Layer-by-Layer (LbL) coatings for controlled delivery of biologicals

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Thesis

Reference

Layer-by-Layer (LbL) coatings for controlled delivery of biologicals

SAKR, Omar

Abstract

Layer-by-Layer technology (LbL), a technique based on the deposition of oppositely charged polyelectrolytes layer-wise on the surface of interest, has gained increasing interest in the field of drug delivery. The presented work in this thesis explores new applications for Layer-by-Layer (LbL) technology in the field of controlled protein delivery. The basic motives for these studies were the protein-friendly mild formulation conditions where only aqueous solutions are used, in contrast to organic solvents typically employed in the fabrication of many other protein formulations, thus correct protein folding and activity are preserved.

Several formulations were designed for the loading and slow release of model and therapeutic biologicals from different carriers. Special attention was paid to testing the activity of released fractions. Moreover, double chambered nanoparticles were developed to host and co-deliver a protein and a small molecule drug to the same cells.

SAKR, Omar. Layer-by-Layer (LbL) coatings for controlled delivery of biologicals. Thèse de doctorat : Univ. Genève, 2015, no. Sc. 4874

URN : urn:nbn:ch:unige-819128

DOI : 10.13097/archive-ouverte/unige:81912

Available at:

http://archive-ouverte.unige.ch/unige:81912

Disclaimer: layout of this document may differ from the published version.

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Layer-by-Layer (LbL) Coatings for Controlled Delivery of Biologicals

THÈSE

présentée à la Faculté des sciences de l’Université de Genève

pour obtenir le grade de Docteur ès sciences, mention sciences pharmaceutiques

par

Omar Shoukry Sakr

du Caire (Egypte)

Thèse N°: 4874

Genève

Atelier de reproduction Repromail 2015

Section de Sciences Pharmaceutiques

Pharmaceutics and Biopharmaceutics

Professeur Gerrit Borchard

Docteur Olivier Jordan

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To those who lost their lives, their families, their freedom, because they dared to say the truth when others remained silent…

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“The formulation of the problem is often more essential than its solution. If I had an hour to solve a problem I'd spend 55 minutes thinking about the problem and 5 minutes thinking about solutions. ”

Albert Einstein

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Acknowledgment

It has been truly a long journey, spanning more than five years, 2 different countries, different working settings, environments and languages but it’s all wrapped together now, way better than I’ve ever dreamt. I feel much in debt for a lot of people, without whom, I’d never make it to the point of writing these lines.

To Prof. Gerrit Borchard, words can’t thank you enough. Gerrit, you have believed in me since the very beginning, and although I didn’t get the first PhD position I applied for in your group, but you insisted on bringing me on board. I will forever remember that early morning phone call when you said “Omar, I have another PhD project and you are in, unless you say no”, and since then, you have been, not only my PhD supervisor, but my advisor, friend and elder brother. Even after my PhD and with my new start‐up, your input as a scientific advisor was remarkably valuable and enlightening.

To the “chemoembolization team”, the amazing Dr. Olivier Jordan and Dr. “soon to be” Katrin Fuchs. We have been together through a lot in the last two and half years. Thank you for all the great work we have done together, for the lengthy discussions, critical opinions, and limitless help.

To my sincere friends in FABIO and FATEC, you are the best! We have been always there for each other and I’m quite sure that we will keep doing so for a lifetime. Special thanks for Tayeb Jbilou for the help with SEM imaging and more importantly French translation; you made my life way easier in Geneva. Thanks are also extended to Yassin Dhif, my very first Master student for his great help and translation of my French summary.

To my elder brother Dr. Hesham Hamed, you were one reason I could make it to Geneva in the first place. You stood beside me during this endless roller coaster and you are still there to advise, help and support. You deserve more thanks than words can convey (ya Beih) and of course, thanks for the tennis lessons.

I would like also to thank the jury members, including Prof. Jörg Huwyler, Prof. Eric Allémann and Dr. Lars Dähne for accepting to review my thesis and for their time spent in this process.

To my dear father and mother, I’m nothing except what you have raised me to be. It all comes down to your endless efforts, support and “Doa’a”. I was told that this might be an end to a long

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learning journey but the fact is that the journey has just started. So, bear with me and be always beside me.

Finally to my lovely wife Salwa, you are the big secret behind all this success and you know it. It’s beyond words to describe your infinite dedication, sacrifice and help. To my three little princesses and prince Hana, Nour and Salman, thank you for always brining a happy smile to my face.

Omar Sakr Geneva, Nov. 2015

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List of Abbreviations

AD: adamantane Alg: Alginate

BCA kit: Bicinchoninic Acid kit Bev: Bevacizumab

CAT: Catalase

cCMG: cationic cholesterol‐modified gelatin

CDP: Cyclodextrin containing Polycations or Cyclodextrin based Polymers CDs: Cyclodextrins

CE: Cholesterol esterase

CGM: cationized gelatin microspheres COX: Cholesterol oxidase

DD: degree of deacetylation DEBs: Drug‐eluting beads dsRNA: double stranded RNA

EPR: Enhanced permeability and retention GA: Glucoamylase

GI: Glucose isomerase GOD: Glucose oxidase

HCC: Hepatocellular carcinoma HSP: Heat Shock Protein

HUVEC: Human umbilical vein endothelial cells

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i.v.: intravenous

IBD: inflammatory bowel disease IFN: Interferon

IL: Interleukin LbL: Layer by Layer Lys: Lysozyme

MF: Melamine formaldehyde Mw: Molecular weight

mRNA: messenger Ribonucleic Acid NGs: Nanogels

NHDF: Normal human dermal fibroblasts

NiMOS: Nanoparticles‐in‐Microspheres Oral System PAH: Poly allylamine hydrochloride

PBS: Phosphate buffered saline PBAE: poly β amino esters

PCI: photochemical internalization

PDADMAC/PDDA: Poly diallyl dimethyl ammonium chloride PDI: Polydispersity index

pDNA: plasmid DNA

PECs: Polyelectrolyte complexes PEG: Polyethylene glycol

PEI: Polyethylenimine

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PEs: Polyelectrolytes pI: Isoelectric point

PLGA: Poly lactic glycolic acid PLL: Poly‐L‐lysine

POD: Peroxidase

PS particles: Polystyrene particles PSS: Polystyrene sulfonate

QCM: Quartz crystal microbalance RISC: RNA‐Induced Silencing Complex RNAi: Ribonucleic Acids interference s.c.: Subcutaneous

siRNA: Small Interference RNA

TACE: Transarterial chemoembolization TF: Transferrin

UV: Ultraviolet

VEGF: Vascular endothelial growth factor Vis: Visible light.

