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5-2011
APPLICATION OF NANOTECHNOLOGY
FOR TARGETED DELIVERY OF
ANTIBACTERIAL ENZYMES AND FOR
ENZYME-BASED COATINGS ON MEDICAL
DEVICES AND IMPLANTS
Rohan Satishkumar
Clemson University, [email protected]
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Recommended Citation
Satishkumar, Rohan, "APPLICATION OF NANOTECHNOLOGY FOR TARGETED DELIVERY OF ANTIBACTERIAL
ENZYMES AND FOR ENZYME-BASED COATINGS ON MEDICAL DEVICES AND IMPLANTS" (2011). All Dissertations.
Paper 740.
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APPLICATION OF NANOTECHNOLOGY FOR TARGETED
DELIVERY OF ANTIBACTERIAL ENZYMES AND FOR ENZYMEBASED COATINGS ON MEDICAL DEVICES AND IMPLANTS
A Dissertation
Presented to
the Graduate School of
Clemson University
In Partial Fulfillment
of the Requirements for the Degree
Doctor of Philosophy
Bioengineering
by
Rohan Satishkumar
May 2011
to
Dr. Alexey Vertegel, Committee Chair
Dr. Naren Vyavahare
Dr. Ken Webb
Dr. Antony Guiseppi-Elie
i
ABSTRACT
The frequency of S. aureus infection and subsequent biofilm formation associated with
vascular catheterization has been increasing in recent years and often begins as a local
colonization at the site of the catheter insertion. Antimicrobial enzymes and peptides, which are
effective against a broad range of pathogens and low rates of resistance, have attracted attention
as promising alternative candidates in treatment of infections caused by antibiotic resistant
bacteria. The use of nanoparticles as carriers for enzymes, in addition to their size, charge, high
surface area per volume etc. offers targeted delivery of enzymes to pathogenic bacteria. We
proposed to use nanoparticles as surfaces for targeted delivery of antibacterial enzymes and as
„surrogate‟ surface coatings on indwelling central venous catheters (CVCs) to inhibit bacterial
colonization and subsequent biofilm formation.
It was shown that nanoparticle charge can be used to enhance delivery and increase
bactericidal activity of an antimicrobial enzyme, lysozyme. In the case of bacterial lysis assay
with a Gram-positive bacterium Micrococcus lysodeikticus, activity of lysozyme conjugated to
positively charged nanoparticles was approximately twice as high as that of free lysozyme. This
was believed to occur through charge-directed targeting of enzyme-nanoparticle conjugates to
negatively charged bacterial cell walls through enhanced electrostatic interactions. In a clinically
more relevant model, we studied antimicrobial activity of lysostaphin against S. aureus for both
lysostaphin-coated and lysostaphin-antibody coated nanoparticle conjugates at different enzyme:
antibody: nanoparticle ratios. At the highest antibody loading, bacterial lysis rates for antibodylysostaphin-coated samples were significantly higher than for plain lysostaphin-coated samples
and free enzyme due to multiple-ligand directed targeting of antibody molecules to bacterial cell
walls (p<0.05).
ii
We also performed in vitro experiments to evaluate the inhibition of bacterial colonies
adhering to a surface. Bacterial infections by S. aureus strains are among the most common
postoperative complications in surgical hernia repair with synthetic polymer meshes. Colony
counting data from the broth count (model for bacteria in wound fluid) and wash count (model
for colonized bacteria) for the enzyme-coated samples showed significantly decreased number of
colony forming units (CFU) when compared to uncoated samples (p< 0.05). A pilot in vivo study
showed a dose-dependent anti-S. aureus efficacy of lysostaphin-coated meshes in a rat model.
Finally, we observed that that coating of nanoparticles overall did not significantly
improve binding yield, leaching, durability and antibiofilm activity of enzymes adsorbed on
catheter segments (p>0.05). Alternatives to coating catheter surfaces using covalent chemistry
through functional groups on nanoparticles either directly or through appropriate crosslinkers
could result in significantly higher enzyme loadings, better stability and long term durability for
future applications. The approach developed here is universal and can potentially be used for
treatment of other medical device-associated infections. Moreover, use of antibacterial enzymeNP conjugates can eventually be expanded for intravenous administration, which will further
broaden its range of application.
iii
DEDICATION
I would like to dedicate this work to my parents who have always been there for me with their
unwavering support, guidance and strength. Thank you for all your love and prayers!
iv
ACKNOWLEDGEMENTS
I would like to sincerely thank my advisor, Dr. Alexey Vertegel, for his guidance and
patience through the past 5.5 years of my graduate studies at Clemson University. His high
expectations, continuous support and mentorship are greatly appreciated. I am grateful to the
other committee members, Dr. Naren Vyavahare, Dr. Antony Guiseppi-Elie and Dr. Ken Webb
for their time and active support in the completion of this dissertation. I would like to thank Dr.
Dan Simionescu and Dr. Martine Laberge for their support and guidance from the very
beginning of my graduate study.
A special thanks to the all the members of the Nanobiomaterials laboratory for being
there during my study at Clemson, sharing all those great moments and engaging in all the
discussions with each and every one of you has been memorable. I would like to acknowledge
Gary Thompson, Sriram Sankar and Vladimir Reukov specifically for their help and support. I
would also like to thank Sravya Durugu for her encouragement and constant support. Dr. David
Bruce and Iris Cordova (BET study), Dr. Meredith Morris (Flow cytometry), Dr. Terry Bruce
(Confocal microscope), Dr. Joann Hudson, Donald Musvee and Dayton Cash (SEM imaging),
Dr. Yuliya Yurko and Dr. Todd Heniford ( In vivo studies at Carolinas Medical Center). To all
the staff in the Bioengineering department for their help and cheerful disposition – Thank you so
much! I would like to personally thank all my friends and colleagues who have studied with me
during my years at Clemson for their help and support.
Finally, I would like to acknowledge the financial support and assistance provided by
Carolinas Medical Center, ViMedrx and the Department of Bioengineering.
