Spin-casting x

O

O Mw = 380 kDa

N

2) UV O

O Si O

N y

y

O OH

O Si O

x = y + z +1

y

Figure 15.11  Benzophenone‐mediate photografting of PEtOXA on functionalized silicon oxide and glass surfaces.

x=z+y+1 Mw = 5, 50, 200, 500 kDa

N

N

x O

F

y z H

N3

F

F

F

F O

s

or

F

F

F

F

HN

O

10 2

O Si O O

O

N3 or

N

F

O F

F

F F

F F

N

N

N F

F F

N3

O

O O

UV

F

O NH NH2 x

y

Figure 15.12  PFPA‐mediated grafting of PEtOXA. Three examples of PFPA‐based grafting promoters are shown: disulfide‐, silane‐, and poly(allylamine)‐based. Source: Wang 2011 [61]. Reproduced with permission of American Chemical Society.

O

470

Polymers for Biomedicine

15.4 ­Poly‐(L‐lysine)‐g‐PMOXA Copolymer Films The fabrication of protein repellent films by electrostatic self‐assembly of graft‐copolymers featuring a poly‐(L‐lysine) (PLL) backbone and biopassive polymer side chains (Figure 15.13) represented a milestone in the development of polymeric bio‐interfaces. The application of PLL‐g‐PEG copolymers first as stabilizers for biopolymer microcapsules and, later as surface modifiers for liv­ ing blood cells, was pioneered by Hubbell and coworkers [63–65]. In several subsequent studies by Textor, Spencer, and Hubbell, the electrostatic interac­ tion between the positively charged PLL backbone and several negatively charged inorganic and organic surfaces was exploited to form PEG interfaces displaying excellent protein‐resistant properties and high lubricity [27,66–74]. These included the functionalization of substrates, such as titania, silicon oxide, glass, and tissue culture polystyrene (TCPS). The morphology of the so‐formed copolymer adlayers presents a comblike configuration, where sur­ face‐attached PLL segments support PEG grafts, which stretch at the interface. Due to this distinctive architecture and the nature of the surface assembly ­process, the molecular characteristics of the copolymers, that is, the side‐ chain  density (copolymer grafting density) and length (PEG molar mass)

(b) O N

(a)

O N O

O N

O

O N

O O O

O

O

O

NH

H N

O x N H

+ O y

+

+

+ +

+

+ +

H N

+

– – – – – – – – – H3N

H3N

Figure 15.13  PLL‐g‐PEG (a) and PLL‐g‐PMOXA (b) graft‐copolymers for the functionalization of negatively charged, inorganic, and organic surfaces.

NH

O x N H

O y

Surface Functionalization by Poly(2-oxazoline)s

finally  determine the adlayer characteristics. These include a number of ­fundamental parameters, such as the interfacial conformation of the PEG grafts (“mushroom” vs brush), their surface coverage and, consequently, the biopassive properties of the adlayers [71,72,75]. Although PLL‐g‐PEG copolymers were successfully applied as coatings in a  wide range of applications, the fading interest in academic research and development in PEG‐based surface modifiers led Konradi and Textor to pro­ pose PMOXA as a variant to this very successful material. Inspired by the synthetic principles exploited for the designing of molecularly engineered PLL‐g‐PEG species, Konradi et  al. reported the synthesis of PLL‐g‐PMOXA copolymers and their successful application for the functionalization of Nb2O5 surfaces [76,77]. PLL‐g‐PMOXA was demonstrated to assemble on the metal oxide surfaces similarly to the PEG‐analogues and eliminated protein adsorp­ tion to a level of