β‐GLS: β‐glucosidase

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Foreword

The increasing development of therapeutic proteins by pharmaceutical industry has highlighted several challenges that should be addressed carefully when deciding on a suitable formulation or delivery carrier. Among these challenges, primary obstacles are short protein biological half‐life, adverse immunogenic side effects, poor loading efficiency of the delivery carrier with only sub‐

therapeutic doses, lack of long‐term stability, and uncontrolled drug release profile. Furthermore, since most of these therapeutic proteins are intended to treat chronic diseases, patients are prescribed multiple injections for long time periods, which often compromises the patient compliance and increase health care costs. Therefore, sustained release formulations are much needed. One promising approach to solve some of these problems would be the local release of proteins from injectable carriers that are able to control the release kinetics of their payload. Many delivery systems have been investigated during the past few decades, and various microstructures such as liposomes, microgels, polymer micelles and nano/micropolymeric particles have been suggested as sustained release protein formulations.

Within this context, work presented in this thesis generally aimed to explore the versatility and potential of Layer‐by‐Layer technology (LbL) as a coating technique to achieve controlled release profile of therapeutic proteins loaded on different micro and nano‐carriers. Special attention was always given to maintaining maximum possible protein activity.

Since its introduction in 1992 by Decher et al., coating of flat surfaces as well as micro‐ and nanoparticles by Layer‐by‐Layer (LbL) technology has become an active area of research and is currently considered a hot topic with many potential applications, especially in drug encapsulation, sustained release dosage forms, protein delivery, and biosensors. Typically, this technique is based on the use of polyelectrolytes of opposite charges assembled layer‐wise on the surface of interest.

Thereby, building up a layered system of tunable characteristics, in terms of composition, nanometer range thickness, surface charge, permeability, and elasticity.

More particularly relevant to protein encapsulation, LbL deposition has the advantage of utilizing mild conditions (e.g., aqueous solutions), which are more favorable to preserve fragile protein folding and activity in contrast to organic solvents typically employed in the fabrication of other protein encapsulation systems. Interestingly, it has been shown that retained water present in LbL

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films, in spite of the drying procedures applied, is important for the preservation of activity of biomolecules.

The first chapter of this thesis gives more details about LbL technology and reviews different approaches and LbL designs applied to immobilize and/or encapsulate various enzymes. One aim was to show the flexibility and the potential of this technique in obtaining different designs and architectures fulfilling a wide range of needs and uses. Another important aim was to shed light on the techniques usually applied to characterize LbL structures and to monitor the process of layer deposition. Methods for testing enzyme activity and stability were of high interest. Moreover, a special part was dedicated to LbL structures encapsulating two or more active proteins where the function depends on the sequential actions of encapsulated enzymes.

In the second chapter, siRNA was chosen as a protein of high therapeutic potential and the use of biopolymers as vectors for its delivery was reviewed. This chapter illustrates many problems related to protein delivery in general and siRNA in specific and moves on to discuss ideal properties of a promising siRNA delivery system, followed by a comprehensive up‐to‐date list of biopolymers used for the delivery of siRNA, highlighting their source, composition, physicochemical properties as related to the current subject. Special attention was paid to the ability of mentioned polymers to protect siRNA fragments, prolong their systemic circulation, and enhance siRNA cellular uptake in vivo.

Work presented in the third chapter explores the potential of Layer‐by‐Layer technology (LbL) as a coating technique to achieve sustained release profile of a model protein (Lysozyme) loaded on microparticulate carriers. The adopted strategy is based on building alternating multilayers of cationic degradable polymers made of poly β‐amino esters (PBAEs) and chondroitin sulfate as a biocompatible negative polyelectrolyte. Loading distribution was monitored by fluorescence imaging, and followed by depositing a series of LbL coatings of different thicknesses. Release of Lys from these formulations was studied and activity of released fraction was determined.

In the fourth chapter, this basic work was taken one step further. Bevacizumab, the anti‐ vascular endothelial growth factor (VEGF) antibody was loaded on embolic beads (DC Bead^TM) using different strategies. This was followed by applying an LbL coating film to slow the release of protein content, and also minimize the burst release phase usually encountered with protein formulations, while maintaining maximum possible activity. Loading kinetics were studied, the deposition of

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polyelectrolyte layers was followed and proven by zeta potential measurements and the coating was visualized using SEM imaging. Finally, the activity of BEV released from DC beads was tested using a 2D MTT assay and fibrin bead assay; a 3D anti‐angiogenic test developed in‐house.

The fifth chapter tackles a different challenge and brings on a different way of thinking. As many combination therapies suggest the use of a protein and a small molecule drug together, as exemplified by the use of siRNA molecules in combination with anticancer drugs to treat different resistant tumours, co‐delivery of both APIs might be very challenging because of the different kinetics of both, leading to distinctive physiological fates and non‐uniform distribution. Therefore, one strategy toward a more effective co‐delivery may be the careful design of a nano‐delivery system that is able to efficiently encapsulate and co‐localize both cargo species in the target cells.

Hereby, the preparation of a novel double chambered, nanoparticulate system was described, applying a simple and less time‐consuming LbL fabrication method using concentration tubes. Small poly lactide‐co‐glycolide (PLGA) nanoparticles loaded with fluorescein base as a model small hydrophobic molecule, represents the “internal chamber” while the “external chamber” was formed of alternating polyelectrolyte layers containing lysozyme as a model protein. These nanocarriers were taken up by MDCK cells in vitro, where a co‐localization of both model compounds was shown by confocal imaging.