v
TABLE OF CONTENTS
TITLE ......................................................................................................................................... i
ABSTRACT .............................................................................................................................. ii
DEDICATION .......................................................................................................................... iv
ACKNOWLEDGEMENTS.........................................................................................................v
TABLE OF CONTENTS .......................................................................................................... vi
LIST OF FIGURES .................................................................................................................. ix
LIST OF TABLES ...................................................................................................................xiv
CHAPTER ONE .........................................................................................................................1
INTRODUCTION ...................................................................................................................1
CHAPTER TWO ........................................................................................................................4
LITERATURE REVIEW ........................................................................................................4
2.1. Bacteria .........................................................................................................................4
2.2. Antibiotic resistance .................................................................................................... 10
2.3. Antimicrobial proteins and peptides ............................................................................ 16
2.4. Enzyme immobilization ............................................................................................... 24
2.5. Nanotechnology for enzyme immoblization ................................................................ 32
2.6. Nanoparticles .............................................................................................................. 35
2.7. Enzyme-nanoparticle conjugates as antibacterials – Targeted delivery......................... 41
2.8. Enzymes as antibacterial coatings on medical devices and implants............................. 43
CHAPTER 3 ............................................................................................................................. 45
SPECIFIC AIMS AND SIGNIFICANCE .............................................................................. 45
3.1. Clinical significance .................................................................................................... 45
3.2. Specific aims ............................................................................................................... 48
vi
CHAPTER 4 ............................................................................................................................. 53
CHARGE-DIRECTED TARGETING OF ANTIMICROBIAL ENZYME-NANOPARTICLE
CONJUGATES ..................................................................................................................... 53
4.1. Introduction ................................................................................................................. 53
4.2. Materials and methods................................................................................................. 54
4.2.2. Methods ................................................................................................................... 56
4.3. Results ........................................................................................................................ 63
4.4. Discussion ................................................................................................................... 73
CHAPTER 5 ............................................................................................................................. 77
ANTIBODY-DIRECTED TARGETING OF LYSOSTAPHIN ADSORBED TO
POLYLACTIDE NANOPARTICLES INCREASES ITS ANTIMICROBIAL ACTIVITY
AGAINST S.AUREUS IN VITRO ........................................................................................ 77
5.1. Introduction ................................................................................................................. 77
5.2. Materials and methods................................................................................................. 79
5.3. Results and Discussion ................................................................................................ 85
CHAPTER 6 ........................................................................................................................... 100
EVALUATION OF ANTIMICROBIAL ACTIVITY OF LYSOSTAPHIN-COATED
HERNIA REPAIR MESHES ............................................................................................... 100
6.1. Introduction ............................................................................................................... 100
6.2. Materials and methods............................................................................................... 102
6.3. Results ...................................................................................................................... 110
6.4. Discussion ................................................................................................................. 120
CHAPTER 7 ........................................................................................................................... 125
APPLICATION OF ENZYMES AS ANTIBACTERIAL COATINGS ON PLAIN AND
NANOPARTICLE COATED CATHETER SEGMENTS.................................................... 125
7.1. Introduction ............................................................................................................... 125
vii
7.2. Materials and methods............................................................................................... 127
7.3. Results and Discussion .............................................................................................. 139
7.4. Conclusion ................................................................................................................ 163
CHAPTER 8 ........................................................................................................................... 165
CONCLUSIONS AND FUTURE RECOMMENDATIONS ................................................ 165
8.1 Conclusions ................................................................................................................ 165
CHAPTER 9 ........................................................................................................................... 169
CHAPTER 10 ......................................................................................................................... 177
References ........................................................................................................................... 177
viii
LIST OF FIGURES
Page
2.1 A) Biofilm growing on a creek and B) S .aureus biofilm growing on a catheter segment.......5
2.2 Bacterial cell walls for Gram positive and Gram negative bacteria…………………………..7
2.3 Alternatives to antibiotics - AQUACEL® Ag..........................................................................14
2.4 Schematic of affinity enzyme immobilization through the Avidin-Biotin system…………..30
2.5 Nanotechnology for enzyme immobilization - Basic structures of nanoscale biocatalysts.....34
2.6 Reaction scheme showing different covalent immobilization techniques of proteins on
polymer nanoparticles through different functional groups ………………..................................38
4.1 Schematic of lysozyme conjugation to nanoparticles..............................................................60
4.2 Efficiency of covalent attachment of lysozyme to polystyrene latex nanoparticles for
samples prepared by (A) direct conjugation and (B) conjugation via PEG spacers......................64
4.3 Results of bacterial lysis assay analyzing effect of enzyme activity for samples directly
conjugated or through PEG spacers purified by centrifugation or dialysis...................................66
4.4 Results analyzing effect of control nanoparticles on bacterial lysis…………………………68
4.5 Tapping mode AFM images comparing the degree of aggregation for samples purified
by dialysis and centrifugation........................................................................................................69
ix
List of Figures (continued…………)
4.6 Lysozyme activity assay with low molecular weight substrate PNP-(GlnAc)5
(A) Direct attachment and (B) attachment via PEG spacer……………………………………...70
4.7 Cartoon showing charge-directed targeting of lysozyme to bacterial cells………………….74
5.1 Schematic showing adsorption of lysostaphin to PLA nanoparticles………………………..86
5.2 Binding yield of A) Alexafluor 594 labeled lysostaphin to PLA NPs alone or in the
presence of S. aureus antibody and B) Alexafluor 350 labeled S. aureus antibody on
the PLA NPs……………………………………………………………………………………..87
5.3 Fraction of leached lysostaphin activity for samples with highest lysostaphin and
antibody coatings………………………………………………………………………………...92
5.4 Bacterial lysis assay comparing activity of lysostaphin conjugated to NPs and free
enzyme…………………………………………………………………………………………...96
5.5 (A) Kinetic constant of bacterial lysis plotted as function of adsorbed lysostaphin
molecules per NP and (B) Kinetic constant of bacterial lysis plotted as function of
adsorbed antibody molecules per NP.............................................................................................98
6.1 Binding yield for Alexa 594 labeled lysostaphin on Ultrapro mesh………………………..110
x
List of Figures (continued….)
6.2 Standard curve plotting the mean rate of lysis (OD per min) against the different
known lysostaphin concentrations……………………………………………………………..112
6.3 Leaching of adsorbed lysostaphin as a function of time for mesh samples treated by
different initial enzyme concentrations…………………………………………………………113
6.4 Turbidimetric enzyme activity assay for the different lysostaphin-coated hernia mesh
samples………………………………………………………………………………………….114
6.5 Fraction of leached sample activity for the aliquots collected at different time
intervals compared against total mesh activity for different coating concentrations…………...116
7.1 Cartoon showing a single lumen catheter segment before and after coating with
nanoparticles……………………………………………………………………………………141
7.2 Binding yield comparing lysostaphin and DispersinB on plain and NP coated catheter
segments either alone or through coadsorption………………………………………………...142
7.3 SEM images of plain and PLA NP coated Polyurethane catheter segments at different
magnifications…………………………………………………………………………………..145
List of Figures (continued….)