In conclusion, results shown in chapter three and four illustrate the versatility of LbL technology and the flexibility of using different building blocks. Moreover, the ability of LbL films to successfully coat loaded hydrogel beads and to sustain the release of its protein drug cargo maintaining an acceptable degree of bioactivity was shown for different proteins. This approach might be very appealing for physicians and patients undergoing treatment by chemoembolization as the efficient local delivery would save patients the need to get high doses of systemic protein injections. In chapter five, successful co‐localization of 2 different species of APIs into the same cell was shown and the designed nanocarrier system might serve as a handy tool for many applications that requires concomitant treatment with a protein drug and a small‐molecule drug.

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Chapter One

Encapsulation of Enzymes in Layer‐by‐Layer (LbL) structures: Latest Advances and Applications

Omar. S. Sakr^1,2 and Gerrit Borchard²

1Capsulution Pharma AG, Volmerstrasse 7b, D‐12489 Berlin, Germany

2School of Pharmaceutical Sciences, University of Geneva, University of Lausanne, quai Ernest Ansermet 30, CH‐1211 Geneva 4, Switzerland

Review Article

Published in:

Biomacromolecules 2013, 14: 2117−2135

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Abstract

Layer‐by‐Layer (LbL) technology has become an active area of research and is currently considered a hot topic with many potential applications, especially in the pharmaceutical and biopharmaceutical fields. This review is providing an overview of current approaches and applications of LbL designs used to immobilize and/or encapsulate various enzymes. One aim was to show the versatility and the potential of this technique in obtaining different designs and architectures fulfilling a wide range of needs and applications. Another important aim was to shed light on the techniques commonly used to characterize LbL structures and to monitor the process of layer deposition. Special attention was given to LbL structures encapsulating multiple enzymes where the function depends on the sequential activities of encapsulated enzymes.

Key words:

Layer by Layer (LbL) – Enzymes – Encapsulation –Sequential action ‐ Protein delivery – Biosensors

Graphical abstract

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1 Introduction

Since its introduction in 1992 by Decher et al. [1], coating of flat surfaces as well as micro‐ and nanoparticles by Layer‐by‐Layer (LbL) technology has become an active area of research and is currently considered a hot topic with many potential applications, especially in drug encapsulation, sustained release dosage forms, protein delivery, and biosensors [2, 3].

LbL deposition is an established method for the fabrication of multicomposite ultrathin films on solid surfaces [1, 4, 5]. Typically, this technique is based on the use of polyelectrolytes of opposite charges assembled layer‐wise on the surface of interest, thereby building up a layered system of tunable characteristics (Figure 1), in terms of composition, nanometer range thickness, surface charge, permeability, and elasticity [2]. Using this approach, a variety of materials, including charged and uncharged species, have been successfully assembled into nanoscale multilayered structures. Multilayers may be formed by using LbL assembly techniques that rely on electrostatic interactions [4], hydrogen bonding [6], coordination bonding [7], charge transfer [8], molecular recognition [9], hydrophobic interactions [10], or a combination of these. As a result, such films may be comprised of a diverse range of components, including but not limited to biomacromolecules [11‐13], nanoparticles [14], dyes [15], dendrimers [16], and conductive polymers [17].

More particularly relevant to protein encapsulation, LbL deposition has the advantage of utilizing mild conditions (e.g., aqueous solutions), which are more favorable to preserve fragile protein folding and activity in contrast to organic solvents typically employed in the fabrication of other protein encapsulation systems such as microparticles, microcapsules, microemulsions, and liposomes [18]. These, in addition, suffer from some limitations including polydispersity, core solidification and residual organic solvents [19]. It has been proven that entrained water is present in LbL films, in spite of the drying procedures applied, which is important for the preservation of activity of biomolecules [20].

Immobilization of enzymes and active proteins is of great scientific and practical importance [21].

Especially when considering microreactor and biosensor development, immobilization and encapsulation of enzymes in LbL structures can provide a means of concentrating and protecting the bioactive molecules in a defined volume, creating a partitioned microenvironment with

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tuneable properties, which could therefore be used to balance the diffusion and reaction as needed for the sensor function [22]. Moreover, encapsulation offers a great advantage to these sensitive structures by protecting encapsulated enzymes from proteolytic enzymes and microbes [23].

Figure 1: Simplified molecular picture of the first two adsorption steps, depicting film deposition starting with a negatively charged surface. Steps 1 and 3 represent the adsorption of a polycation and a polyanion, respectively, and steps 2 and 4 are washing steps.

LbL encapsulation of proteins and enzymes is superior to previously used techniques, where proteins have traditionally been immobilized onto solid surfaces by physical adsorption, solvent casting, covalent binding, and electropolymerization [24]. However, these methods often produce irregular films at low protein density. Ordered protein multilayer films with a high protein density have been constructed by using Langmuir‐Blodgett deposition methods [24, 25] or by exploiting biospecific interactions[26]. Denaturation of the immobilized proteins, restricted film permeability to substrates, and the highly specific nature of the assembly limit the wide applicability of all these methods [27].

Recently, basic LbL principles [5], their use to functionalize nanoparticles [2], to control and modify permeability and release kinetics [28‐30], to deliver biotherapeutics such as antigens and genes [31], and to fabricate stimuli responsive capsules [32, 33], were reviewed.

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The present work is devoted to review different approaches and LbL designs applied to immobilize and/or encapsulate various enzymes. One aim was to show the versatility and the potential of this technique in obtaining different designs and architectures fulfilling a wide range of needs and uses (Table 1). Another important aim was to shed light on the techniques usually applied to characterize LbL structures and to monitor the process of layer deposition. Methods for testing enzyme activity and stability were of higher interest. Moreover, a special part is dedicated to LbL structures encapsulating two or more active proteins where the function depends on the sequential actions of encapsulated enzymes. It is noteworthy to mention that, due to the great interest LbL encapsulation techniques have gained; considerable research work is published on enzyme encapsulated in LbL structures. This review attempts to cover the different LbL designs and structures supported with selected examples from the literature.

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Table 1: Summary of main LbL designs used to encapsulate enzymes reviewed in this article

Enzyme LbL design Layer components^(a) Ref.

Catalase (CAT)

Microspheres: based on sacrificial melamine formaldehyde particles, partially decomposed after layer deposition.

dextran sulfate / protamine

[35]

Capsules: based on building LbL multilayers on CAT crystals

PSS/PAH [37]

Mixed design: CAT microcrystals were first encapsulated in PSS/PAH, then films of alternating layers of CAT capsules and oppositely charged PEs were deposited on planar surfaces

PSS/PAH/

Cat‐capsules

[39]

Mixed design: CAT microcrystals were first encapsulated in PSS/PAH, then thin films of alternating layers of CAT capsules and oppositely charged PEs were deposited on gold electrodes for biosensing of H2O2.

PSS/PAH/

Cat‐capsules

[40]

Coating films: CAT was first encapsulated in small gold nanoparticles, and then electrostatically assembled with cationic PE on planar surfaces and colloidal particles

PSS/PAH/

CAT‐AUNPs

[41]

α‐

Chymotrypsin

Capsules: based on building LbL multilayers on α‐chemotrypsin crystals

PSS/PAH [21,

42]

Hollow capsules: based on sacrificial melamine formaldehyde particles, decomposed after layer deposition.

PSS/PAH

alginate/protamine [23]

[44]

Cholesterol oxidase (COX)

Coating films: cholesterol biosensor based on immobilization of COX in LbL films deposited on glass substrates.

PSS/PEI/COX [45]

Coating films: cholesterol biosensor based on immobilization of COX in LbL films deposited on glass substrates.

COX/PAH [20]

Coating films: cholesterol biosensors based on immobilizing COX in LbL films deposited on electrospun polyaniline nanofibers.

COX/ PDDA [46]

Glucose isomerase (GI)

Coating films: Both on planar surfaces and colloidal particles.

GI/Bi‐cationic pyridine salt

[52]

β‐glucosidase (β‐GLS)

Coating films: on colloidal particles. PAH/PSS/ β‐GLS [54]

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Urease

Hollow capsules:based on sacrificial melamine formaldehyde particles, decomposed after layer deposition.

PSS/PAH [58]

Coating films: on colloidal particles. PEI/Urease PDDA/Urease Urease/PSS Fe3O4

[60]

Peroxidase (POD)

Coating films: on colloidal particles. PSS/POD PSS/(POD‐PSS)

[27]

Hollow capsules: based on sacrificial melamine formaldehyde particles, decomposed after layer deposition.

PSS/ PDADMAC dextran sulfate/

protamine

[36, 70]

Glucose oxidase (GOD)

Coating films: on planar quartz slides. PSS/PEI/GOD [73]

Coating films: on colloidal particles PEI/GOD

PSS/ PAH/GOD/

Fe3O4 NP3/

PDADMAC

[27, 76]

Hollow capsules: based on sacrificial MnCO3 particles, decomposed after layer deposition.

Diasoresin/PSS/PAH [22]

Coating films: Glucose biosensor based on a pyrolytic graphite electrode with a modified surface by LbL deposition of GOD and PEI

GOD/PEI [80]

Capsules: glucose‐sensitive multilayer shells made of GOD/ PDMAEMA, built on insulin crystals.

GOD/PDMAEMA

[91]

Peroxidase

(POD) and Glucose

oxidase (GOD)

Coating films: on planar quartz slides.

PSS/POD GOD/PEI

[84]

Capsule in Capsule: based on coprecipitating the 2 enzymes with CaCO3 in 2 successive steps, separated by deposition of a thick multilayer membrane in between, and followed by dissolving the CaCO3.

PSS/PAH [86]

Glucose

oxidase (GOD) and

Glucoamylase (GA)

Coating films: on planar filter surfaces. GA/GOD/PEI/PSS [85]

Coating films: maltose sensor based gold electrode coated with GOD and GA sandwiched between bipolar quaternary ammonium salt

GA/ bipolar quaternary

ammonium salt / GOD

[88]

Catalase (CAT) and Glucose oxidase (GOD)

Capsules: glucose‐sensitive multilayer shells made of GOD/CAT held together via glutaraldehyde cross‐linking, built on insulin crystals.

GOD/ CAT [92]

(a) not indicating the order of addition.

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2 Catalase (CAT)

Catalase (CAT) is a common enzyme found in nearly all aerobic organisms. It catalyzes the decomposition of hydrogen peroxide to water and oxygen [34]. CAT was used by Balabushevich et al. [35] as a model protein of high molecular weight for encapsulation. Polyelectrolyte microspheres were obtained by alternating adsorption of dextran sulphate and protamine on melamine formaldehyde (MF) particles followed by partial hydrolysis of the MF core. In fact, the main difference between the fabrication of hollow capsules and microspheres prepared by this method employing MF cores is the core hydrolysis step (Figure 2). While the complete hydrolysis of the MF core produces hollow capsules, during the slow partial hydrolysis of MF core under mild conditions, the newly formed and positively charged amino groups interact with polyanionic structures of the first layer of the microcapsule shell resulting in the redistribution of membrane PEs and formation of a homogeneous, weakly cross‐linked, charged gelatinous matrix inside the microspheres [36].

CAT was encapsulated in the microspheres by simple incubation and mixing for sufficient time. The loaded microspheres were collected by centrifugation and washed. Finally, an additional layer of dextran sulphate was added to protect the enzyme adsorbed at the surface of the positive protamine layer. CAT was entrapped in microparticles at high efficiencies (70‐100%), depending on its original concentration in the incubation medium. The specific activity observed was dependent on the amount of protein entrapped in the microspheres. The authors reported that electrostatic and hydrophobic interactions were responsible for the interaction of the protein with the microspheres’ gelatinous core. Therefore, encapsulation and release of proteins from the proposed LbL system can be controlled via adjustment of pH, ionic strength, and temperature of the incubation medium [35].

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Figure 2: Scheme of production of microspheres and hollow microcapsules using commercially available melamine formaldehyde particles and alternating adsorption of polyelectrolytes.³⁵ “Source: Biochemistry (Mosc) 69, (7), 2004, 763‐9, Encapsulation of catalase in polyelectrolyte microspheres composed of melamine formaldehyde, dextran sulfate, and protamine, Balabushevich, N. G.; Zimina, E. P.; Larionova, N. I, Figure 1, © 2004 MAIK “Nauka/Interperiodica”. Reproduced with kind permission from Springer Science and Business Media.”