xi
7.4 Activity assay data comparing the kinetic constant of bacterial lysis against the
highest and lowest lysostaphin coating concentrations for plain and NP coated
catheter segments……………………………………………………………………………….148
7.5 Activity assay data comparing the kinetic constant of hydrolysis against the highest
and lowest DisperinB coating concentrations for plain and NP coated catheter segments..…...150
7.6 Leaching of enzyme as a function of time for plain and NP coated catheter segments
at different Coating concentrations for lysostaphin and DispersinB ………………………… .151
7.7 Direct cytotoxicity results of MTT assay that shows the percentage of cell viability after
48 h of all the catheter samples coated with the highest coating concentration that were
placed in contact with 3T3 fibroblasts………………………………………………………….154
7.8 Percentage of Biomass on catheters for catheter samples coated with highest enzyme
concentration…………………………………............................................................................156
7.9 Digital camera images showing biofilm mass on control and sample catheter
segments………………………………………………………………………………………...158
7.10 Mean rate constants of bacterial lysis for lysostaphin coated on both plain and
NP coated catheter samples…………………………………………………………………….160
xii
List of Figures (continued……………)
7.11 Mean rate constants of hydrolysis for DispersinB coated on plain and NP coated
catheter samples………………………………………………………………………………...162
xiii
LIST OF TABLES
Page
2.1 List of some of the antibacterial peptides and proteins and their enzymatic mode of
action……………………………………………………………………………………………22
4.1 Properties of Polystyrene nanoparticles with different functional groups…………………..56
4.2 Initial rates of the bacterial lysis study comparing samples conjugated directly or
through PEG spacers purified by dialysis and centrifugation……………………………………67
4.3 Rates of hydrolysis using low molecular weight substrate PNP-(GlnAc)5 comparing
activity of lysozyme conjugated directly or through PEG spacers………………………………71
4.4 Zeta potential analaysis………………………………………………………………………72
5.1 Characteristics of samples with different nanoparticle (NP): lysostaphin (LS)
: antibody (Ab) ratios…………………………………………………………………………….89
5.2 Size and zeta potential of all enzyme-NP conjugates and controls.........................................90
5.3 Bacterial turbdimetric assay comparing activity of enzyme-NP conjugates against
free enzyme………………………………………………………………………………………93
6.1 Specific enzyme activity of samples with different enzyme coating concentrations………115
List of Tables (continued…………)
xiv
6.2 Effect of lysostaphin coating concentration on log reduction of colonies for the broth
supernatant and wash count……………………………………………………………………118
6.3 In vivo rat study results……………………………………………………………………..120
xv
CHAPTER ONE
INTRODUCTION
My graduate research at the department of Bioengineering in Clemson focused on
the specific application of Nanotechnology for targeted delivery of antibacterial enzymes
and for enzyme-based coatings on medical devices and implants. Using different model
studies, further discussed in various chapters of my dissertation, the goal of my research
was to use nanotechnology to address specific issues and drawbacks involved in the
delivery of antibacterial enzymes as infections and as coatings on implants to prevent
bacterial colonization and biofilm formation. Chapter 2 of this dissertation presents a
detailed literature review on the current problems associated with treatment of nosocomial
infections with antibiotics specifically focusing on multidrug resistant S. aureus, enzymes
and peptides as alternatives to antibiotics and its drawbacks and finally the need for
enzyme immobilization using nanotechnology through various techniques.
Chapter 3 lists the specific aims and goals of our research project by using model
studies to test our hypothesis and fundamentally address specific issues involved in the
application of enzymes for - a) targeted delivery using polymer nanoparticles (NPs) and b)
as coatings on hernia repair polymer meshes and Central venous catheter segments.
Our first model study in Chapter 4 aimed at comparing the influence of
nanoparticle charge on antibacterial activity of enzyme-nanoparticle conjugates.
Specifically, we covalently attached a model antibacterial enzyme hen egg lysozyme to
two types of polystyrene latex nanoparticles: positively charged, containing aliphatic
1
amine surface groups, and negatively charged, containing sulfate and chloromethyl surface
groups. We tested lysozyme activity against its substrate, Micrococcus lysodeikticus and
compared antibacterial activity of free enzyme against enzyme conjugated to NPs. We
further evaluated the method of enzyme purification and effect of PEG spacers on
antibacterial activity of enzyme-NP conjugates
In a clinically more relevant model (Chapter 5), we studied whether co-attachment
of different ligands such as an antibody (against the pathogen) along with antibacterial
enzymes to NPs enhances in vitro antimicrobial activity of the enzyme. We compared the
anti S. aureus activity of lysostaphin adsorbed on the surface of PLA (poly (lactic acid))
nanoparticles to that of the free enzyme at different initial enzyme: NP molar ratios. A
rabbit polyclonal S. aureus antibody was coadsorbed on PLA nanoparticles along with
lysostaphin to evaluate the effect of delivery based on antibody-directed targeting.
Bacterial infections by antibiotic-resistant S. aureus strains are known to be one of
the most common postoperative complications in surgical hernia repair. In our third model
study, we evaluated the in vitro antimicrobial susceptibility of S. aureus to lysostaphincoated hernia repair polymer meshes and study the effect of enzyme coating concentration
on antimicrobial activity. In vivo antimicrobial efficacy of lysostaphin-coated meshes was
further evaluated by our clinical collaborators at Carolinas Medical Center in a rat hernia
repair model (Chapter 6).
It is well known that bacteria can either colonize the implant surface or exist in a
planktonic free floating form. Both these chapters (4 and 5) provided insight into taking
advantage of special properties of NPs such as size, charge, surface area etc. to improve
2
functionality of enzyme-NP conjugates, specifically towards targeting to planktonic cell
suspensions.
Colonization is the first stage of device associated-infection. Bacteria in colonies
(biofilms) present more danger than planktonic bacteria and are also less susceptible to
treatment by antimicrobial agents. Although coating medical devices with plain
antibacterial enzymes could work just as effectively, we hypothesized that coating catheter
surfaces with nanoparticles initially may be advantageous due to larger surface area for
effective enzyme coating, higher stability, durability and low enzyme leaching. Thus the
goal of our final model study was the characterization and comparison of the in vitro
performance for plain and NP coated catheter segments using antimicrobial and
antibiofilm enzymes (Chapter 7). Coating was achieved via adsorption of two model
enzymes, DispersinB (antibiofilm) and lysostaphin (antimicrobial) on the surface of plain
and poly ((lactic acid)) nanoparticle coated polyurethane catheter segments. A series of
experiments evaluating enzyme binding efficiency, activity, leaching, cytotoxicity,
antibiofilm efficacy and durability were performed to compare plain and NP coated
catheter segments.
Chapters 8 and 9 provide a conclusion to our studies and future recommendations
to design more effective antibacterial surface coatings on devices and implants. By
manipulating the available technology today, we do not just limit ideas only to
antibacterial coatings but also hopefully expand to a wider range of biomedical
applications.
3
CHAPTER TWO
LITERATURE REVIEW
2.1. Bacteria
Bacteria were one of first types of cell to evolve in nature. They have been
described as a „prokaryotic‟ organism because their cells lack a membrane bound nucleus
and their genetic material is typically a single circular chromosome located in the
cytoplasm in an irregularly shaped body called the nucleoid [1]. Despite their small size,
bacteria have an enormous range of metabolic capabilities, and a lot of them can be found
in some of the extreme environments on earth. All of the various surface components of a
bacterial cell are important in its ecology since they mediate contact of the bacterium with
its environment. It must use its own surface components to assess the environment and
respond ways that supports its own existence and survival in that environment. The
surface properties of a bacterium are determined by the molecular composition of its
membrane and cell wall. In most prokaryotes, the primary function of the cell wall is to
protect the cell from internal turgor pressure caused by the much higher concentrations of
proteins and other molecules inside the cell compared to its external environment.
Most bacteria also contain some sort of a polysaccharide layer outside of the cell
wall or outer membrane. In a general sense, this layer is called a capsule. A true capsule is
a discrete detectable layer of polysaccharides deposited outside the cell wall. A type of
capsule found in bacteria called a glycocalyx is a thin layer of polysaccharide fibers which
is mostly observed on the surface of cells growing in nature. Capsules are known to
protect bacteria from engulfment by phagocytes and from attack by antimicrobial agents
4
[2]. In nature, and in many medical conditions, colonies of bacteria live in a biofilm,
primarily composed of an outer slimy layer.
These films can range from a few micrometers to up to tens of centimeters in
thickness. Bacteria living in biofilms display a complex arrangement of cells and
extracellular components, forming secondary structures such as microcolonies, through
which there are networks of channels to enable better diffusion of nutrients [3, 4]. Also,
bacteria communicate with each other through an internal cell signaling process called
Quorum sensing, thus regulating a host of cell metabolic activities, similar to a
communication network. In natural environments, such as soil or the surfaces of plants,
the majority of bacteria are bound to surfaces in biofilms [5]. Biofilms are also important
for chronic bacterial infections and infections of implanted medical devices, as bacteria
protected within these structures are much harder to kill than individual bacteria [6].
A
B
Figure 2.1. A) Biofilm growing on a creek (http://toxics.usgs.gov) and B) S .aureus
biofilm growing on a catheter segment ((http://en.wikipedia.org)
5
2.1.1. Bacterial cell wall
The bacterial cell wall has been one of the important clinical targets for antibiotic
agents since the first use of penicillin in World War II. Bacterial cell walls are made of
peptidoglycan (also called murein), which is made from polysaccharide chains crosslinked by unusual peptides containing D-amino acids [7]. Bacterial cell walls are different
from the cell walls of plants and fungi which are made of cellulose and chitin,
respectively[8]. Although the primary function of the cell wall is to provide a rigid
exoskeleton for protection against both mechanical and osmotic lysis [9], it also serves as
an attachment site for proteins that interact with the bacterial environment.
Based on the structure of their cell wall, bacteria are classified into two types – a)
Gram-positive and b) Gram-negative bacteria. The names originate from the reaction of
cells to the Gram stain, a test long-employed for the classification of bacterial species.
Gram-positive bacteria have a thick mesh-like cell wall made of peptidoglycan, which
stain purple and Gram-negative bacteria have a thinner layer, which stain pink. Although
the vast majority of bacteria adhere to the color differentiation of the Gram stain, some
bacteria do not; these are called Gram-variable bacteria [10].
The cell wall of Gram-positive bacteria is a peptidoglycan macromolecule with
attached accessory molecules such as teichoic acids, teichuronic acids, polyphosphates, or
carbohydrates [9, 11]. Gram-positive bacteria have much thicker peptidoglycan layer than
their Gram- negative counterparts and no external outer membrane. They lack a distinct
periplasmic space, which is usually defined as the region between the cytoplasmic
membrane and the outer membrane. The glycan strands of the cell wall consist of the
repeating disaccharide N-acetylmuramic acid-(b1-4) - N-acetylglucosamine (MurNAc6
GlcNAc) [12, 13]. Glycan strands vary in length and are estimated to contain 5 to 30
subunits, depending on the bacterial species [14-16].
The muramic acid is a unique substance associated with bacterial cell walls and
these chains are cross-linked by short polypetide chains consisting of both L- and Daminoacids. Wall peptides are cross-linked with other peptides that are attached to a
neighboring glycan strand [17-20], thereby generating a three-dimensional molecular network that surrounds the cell and provides the desired exoskeletal function [21]. The
teichoic acids are polyols consisting predominantly of glycerol, ribitol and mannitol,
covalently linked to the peptidoglycan through phosphodiester bonds and can be
substituted by sugars, aminosugars or D-alanine residues. These anionic polymers account
for 10–60% (by weight) of the bacterial cell wall, with the relative amount depending on
the culture conditions [22, 23].
7
Figure 2.2. - Bacterial cell walls for Gram positive and Gram negative bacteria
(figure modified from Cabeen et al; 2005 [24].
Although they are structurally diverse[25, 26] ,the negative charge of these anionic
polymers mainly originates from phosphate or carboxyl groups, and they may also
elaborate acidic side chains containing glycerol, phosphate, organic acids (e.g. pyruvic and
succinic acid), or sulphates. They play a very crucial role in the binding of divalent
cations, balance of metal ions for membrane functionality, folding of extracellular
metallo-proteins and formation of a barrier to prevent diffusion of nutrients and
metabolites [23, 27-30]. In addition, the peptidoglycan is a well-known target for almost
all clinically useful antibiotics that inhibit bacterial cell wall synthesis [31]. Being an
essential and unique cell wall component of virtually all bacteria and not eukaryotic cells,
it is an excellent target for recognition by the innate immune system of the host [31, 32].
The cell walls of Gram negative bacteria are composed of three layers, an inner
bilayer phospholipid membrane, a thin peptidoglycan layer, and the outer bilayer
membrane, composed of lipopolysaccharides [31, 32]. Along with the plasma membrane
and the cell wall (outer membrane, peptidoglycan layer, and periplasm) constitute the
Gram-negative envelope [33, 34].The envelope of E-coli and other Gram-negative bacteria
contains two distinct lipid bilayers that are separated by the peptidoglycan-containing
periplasmic space that represent the inner membrane (IM) and the outer membrane (OM)
respectively [35, 36]. The inner membrane is composed of phospholipids in both its inner
and outer leaflets as well as integral and peripheral membrane proteins. It carries out a
variety of functions typically assigned to both the plasma membrane and specific
organelles in higher organisms that includes cell signaling, biosynthesis, electron transport
8
and ATP synthesis [37] . The OM is a unique asymmetric lipid bilayer consisting of an
inner face of phospholipids and an outer face of lipopolysaccharide (LPS). The outer
membrane (OM) plays an important role in nutrient uptake but in addition provides the
organism with a remarkable permeability barrier, conferring resistance to a variety of
agents including antibiotics [38].
2.1.2 Staphylococcus aureus
Staphylococcus aureus is facultative anaerobic Gram-positive coccus which occurs
singly, in pairs, and in irregular clusters. It is nonmotile, non-spore forming, catalase and
coagulase positive bacteruim. The term Staphylococcus is derived from the Greek term
staphyle, meaning "a bunch of grapes." It can colonize and infect both healthy,
immunologically competent people in the community and hospitalized patients with
decreased host defenses [39]. It‟s repertoire of virulence factors is extensive, with both
structural and secreted products playing a role in the pathogenesis of infection. Upregulating the production of virulence factors enables it to persist in the blood stream,
adhere and colonize the skin and mucosa, evade the host immune response, form
protective biofilms through quorum sensing and also develop resistance against a wide
range of antibiotics.
S. aureus has an intrinsic ability to form biofilms on damaged tissue, prosthetic
materials such as medical devices and heart valves, thereby making it much more difficult
to treat than planktonic bacteria. The biofilm matrix is usually composed of biopolymers
such as polysaccharides, proteins and lipids which build a firm consistency thus providing
protection against the immune cells and penetration of antimicrobial agents [40]. It is also
9
known that cells living in biofilms display a complex arrangement of various extracellular
components and form secondary structures such as microcolonies and small-colony
variants (SCVs) that demonstrate almost complete resistance against conventional
antibiotics [41].