A different approach was adopted by Caruso et al. [37], where CAT crystals themselves were used as templates to deposit alternating layers of PSS/PAH to form the polymer capsule encapsulating the enzyme in the core at very high efficiency. The method takes advantage of the fact that the enzyme is present as a crystalline suspension in water at pH 5‐6 and may therefore be treated as a colloidal particle. It should be noted that enzyme crystal templating, however, presents several

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challenges that do not apply when templating, e.g., latex particles. First, the crystals are formed only under strictly defined conditions. Therefore, suitable conditions that facilitate polymer multilayer deposition on the crystal surface and do not destroy the enzyme crystal morphology (i.e.

to avoid its solubilization) need to be determined. Second, the permeability of the polymer capsule walls must be such that it permits encapsulation of the enzyme. In addition, since the primary usefulness of enzymes is their biological function, their activity must be preserved during encapsulation.

Practically, CAT crystals were separated from solubilized protein by washing and centrifugation several times with potassium acetate buffer of pH 5 at 4 °C. Chilled solutions were used to avoid significant solubilization of the enzyme crystals. CAT crystals exhibited a positive surface charge in water at pH 5 (+20 mV), as determined by electrophoretic mobility measurements. This positive charge at the surface of the crystals in principle makes them suitably charged templates for the deposition of polyelectrolyte layers of PSS and PAH. The successful deposition was proven by the reversal of the surface charge after each cycle of deposition, which is a characteristic of polyelectrolyte multilayer growth on colloidal templates [37].

To investigate the effect of the encapsulation process on the activity of CAT, the activity was measured after solubilization of CAT (by changing pH) and release from the polymer capsules. A recovered specific activity of 97% was obtained, compared with 100% for the uncoated CAT. This shows that the polymer multilayer coating of the CAT crystals proceeded without causing any significant loss of enzyme activity. Another important point was the ability of the capsule wall to protect the enzyme against proteolytic activity. As shown in Figure 3, solubilized, uncoated CAT (curves d and e) was inactivated by protease to more than 90% during an incubation time of 100 min. In contrast, no measurable loss in enzyme activity was observed for the polymer‐encapsulated (solubilized) CAT within 100 min under the same conditions (curves b and c). These results clearly demonstrate that a thin polymer coating of four layers (thickness of about 8 nm) is sufficient to prevent proteolysis of polymer encapsulated CAT [37]. These findings are consistent with the observation that proteins of molecular sizes greater than approximately 5 nm do not penetrate polyelectrolyte multilayer films [38]. Compared to traditional LbL methods, where solubilized charged enzyme molecules are deposited among the layers, encapsulated enzyme crystals display an up to 50‐fold increased biocatalytic activity, thus making them attractive candidates for various biotechnological applications [39].

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Figure 3: Stability of (a, d, e) solution‐solubilized catalase and (b, c) polymer‐multilayer encapsulated (solubilized) catalase with respect to proteolysis: (a) solution‐solubilized catalase crystals, no protease incubation (control); (b) [(PSS/PAH)2]‐coated (four layers) catalase, protease incubation; (c) [(PSS/PAH)4]‐

coated (eight layers) catalase, protease incubation; (d) and (e) repeat experiments for solubilized catalase, protease incubation. Proteolysis of the catalase was determined by measuring the decrease in the catalase enzyme activity [37] “Reprinted with permission from reference 37. Copyright 2000 American Chemical Society.”

Due to the interesting advantages of encapsulating enzyme crystals, such as high enzyme loading, preserved bioactivity of the encapsulated enzyme, the ability of the semipermeable PE coating to prevent the solubilized enzyme from leakage while simultaneously permitting the diffusion of small (substrate) molecules for enzyme reaction, this idea was taken a step forward by Jin et al. [39], where a mixed approach was adopted. CAT microcrystals were first encapsulated by the alternate adsorption of PSS and PAH on their surface, yielding an extremely high loading of active enzyme in the polyelectrolyte multilayer capsule. Then, multilayer films were constructed on planar surfaces (quartz crystal microbalance (QCM) electrodes or quartz slides) by LbL deposition of the polyelectrolyte‐coated CAT crystals and oppositely charged polyelectrolyte.

Moreover, this mixed approach was adopted by Yu et al. [40], where LbL encapsulated CAT microcrystals were assembled onto gold electrodes by sequential deposition with oppositely charged PEs, utilizing electrostatic interactions to form thin enzyme films for biosensing of H2O2. In

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addition to the aforementioned advantages of this design, the authors found that the PE layers encapsulating the enzyme effectively increase the surface charge density of the enzyme microcrystals, rendering them suitably charged components for the construction of biofunctional thin films. The PSS/PAH encapsulated CAT was shown to retain biological as well as electrochemical activity. Direct electron transfer between CAT molecules and the gold electrode was achieved without the aid of any electron mediator. As a H2O2 biosensor, films consisting of one layer of the encapsulated CAT displayed considerably higher (~5‐fold) and more stable electrocatalytic responses to the reduction of H2O2 than did corresponding films made of a single layer of non‐

encapsulated CAT or solubilized CAT. An increase in either the number of “precursor” PE layers between the gold electrodes and the CAT microcrystal layers in the film or the number of PE layers encapsulating the CAT microcrystals was found to decrease the electrocatalytic activity of the electrode. At low precursor PE layer numbers (~2) and encapsulating PE layers (~4), the current response was proportional to the H2O2 concentration in the range of 3.0 x 10^‐6 to 1.0 x 10^‐2 M. The overall electroactivity of the multilayer film increased for the first two layers of encapsulated CAT, after which a plateau was observed. This was attributed to the increasing difficulty of electron transfer and substrate diffusion limitations. Using immobilized PE‐encapsulated enzyme microcrystals for biosensing was shown to provide a versatile method to prepare films of high concentrations and tailored activities of enzymes.