S. aureus biofilm formation involves two stages – initial adhesion of cells to a
surface mediated by a number of surface proteins that serve as an anchor, followed by
production of various extracellular factors that enable cell multiplication and maturation
into a structured community. „Quorum Sensing‟ or intracellular signaling between cells
via the accessory gene regulatory (agr) system has also been strongly implicated in the
pathogenesis of biofilm-associated S. aureus infections [42]. The (agr) quorum-sensing
system decreases the expression of several cell surface proteins and increases expression
of many secreted virulence factors based on bacterial density [43]. It regulates the
detachment of cells from the established biofilm and then allows them to colonize new
sites or even enter the blood stream. The rates of infections caused by both community
and hospital-acquired strains are increasing steadily and subsequent treatment of these
infections is also becoming more difficult because of the increasing prevalence of
multidrug-resistant strains [44].
2.2. Antibiotic resistance
Antibiotics have traditionally been used in modern medicine as chemotherapeutic
agents to treat infections thus becoming indispensable in the modern health care system.
However the rates of multi-antibiotic resistance among bacteria that infect wounds are
constantly on the rise and surgical site infections have become a substantial burden of
10
disease for patients and health services [45-48]. The US Centers for Diseases Control and
Prevention (CDC) estimates that about 500,000 surgical site infections occur annually in
the United States [48-51]. There are several factors contributing to surgical site infections
namely the inoculum of bacteria introduced into the wound during the procedure, the
unique virulence of contaminants, the microenvironment of each wound, and the integrity
of the patients host defense mechanisms [48, 50].Consequently, rapid control of wound
infections and monitoring of prophylactic and therapeutic strategies have recently been
proposed [49, 51] . Antibiotic resistance in nosocomial infections is thus an everincreasing problem.
Antibiotics have been shown to act in different ways, either by targeting the
pathogen„s physiology directly or by disrupting the cellular structure of bacterial cells.
There are 4 major modes of action: (1) interference with cell wall synthesis, (2) inhibition
of protein synthesis, (3) interference with nucleic acid synthesis, and (4) inhibition of a
metabolic pathway. Antibacterial drugs that work by inhibiting bacterial cell wall
synthesis include the b-lactams, such as the penicillins, cephalosporins, and carbapenems
[52, 53]. They inhibit synthesis of the bacterial cell wall by interfering with the enzymes
required for the synthesis of the peptidoglycan layer [52]. Macrolides, aminoglycosides
and tetracyclines produce their antibacterial effects by inhibiting protein synthesis by
binding to different subunits of the Ribosome [52, 53]. Fluoroquinolones exert their
antibacterial effects by disrupting DNA synthesis and causing lethal double-strand DNA
breaks during DNA replication [54]. Antibacterial drugs such as folic acid analogues
inhibit different metabolic pathways.
Disruption of the bacterial membrane is
11
characteristic of polymyxins that exert their inhibitory effects by increasing bacterial
membrane permeability, causing leakage of bacterial contents [55].
Several mechanisms of antimicrobial resistance occur in different bacterial genera.
First, the organism may acquire genes encoding enzymes, such as β-lactamases, that
destroy the antibacterial agent before it can have an effect. Susceptible bacteria can
acquire resistance to an antimicrobial agent via new mutations [56]. Such spontaneous
mutations may cause resistance by altering the target protein to which the antibacterial
agent binds by modifying or eliminating the binding site (e.g., change in penicillin-binding
protein 2b in pneumococci, which results in penicillin resistance), upregulating the
production of enzymes that inactivate the antimicrobial agent (e.g., erythromycin
ribosomal methylase in staphylococci) and altering an outer membrane protein channel
that the drug requires for cell entry or by upregulating pumps that expel the drug from the
cell [56]. Therefore strains of bacteria carrying resistance conferring mutations are
selected by continuous antibiotic use, which kills the susceptible strains but allows the
newly resistant strains to survive and grow.
Bacteria also develop resistance through the acquisition of new genetic material
from other resistant organisms. Mutation and selection, together with mechanisms of
genetic exchange, enable many bacterial species to adapt quickly to the introduction of
antibacterial agents into their environment [57]. Resistance to multiple antibiotics is
increasingly reported in a number of Gram-negative pathogens, especially the
Enterobacteriaceae [58-61], E-coli [62, 63], Pseudomonas aeruginosa [64-66]etc.
Multidrug resistance occurs as a result of accumulation of multiple mutations and/or
resistance genes, though single mutations can also promote multidrug resistance
12
Also, drugs come with adverse effects and antibiotics are no exception. Common
side-effects are gastrointestinal symptoms, skin rashes, and thrush. A common problem in
clinical practice is in determining the required antibiotic to treat infection in a patient who
has been labeled as being allergic to the antibiotic. In many cases, such patients are
prescribed antibiotics that are less effective or more toxic, have a broader spectrum, or are
more expensive than the drug of choice for their condition. The occurrence of overgrowth,
or infection, as a direct result of antibiotic consumption is also less well understood by
physicians. An antibiotic will inhibit, or kill, a whole range of bacteria, which creates
space on mucosal and other surfaces for other organisms to proliferate [67]. These
survivors might show various rates of acquired resistance, are more difficult to treat and
almost impossible to eradicate in some patients [68].
2.2.1 Alternatives to antibiotics
There is therefore an urgent need for the use of alternative approaches to
antibiotics. Widespread application of an effective topical antimicrobial agent
substantially reduces the microbial load on an open wound surface and reduces the risk of
infection [69-71]). However the selection of topical antimicrobial therapy should be based
on the agent's ability to inhibit the microorganisms recovered from wound cultures and
monitoring of the nosocomial infections acquired in hospitals and medical centers. For e.g.
several topical antimicrobials have also been used for topical burn therapy, including
gentamicin sulfate (0.1% water-soluble cream), betadine (10% povidone-iodine ointment),
bacitracin-polymyxin ointment, and nitrofurantoin [72].
13
Fig 2.3. Images of AQUACEL® Ag - ionic silver (Convatec, NJ, USA). This
material provides an effective barrier to bacterial penetration to help reduce
infection. AQUACEL® Ag is indicated for use on acute and chronic wounds
including burns, surgical wounds, diabetic foot ulcers, pressure ulcers and leg
ulcers. The power of ionic silver in AQUACEL ® Ag kills a broad spectrum of
wound pathogens in the dressing that can cause infection - including
Staphylococcus aureus, methicillin-resistant Staphylococcus aureus (MRSA),
vancomycin-resistant Enterococcus (VRE), and other pathogens (Figure source :
http://www.convatec.com).
However, these compounds are no longer used extensively because significant
resistance has developed and/or they have been shown to be toxic or ineffective at
controlling localized burn wound infections [73]. Toxicity from silver is observed in the
form of argyria or skin discoloration, usually when there is a large amount of silver ions
are used in the wound dressing. However, a delayed wound healing response has also been
observed in a clinical setting. It has been shown that exposure of human dermal fibroblasts
and epidermal keratinocytes to silver sulfadiazine in vitro
produced a significant
reduction in cell proliferation and increased cytotoxicity [74, 75].
It has also been
reported to inhibit collagen synthesis in human rheumatoid synovial cells in culture [76].
14
Novel vaccines were developed as they do not suffer the problem of resistance
instead work to enhance the body's natural defenses. Nevertheless, it is possible that new
strains may evolve that escape immunity induced by vaccines. Vaccines also do not
guarantee complete protection from a disease. This may be due to a lowered immunity in
general (diabetes, steroid use, HIV infection) or because the host's immune system does
not have B-cells capable of generating antibodies to that antigen.
Bacteriophage therapy has been recently studied as an emerging alternative to
antibiotics [77]. Phages were found to possess several potential disadvantages when
compared with antibiotics. Although they are specific for the target bacterium, which
reduces the disruption to the natural flora of the host, the need for isolation of a phage
specific to the target bacteria leads to the increased costs and development time. Also,
phages that are injected into the bloodstream are immunogenic [78]. Some of them are
cleared very quickly and, after a certain period, antibodies against the phages are produced
by the body [78]. Funding for phage therapy research and clinical trials is also generally
insufficient and difficult to obtain, since it is a lengthy and complex process to patent
bacteriophage products. Phage therapy has been used publicly for many years; therefore,
the technique itself would not be patentable. Additionally, it is unlikely that it would be
possible to patent individual phage, and the wide range of phage targeting each species of
bacteria would mean that isolating a range of unique phage would be relatively
straightforward.
15
2.3. Antimicrobial proteins and peptides
Use of proteins and peptides as antimicrobial agents is inspired from nature and
has recently attracted much attention as an antibiotic-free approach to treat bacterial
infections [79-81]. Naturally occurring antimicrobial peptides and enzymes have are
characterized by higher activity, broad antimicrobial spectrum, and the low rate of
resistance to these agents by bacteria. They may be expressed constitutively or sometimes
inducibly in response to certain specific pathogenic challenges [82].
The current promising candidates of antibacterial enzymes are functionally
classified as either hydrolases or oxidoreductases [83]. Hydrolases are bacteriolytic
enzymes that function by degrading key structural components of the cell walls of
bacteria. Peptidoglycan is the major structural component of bacterial cell walls.
Degradation of bacterial cell walls results in lysis due to rupture of osmotic balance in the
cell. These enzymes are further grouped into three different classes –1) Nacetylhexosaminidases that catalyze the cleavage of the β (l-4)-glucosidic linkages in the
carbohydrate backbone of the peptidoglycan, 2) N-acetylmuramyl-L-alanine amidases that
catalyze the cleavage between the carbohydrate moiety and the peptide moiety of the
peptidoglycan, and 3) Endopeptidases that hydrolyze the peptide crosslinks of the
peptidoglycan [84]. Lysostaphin is an antibacterial enzyme which specifically cleaves
cross-linked pentaglycine bridges in the peptidoglycan of S. aureus, thereby hydrolyzing
the cell wall and lysing the bacteria.
Oxidoreductases on the other hand do not possess any antimicrobial activity by
themselves. However they exert their effect through the generation of reactive species that
are known to be toxic to bacteria. Glucose Oxidase, Lactoperoxidase and
16
Myeloperoxidase are some well-studied oxidoreductase systems [83, 85, 86]. These
enzymes (Peroxidases) are synthesized and secreted by ductal epithelial cells of the
mammary gland and other exocrine glands. They also probably provide protection against
microbial infection to the mucosal surfaces of eye, nose, mouth, trachiobronchial tree and
intestinal tract [87]. Both salivary peroxidase and myeloperoxidase, the two peroxidase
enzymes in human mixed saliva, have been purified but only for research purposes. Largescale purification from human saliva or from human polymorphonuclear leukocytes is
difficult and far too expensive for commercial purposes. Therefore, for commercial
purposes bovine lactoperoxidase (purified from milk or colostrum) has been used because
this enzyme is structurally and catalytically very close to human salivary peroxidase [87].
The lactoperoxidase system has been incorporated into commercial products such as tooth
pastes and mouthwashes like Biotene ®, BioXtra® and Oralbalance® [87]. It is also well
understood that bacteriolytic enzymes usually offer protection against only a few bacteria,
owing to their specificity. The oxidoredctuase systems may affect a broader spectrum of
microorganisms but suffer from the drawback of being dependent on the availability of
substrates for the generation of antimicrobial reaction products and toxicity towards
eukaryotic cells.
2.3.1 Antimicrobial peptides (AMPs)
These are a group of short 10–40-residue polypeptides that are part of the innate
immune system in many organisms including bacteria, insects, plants, and vertebrates[88].
In mammals, these peptides are present mainly in phagocytic cells and in mucosal
epithelial cells. In addition to their antimicrobial role, antimicrobial peptides (AMPs) also
17
serve as important molecules involved in inflammation, immune activation, and wound
healing [89-91].
In humans, three distinct groups of antimicrobial peptides are usually
distinguished: defensins, cathelicidins, and histatins [92, 93]. Histatins will not be
considered here because their primary function is antifungal, rather than antibacterial. A
characteristic feature of defensins (molecular weight 3-5 kDa) is the presence of six
cysteine residues forming three intramolecular disulfide bridges. Defensins are further
subdivided into - and -defensins based on the distribution of the cysteines and linkages
of disulfide bridges. Cathelicidins are a family of proteins containing an N-terminal
cathelin domain and a C-terminal cationic antimicrobial domain that becomes active after
cleavage [94]. Only one human cathelicidin is known. It is a cationic 18 kDa protein
precursor, hCAP18. Its mature antibacterial peptide named LL-37 is liberated through
cleavage by elastase and proteinase 3 [95]. LL-37 has a broad-spectrum antimicrobial
activity against a variety of Gram-positive and Gram-negative bacterial, fungal, and viral
pathogens.
Amphipathicity (presence of both hydrophilic and hydrophobic groups) and net
charge are characteristics conserved among many antimicrobial peptides [88]. All
antimicrobial peptides bear a large positive charge, which results in their preferential
targeting to bacteria, rather than to eukaryotic cells. It is generally understood that the
bactericidal activity of antimicrobial peptides is due to permeation of bacterial cell
membranes [88, 96]. It is also likely that antimicrobial enzymes and peptides may use
more than one mechanism of action, such as destabilization of the cell membrane
combined with inhibition of one or more intracellular targets [97]. This “multitarget”
18
mechanism hypothesizes that highly charged cationic peptides and enzymes can bind to
anionic molecules within the cytoplasm, such as nucleic acids or enzymes with ionic
surfaces, thus interfering with processes in which these molecules are involved.
2.3.2 Synergy
Interactions between antimicrobial enzymes and peptides may be synergistic,
additive, or antagonistic in nature. Because of their different modes of action,
antimicrobial enzymes and peptides may act cooperatively by simultaneously attacking
essential structures in microorganisms [98]. Synergy among individual peptides, was first
observed with frog peptides, including members of the dermaseptin family [99] and the ahelical peptides magainin and PGLa [100], and between b-defensins and the cationic
protein BPI [101].