More recently, another novel and versatile approach for the preparation of multilayers was introduced, where CAT was first encapsulated in small gold nanoparticles (CAT‐AuNPs), and then electrostatically assembled with anionic and cationic PEs on colloidal silica particles [41]. CAT‐AuNPs were synthesized directly from CAT stabilized gold suspensions and the diameter of the obtained particles was about 9 ± 3.5 nm. Since the pI of CAT is 5.6 [37], CAT‐AuNPs were positively charged at pH 3 and were assembled with the anionic polymer PSS. However, when the pH of the CAT‐AuNPs solution was changed from 3 to 9, charge reversal took place and the anionic CAT‐AuNPs were bound electrostatically to cationic PAH. As shown in Figure 4, an interesting feature of CAT‐AuNPs is that the pH dependent electrostatic properties of CAT‐AuNPs can control the structure of the hybrid nanocomposite, transforming them from well dispersed small nanoparticles at pH9 (due to repulsion forces between the similarly charged particles) to agglomerated colloidal particles with a diameter of about 50 nm or network‐structured composites, depending on the initial concentration of gold precursor [41].

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Figure 4: Schematic and TEM images for the preparation of CAT‐Au_NP with (a) dispersed, (b) colloidal and (c) network structures [41]. "Source: S. Kim, J. Park, J. Cho, Layer‐by‐layer assembled multilayers using catalase‐

encapsulated gold nanoparticles. Nanotechnology 21(37) (2010): 375702. (doi:10.1088/0957‐

In addition, this structural transformation had a significant effect on the surface morphology of CAT‐Au nanocomposite becoming rougher with fibrillary structures, especially in case of network‐

structured CAT‐AuNP. It was found that the total adsorbed amount of (PE/CAT‐AuNP network)5 multilayers was about 5.7 times higher than for (PAH/CAT)5 multilayers at the same solution concentration. Consequently, the higher CAT adsorption led to higher catalytic activity toward H2O2.Considering using these CAT‐AuNPs to coat an electrode surface, the rugged and fibrillary structure of PE/CAT‐AuNP colloids and PE/CAT‐AuNP network multilayers has an increased surface

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area, and as a result, increases the area of contact between the probe molecules and the CAT as well as the effective electron transfer rate. Therefore, this structural morphology may assist in increasing electrochemical sensitivity, which may be beneficial to a variety of biocatalytic applications [41].

3 α‐Chymotrypsin

α‐Chymotrypsin, a serine proteinase found in the intestinal tract and whose activity can be easily

monitored, has been chosen as a model enzyme for the evaluation of the feasibility and possible applications of LbL microcapsules.

Two main strategies were followed: in the first method [21, 42], α‐chymotrypsin was salted out from its acidic solution by mixing with an appropriate volume of a saturated solution of NaCl yielding particles with typical dimensions of 0.1–0.4 mm. Being positively charged at pH 2‐3, chymotrypsin (pI: 8.5) can be coated with a series of oppositely charged PEs such as PSS/PAH, with intermediate washing steps to remove excess unattached polymers. Although one can easily accept that the first layer would be the negatively charged polymer PSS, Balabushevitch et al. [42] found the possibility of starting the deposition with the positively charged polymer PAH. This was attributed to strong hydrophobic forces between PAH and the salted‐out protein core.

This preparation of microencapsulated α‐chymotrypsin retained 73% active site content after storage for 6 days at pH 3.0 at 4°C, while free enzyme had already lost 57% active site content after 6 days. Basic pancreatic trypsin inhibitor (BPTI, M.W. 6500), one of the known proteinase inhibitors[43], was used to challenge the ability of the LbL film to protect the encapsulated cargo.

BPTI was able to suppress 85% of free enzyme activity compared to only 13% of microencapsulated enzyme activity [42].

In the second method, hollow polyelectrolyte capsules were fabricated prior to loading with the model enzyme. For this purpose, small micron‐sized melamine formaldehyde (MF) particles were chosen as sacrificial cores. MF particles were coated with alternating layers of PSS/PAH [23] or alginate/protamine [44] at neutral pH, then the cores were dissolved by 0.1 M HCl solution and the empty shells were washed with water to remove MF residues to finally obtain hollow capsules. To load the enzyme α‐chymotrypsin into the hollow capsules, a small volume of the empty capsule suspension was centrifuged and the supernatant removed. Successively, capsules were

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resuspended in the enzyme solution of appropriate buffers and mixed well. After sufficient incubation of the capsules in the enzyme solution, the mixture was centrifuged and the capsules were washed three times with water. Confocal fluorescence images of labelled ‐chymotrypsin within capsules after mixing the protein and microcapsule suspensions indicated clearly the protein penetration through the open walls into the hollow capsules and the successful encapsulation of the enzyme.

Tiourina et al. [44] used rhodamine 110 [bis‐(succinoyl‐L‐alanyl‐L‐alanyl‐L‐prolyl‐L‐phenylalanyl amide)] to investigate the activity of the encapsulated enzyme, which was found to retain about 70% of its activity. It is important to mention that the enzymatic reaction inside the microcapsules requires substrate diffusion through the polyelectrolyte membrane while the reaction product is released into the surrounding medium [23].

In a different report [23], the author challenged the encapsulated α‐chymotrypsin with one of its inhibitors BPTI. While the native enzyme was completely inhibited at a [BPTI]/[‐chymotrypsin]

ratio of 1:1 after 5 min, the inhibitor did not influence the activity of the encapsulated α‐

chymotrypsin at a [BPTI]/[‐chymotrypsin] ratio of 2:1 after 30 min and the enzyme lost only 40%

activity at a [BPTI]/[‐chymotrypsin] ratio of 50:1 after 30 min. These results indicate that polyelectrolyte shells possess protective properties against high molecular weight inhibitors. The protection is mostly a result of the steric hindrance created by branched polymer molecules because protein‐protein interaction requires multi point contacts, which were apparently unavailable in the mentioned cases.

4 Cholesterol oxidase (COX)

Biosensors employing immobilized cholesterol oxidase (COX) for the detection of cholesterol may be more advantageous in comparison to standard methods such as spectrophotometry, gas–liquid chromatography and HPLC, owing to the simplicity and low costs involved. This is relevant to the development of reliable methods of cholesterol detection in blood, which is a fundamental parameter to identify disorders such as hypercholesterolemia, and to control the cholesterol level in foodstuff for human intake [20, 45, 46].