The synthetic peptide LL-37 demonstrated antibacterial activity against a number
of Gram-negative and Gram-positive organisms including P. aeruginosa and was
synergistic with lactoferrin and lysozyme [102]. A synergistic antibacterial effect between
LL-37, β-defensins, and lysozyme was initially described by Nagaoka et al. [103]and later
comprehensively investigated by Chen et al [104]. They observed that the antibacterial
agents- hBDs (human beta defensins), LL-37 and lysozyme exhibit antimicrobial activities
synergistically against invading microorganisms, in particular S. aureus that frequently
colonizes the skin [104]. The synergistic effect found was strong enough to speculate that
defensins may not be able to act as antibacterial molecules by themselves, but only in
synergism with cathelicidins [103]. In another study, synergism in various combinations
of six antimicrobial factors, including lysozyme, lactoferrin, LL-37, HBD-1, HBD-2, and
19
secretory leukocyte protease inhibitor (SLPI), had been evaluated by Welsh and coauthors [105]. In this work, synergy of lysozyme with LL-37 has been confirmed, while
no synergy has been observed in the cases of lysozyme combined with either HBD-1 or
HBD-2.
Thus synergistic interactions appear to be important evolutionary survival
mechanisms that can be cleverly manipulated in the design of antibacterial combinations.
Combining
multi-targeting
functions
through
different
peptides
and
proteins
simultaneously ensures their antimicrobial action to a wide range of microorganisms. Due
to widespread antibiotic resistance, such multiple combinations, similar to that of an
antibiotic cocktail, are often required to overcome infectious diseases. Implementing such
synergistic principles into the design of next generation therapeutics could prove to be an
important determinant in the effectiveness of using an antibiotic-free approach.
2.3.3 Antibiofilm activity
The biofilm matrix is composed of an assortment of cells and various extracellular
polymers of different polysaccharides such as glucose, dextrans, cellulose, alginate etc.
and also of some glycoproteins [40]. Antimicrobial enzymes have been used in the
inhibition of biofilm formation. However due to the heterogeneity of these
polysaccharides, a combination of enzymes is necessary to achieve a synergistic effect for
biofilm inhibition. For instance, Pectinex UltraSP (a mixture of enzymes containing
cellulose, pectinase etc.) has been used for enzymatic degradation of S. aureus, P.
aeruginosa, P. fluorescens and S. epidermis biofilms with S. aureus being the most
sensitive [106].
20
Some of these polysaccharide-hydrolyzing enzymes are only known for dispersion
and inhibition of biofilms but do not have a significant bactericidal activity. For example
DispersinB
(patented
by
Kane
Biotech
Inc.),
a
naturally
occurring
N-
acetylglucosaminidase isolated from A. actinomycetemcomitan is a highly active and
stable glycoside hydrolase that specifically glycosidic linkages of poly-b-1, 6-Nacetylglucosamine in polysaccharide adhesins of bacteria needed for biofilm formation
[107, 108]. As DispersinB can only inhibit or disperse bacterial biofilms without affecting
their growth, it would be necessary to combine it with another antimicrobial in order to
inhibit the growth and proliferation of biofilm-embedded bacteria [107, 109]. When used
in combination with glucose oxidase; it increased the sensitivity of S. epidermidis biofilm
to glucose oxidase [110]. Johansen et al also similarly showed that combining
oxidoreductases such as glucose oxidase and lactoperoxidase with polysaccharidehydrolyzing enzymes resulted in bactericidal activity as well as removal of the biofilm
thus citing another example of a synergistic effect [106]. Apart from the
exopolysaccharide layer, extracellular DNA has also been as a target used to inhibit
biofilm formation. For e.g Bovine DNase1 was shown to suppress P. aeruginosa,
Streptococcus intermedius and S. mutans biofilm formation [111]. The antibiofilm activity
of these DNases involves degradation of extracellular DNA, which is an essential
component of biofilms.
21
Peptide and proteins
Source
Lysozyme
Human, Chicken
Lactoferrin
Human, Bovine
α and β defensins
Mammals;
analogues in
insects and fungi
LL-37
Human
Indolicidin
Bovine
Nisin
Bacteria
Magainin
Frog
Protegrin
Human, Porcine
Cecropin A
Silk moth
Proposed antibacterial mechanism
References
Enzymatic lysis of peptidoglycan layer; Non
enzymatic mode of action
[112, 113]
Sequestering iron essential for microbial respiration ;
Directly microbicidal due to its N-terminal cationic
fragment “lactoferricin.”
Cell membrane and intracellular targets, inhibits
macromolecular synthesis
[114, 115]
[116, 117]
Membrane permeabilization
[118, 119]
Inhibits macromolecular synthesis, Ca2_-calmodulin
interaction
[120, 121]
Membrane permeabilization; binds to lipid 2 of
peptidoglycan layer; reduction of ATP; collapse of
pH gradient
[122, 123]
Membrane permeabilization
[124, 125]
Very potent, membrane permeabilization
[126]
Membrane destabilizing
[127, 128]
Table 2.1. below shows list of some of the antibacterial peptides and proteins
2.3.4 Challenges associated with application of antibacterial proteins and peptides
Overall great enthusiasm exists regarding the prospect of developing antimicrobial
proteins and peptides as a new generation of „antibacterial agents‟ for treatment of
implant-associated infections. In spite of this, use of enzymes and peptides in a clinical
setting is associated with several potential drawbacks, which includes 1) high
22
manufacturing cost, especially of peptides, 2) short half life in vivo, 3) loss of activity in
physiological conditions (especially at high ionic strength, which may interfere with their
charge-directed targeting and mechanism of action), 4) unwanted systemic reactions
(aggregation, immunoreactivity), 5) interference with normal bacterial flora and even 6)
site-specific delivery that may arise when attempts are made to use them for targeted
delivery as antibacterial agents. It is also known that small molecules (lesser than 70kDa)
such as antimicrobial enzymes and peptides have a relatively short half-life in blood after
intravenous administration. Their rapid clearance from circulation may reduce its efficacy
as an antimicrobial. Also, some of the enzymes such as lysostaphin and DispersinB are
derived from bacterial species and could potentially show immunogenicity in humans
which further stimulates its clearance from the blood stream. This is mainly due to the
induction of a specific humoral response [129, 130]. Antibodies from the host might
neutralize these enzymes and compromise the efficacy of the drug and there could also be
other effects related to cross-reactivity with other autologous proteins. Thus, their short
circulating half-life cannot be effectively countered by increasing the amount or frequency
of dosage. There are also a number of MMPs (Matrix Metalloproteinases), Serine
proteases etc. that could also result in protein degradation through hydrolysis. More
expensive strategies such as synthesis of recombinant protein that is resistant to
proteolytic degradation have been reported [131].