Ram et al. [45] reported the formation of a cholesterol biosensor electrode via immobilization of COX in LbL films. All solutions were adjusted at pH=7.5 as COX (pI between 4.6 and 5.2) can be used

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as a polyanion. Initially, a layer of polyanion PSS was adsorbed, followed by a layer of polycation PEI. Then, the LbL film was constructed by consecutive adsorption of polycation PEI and negatively charged proteins (COX) and cholesterol esterase (CE). The assemblies studied can be denoted as PSS/PEI/COX, PSS/PEI/COX/PEI/CE, PSS/PEI/COX‐CE/PEI. To monitor native and esterified cholesterol levels both COX and CE enzymes were employed simultaneously. Whereas COX catalyzes the oxidation of cholesterol, CE catalyzes the hydrolysis of esterified cholesterol, which is an important factor for the determination of total cholesterol, since about 70% of the cholesterol in blood is found to be esterified cholesterol [47, 48].

During preliminary studies, UV spectroscopy, QCM and electrochemical investigation clearly gave evidence for uniform enzyme immobilization on various substrates. The cell used to determine the response current consisted of cholesterol oxidase LBL film as working electrode, a platinum wire as counter electrode and Ag/AgCl as reference electrode in a solution of 100 mM phosphate buffer containing 1% of Triton X‐100. Considering the decrease of the activity of cholesterol oxidase at high concentrations of Triton X‐100 and low solubility of cholesterol at low concentrations of Triton X‐100, 1% of Triton X‐100in the buffer solution was used in this study. Measurements indicated a linear response to cholesterol up to concentrations of 1 mM, and further increase in cholesterol concentration showed a slower rise of the current. The stable current data were used for the calibration curve for actual measurements [45].

Moraes et al. [20] reported another design of COX based cholesterol biosensors via the immobilization of COX in LbL films, in alternation with layers of PAH. In preliminary studies, COX was alternately deposited with PAH onto previously cleaned glass substrates using the LbL technique. Before the PAH/COX film fabrication, two bilayers of PAH/polyvinyl sulfonic acid (PVS) were deposited onto the solid substrates to reduce substrate effects. Layer growth and film morphology were studied and the kinetic analysis of the adsorption process showed a two‐step process. Impedance spectroscopy measurements were used to detect cholesterol in liposomes (made from egg phosphatidyl glycerol) and natural egg yolk. Three electrodes were compared:

bare, PAH/PVS and PAH/COX coated gold electrodes. Owing to the specific interaction with COX in the LbL film, only gold electrodes coated with PAH/COX were able to detect cholesterol in aqueous solutions with a significantly higher sensitivity reaching 10⁻⁶ M [20].

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More recently, Shin et al. [46] proposed a novel architecture of cholesterol biosensors based on immobilizing COX in LbL films deposited on electrospun polyaniline nanofibers using poly diallyl dimethyl ammonium chloride (PDDA) as a counter ion. The nanofibers were composed of polystyrene as a core and polyaniline as a conducting polymer. Nanofibers were produced under a high voltage electrostatic field between a metallic nozzle of a syringe and a metallic collector. The charged polymer solution is jetted from the metal needle to the grounded collector. At working distance, the polymer jet elongates, solidifies, and deposits on the collector. The fibers are deposited in the form of a non‐woven fabric onto the target collector. Such nanofibrous membranes have many attractive features when used as supports for enzyme immobilization.

These include a large surface area for the attachment of enzymes, a nanofibrous morphology to improve the mass‐transfer rate of the substrate, and a membrane‐like structure for easy recovery from the reaction media and continuous operations in a bioreactor. The high surface area to volume ratio makes electrospun conducting polymer nanofibers particularly interesting for sensing applications [49, 50]. One layer of TiO2 was coated onto this nanofiber, and five alternate cycles of PDDA and COX adsorption were carried out (Figure 5).

Figure 5: (a) SEM of nanofibers for the blend of polyaniline and polystyrene before LbL coating (bar represents 5 µm), (b) SEM of nanofibers that COX was immobilized via LbL coating (bar represents 5 µm).

A B

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A cholesterol biosensor was fabricated using a standard one‐compartment three‐electrode cell and used for all electrochemical experiments. The sensing electrode was fabricated in 40 mL of 0.1 M phosphate buffer solution at pH 6.3 containing 1% Triton X‐100, using the nanofiber mat onto which COX had been immobilized as a working electrode, the reference electrode was Ag/AgCl (3 M KCl) and the counter electrode was a platinum wire (20 cm). Measurements showed a linear electrical response up to a concentration of 0.35 mM cholesterol. Accurate data could not be obtained using the nanofiber mat onto which two layers of COX had been deposited, and best results were obtained when more than five layers of COX had been deposited on the nanofiber mat [46].

5 Glucose isomerase (GI)

Glucose isomerase (GI, MW of ~ 160000, pI: 4.7) converts D‐glucose to D‐fructose in a reaction that is industrially applied to the production of high‐fructose corn syrup [51]. GI was immobilized by Kong et al. [52], in ultrathin films both on planar surfaces and on porous p‐trimethylamine‐

polystyrene (TMPS) beads. Negative charge was induced to GI molecules by adjusting the solution pH to above its pI. GI was deposited in alternation with bi‐cationic pyridine salt (Figure 6). The alternating molecular deposition of glucose isomerase and bipolar pyridine salt was followed by means of UV/VIS absorption spectra and the linear increase of the optical density of the films with increasing number of layers proved the process of deposition. The glucose isomerase activity was assayed following Tomas's cysteine‐carbazole method for fructose [53]. The glucose isomerase activity was preserved in the film, and the activity increased with the number of enzyme layers.

However, the average activity per enzyme layer decreased with the number of enzyme layers, because the more enzyme layers are deposited, the larger is the limitation of the diffusion of the substrate to reach the enzyme, which resulted in a great decrease of the specific activity of the enzyme (Figure 7).

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Figure 6: Model for alternating deposition film of glucose isomerase and bipolar pyridine salt through electrostatic interaction Bi‐cationic pyridine salt [52]. “Source: Macromol Rapid Commun, 15, 1994, 405‐409.

Kong, W.; Zhang, X.; Gao, M. L.; Zhou, H.; Li, W.; Shen, J. C. Copyright [1994]. This material is reproduced with permission of John Wiley & Sons, Inc.”.

Figure 7: Relationship between activity of glucose isomerase and the number of the enzyme layers. Total activity of enzyme in the slide (circles) average activity per enzyme layer (squares) [52] “Source: Macromol Rapid Commun, 15, 1994, 405‐409. Kong, W.; Zhang, X.; Gao, M. L.; Zhou, H.; Li, W.; Shen, J. C. Copyright [1994]. This material is reproduced with permission of John Wiley & Sons, Inc.”.

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6 β‐glucosidase (β‐GLS)

β‐glucosidase (β‐GLS) enzyme layers, each separated by oppositely charged polyelectrolyte PSS,

were deposited onto polystyrene (PS) latex particles using the layer‐by‐layer adsorption technique, as a new means to perform enzymatic glucosidation [54]. The purpose of o‐glycosyl hydrolases such as β‐GLS is the hydrolysis of glycosidic bonds. Reverse hydrolysis on its part is known to be an elegant method to form glycosidic bonds via enzymatic catalysis, yielding stereoselective glycosidation. This is achieved by performing the glycosidation reaction in organic solvents, thereby keeping the water content of the system low. In this report, β‐GLS was applied to form glycosidic bonds between carbohydrates and non‐carbohydrate, hydrophobic moieties (long chain alcohol).

Four alternating PAH and PSS layers were deposited onto the PS particles (first layer PAH), resulting in negatively charged particles with PSS as the outermost layer. As the pI of the enzyme is at 5.5, β‐

GLS is positively charged under the experimental conditions employed (pH 4.8). Four enzyme layers were deposited, each separated by one PSS layer.

Successful deposition of layers on the particles surface was proven by showing surface charge reversal, and the growth of the PS latex particle diameter with layer deposition as followed by single‐particle light scattering (SPLS). By recording the light scattered from many individual particles, histograms of particle number versus scattering intensity (or SPLS intensity distributions) were obtained. These intensity distributions were then fitted using the Rayleigh–Debye–Gans theory [55], and refractive indices for the enzyme and polyelectrolyte layers [56], thereby providing a quantitative measure of the layer thickness. Thicknesses were measured for particles with PSS as the outermost layers, since these particles were found to have a higher colloidal stability. Regular multilayer film growth occurred: linear enzyme multilayer growth was observed with an average thickness of 3.5 ± 0.6 nm for a β‐GLS/PSS layer pair.

Enzymatically catalyzed glucosidation reactions carried out with multilayer‐coated particles showed distinct differences in overall yield. Quantitative determination of dodecyl glucoside in the reaction mixtures revealed a regular increase in yield with increasing β‐GLS layer number for particles with an outermost PSS layer. Particles with one enzyme layer yielded only traces of the product, which could not be quantified. The increasing amount of dodecyl glucoside observed for particles with two, three, and four β‐GLS layers indicated that enzyme in the inner layers also takes part in the

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catalytic process. This in turn suggests that the substrate can diffuse into the layers to interact with the active sites of immobilized enzyme.

It is worth mentioning that particles covered by enzyme as the outermost layer showed a higher tendency to aggregate compared to those with PE as the outermost layer. The colloidal stability of protein/polyelectrolyte coated particles is known to increase when the polyelectrolyte forms the outer layer, the reason most probably being the electrostatic and steric stabilization of the particles conferred by the polyelectrolyte [56]. However, catalysts with β‐GLS as the outermost layer yielded higher amounts of dodecyl glucoside (ca. 2–5 times more) than those having the enzyme layer coated with PSS. This may be explained by a loss of enzyme from the surface into solution (i.e. the reaction mixture) when the enzyme is not covered by polyelectrolyte, resulting in higher amounts of product.

7 Urease

Urease is a metalloenzyme that catalyzes the hydrolysis of urea to yield ammonia and carbon dioxide. It is found in a wide variety of organisms including plants, fungi and bacteria [57]. Stable hollow polyelectrolyte capsules containing urease (Mw 480 kDa) were produced by means of the layer‐by‐layer assembling of PAH and PSS on melamine formaldehyde microcores followed by the core decomposition at low pH [58]. The authors introduced a new method of loading urease enzyme into the particles depending on a finding that capsules were non‐permeable for urease in water and became permeable in a water/ethanol mixture. As shown in Figure 8 (left), confocal microscopy imaging illustrates that FITC‐labelled urease was excluded from the polyelectrolyte shells. The interior of the capsules remained dark and outside the capsule the background containing FITC‐urease was fluorescent. This observation indicated the closed state of the capsules.

Figure 8 (center) shows a confocal fluorescence image of labelled urease with capsules after addition of ethanol (1:1 water/ethanol mixture). In this case, the fluorescence signal localized in the interior of the capsule has the same intensity as the outside background fluorescence. This indicated the penetration of the enzyme into the capsules and requires an open state of the capsule wall. When the capsules were transferred back into water, the polyion shells became closed, and the urease was captured inside, as illustrated in Figure 8 (right). The interior of the capsule is bright and constant over time, and there is no fluorescence signal from the solution.

Thus, urease filled the capsules. The images did not change with time, indicating that urease is

Layer-by-Layer (LbL) coatings for controlled delivery of biologicals

Thesis

Reference

Layer-by-Layer (LbL) coatings for controlled delivery of biologicals

Layer-by-Layer (LbL) Coatings for Controlled Delivery of Biologicals

THÈSE

To those who lost their lives, their families, their freedom, because they dared to say the truth when others remained silent…

“The formulation of the problem is often more essential than its solution. If I had an hour to solve a problem I'd spend 55 minutes thinking about the problem and 5 minutes thinking about solutions. ”

Albert Einstein

Acknowledgment

Table of Contents

List of Abbreviations

Foreword

Chapter One

Encapsulation of Enzymes in Layer‐by‐Layer (LbL) structures: Latest Advances and Applications

Abstract

1 Introduction

2 Catalase (CAT)

3 α‐Chymotrypsin

4 Cholesterol oxidase (COX)

5 Glucose isomerase (GI)

6 β‐glucosidase (β‐GLS)

7 Urease