Several research-related issues also must be resolved for better understanding of
the fundamental aspects of the processes associated with clinical applications- 1)
Development of standardized techniques to assess the activity of enzymes and peptides 2)
Targeting of agents to the site of action. 3) Addressing toxicity in more detail, for e.g. the
23
positive charge of AMPs may result in their strong interactions with anionic components
of both bacterial and host cells. 4) Understanding of the role and expression of
antimicrobial enzymes and peptides in both health and disease. Focusing on these issues is
expected to lay the foundation for the future applications of enzymes and peptides as
highly effective antibacterial agents for treatment of device-related infections.
2.4. Enzyme immobilization
Immobilization of enzymes on surfaces is of great importance for a wide range of
biomedical applications because of their recyclability, decrease in systemic toxicity,
carrier specificity and improvement in enzyme stability [132]. Enzymes are highly
specific and efficient biocatalysts with a wide range of biotechnological and biomedical
applications [133, 134]. There are however quite of a few issues that need to be addressed
towards their application for treatment of bacterial infections. These include substrate and/
or site specificity, systemic toxicity and loss of enzymatic activity in vivo due to low
stability. In order to overcome these problems, numerous efforts have been devoted to the
development
of
several
immobilization
procedures.
Presently,
immobilized
proteins/enzymes are used routinely in the medical field, such as in the diagnosis and
treatment of various diseases. For example, immobilized antibodies, receptors, or enzymes
are used in biosensors and ELISA for the detection of various bioactive substances in the
diagnosis of disease states; antibody directed drug delivery using colloidal systems [135137].
Immobilization can be defined as the attachment of molecules to a surface
resulting in reduction or loss of mobility. In fact sometimes, immobilization may lead to
24
partial or complete loss of protein activity, due to random orientation and structural
deformation [138]. There is a multipoint attachment between enzyme molecules and the
surface that reduces unfolding, and hence improves stability [139]. In order to retain
biological activity, enzymes should preferably be attached onto surfaces without
significantly affecting their conformation and function.
Generally, the choice of a suitable immobilization strategy is determined by the
physicochemical and chemical properties of both surface and enzyme. The stability of an
immobilized enzyme is dictated by many factors such as the number of bonds formed
between the enzyme and carrier, the nature of the bonds (i.e. covalent, non-covalent or
different types of covalent or non-covalent bonds), the degree of confinement of enzyme
molecules in/on the carrier, the microenvironment of the enzyme and carrier, and the
immobilization conditions [138]. For e.g. nonporous materials, to which enzymes are
attached to the surfaces, are subject to minimum diffusion limitation while enzyme
loading per unit mass of support is usually low. On the other hand, porous materials can
afford high enzyme loading, but suffer a much greater diffusional limitation of substrate.
Many surface immobilization techniques have been developed in the past years,
which are mainly based on the following three mechanisms: physical, covalent, and
bioaffinity immobilization [138]. It is unlikely that one immobilization technique will be
considered optimal for all proteins. Also, the most suitable strategy for protein
immobilization is also particularly dependent upon the biomedical application. Hence,
several unsolved challenges are involved in immobilization techniques. Correspondingly,
many methods have been developed to improve the enzyme stability during
immobilization. Engineering the microenvironment of the enzyme molecules has been
25
increasingly used to improve enzyme stability. For e.g. in the case of covalent enzyme
immobilization, alteration of microenvironment can be easily achieved by quenching the
remaining excess active functionalities using blocking agents [140], which can be simply
classified into two groups: small molecules (e.g. amino acids or other amines), and
macromolecules (e.g. bovine serum albumin, gelatin, polyethyleneimine (PEI) [141] , and
polyethylene glycol (PEG)). These blocking agents (or quenching agents) can easily react
with the active functionalities, such as epoxy rings or aldehyde groups, of the commonly
used carriers without the use of additional activating chemistry.
Other methods of immobilization include entrapment via inclusion of an enzyme in
a polymer network (gel lattice) such as an organic polymer or a silica sol-gel, or a
membrane device such as a hollow fiber or a microcapsule. Immobilization in silica sol
gels prepared by hydrolytic polymerization of tetraethoxysilane, in the presence of the
enzyme [142] and has been used for the immobilization of a wide variety of enzymes
[143]. Also, it has been shown that cross-linking of dissolved enzymes via reaction of
surface NH2 groups with a bifunctional chemical cross-linker, such as glutaraldehyde,
results in insoluble cross-linked enzymes with retention of catalytic activity [144].
However, this method of producing cross-linked enzymes had several drawbacks, such as
low activity retention, poor reproducibility, low mechanical stability, and difficulties in
handling the immobilized enzyme. Some methods of immobilization are described below.
2.4.1. Physical adsorption
The type of intermolecular forces that take part in the adsorption process will
depend on the particular protein and surface involved. Protein adsorption on surfaces is
predominantly governed by electrostatic, hydrophobic interactions and to certain extent
26
even hydrogen bonding. The resulting layer is likely to be heterogeneous and randomly
oriented, since each protein molecule can form many contacts in different orientations for
minimizing repulsive interactions with the substrate and previously adsorbed proteins
[145].
The arrival of protein at the interface is assumed to be driven solely by diffusion
processes, which are dependent on bulk concentration and diffusion coefficient [145]. The
particular surface chemistry, intermolecular forces between adsorbed molecules, solvent–
solvent interactions, topology and morphology of the protein-adsorbent interface also
further dictates the adsorption kinetics. Protein adsorption process includes the
redistribution of charged groups in the interfacial layer, hydrophobic interactions between
protein and material‟s surface, and structural rearrangements of protein molecules [145].
Drawbacks of the adsorption are random orientation resulting in significant loss of
activity and weak attachment followed by subsequent protein desorption from the surface.
The structure of a protein is closely related to its function therefore unspecific protein
adsorption and surface-induced conformational changes are important issues in
biocompatibility of materials [146]
2.4.2. Covalent Immobilization
This method confers the immobilized enzyme with greater stability against
environmental changes such as pH and temperature, as well as higher selectivity toward
the substrates. However, the coupling protocols could involve complicated synthetic
procedures, which are often carried out under harsh experimental conditions, and with use
of toxic and expensive reagents. Furthermore, functional groups of the enzyme are used to
27
link the solid supports, resulting in a significant loss in enzymatic activity [147]. Covalent
bonds are mostly formed between side-chain-exposed functional groups of proteins with
suitably modified supports, resulting in an irreversible binding and producing a high
surface coverage [148]. The functional groups on the support are generated by chemical
treatment, and several pretreated surfaces are already commercially available.
Covalent attachment is permanent, leaving no unbound material after clean-up. It
may prevent elution of bound protein during storage, thus increasing shelf life. Covalent
coupling on surfaces enables a uniform coating procedure [149, 150]. As a consequence of
having reactive groups over the entire surface of the material, it is possible to completely
cover the surface with protein. Protein coverage is also more easily controlled by covalent
coupling, especially when the desired quantity of adsorbed protein is low. Achieving the
correct spatial orientation for the bound protein can be difficult via physical adsorption.
Covalent attachment, on the other hand, can orient the molecule properly, if the correct
coupling chemistry is chosen, improving the activity of the bound proteins and resulting in
lower reagent consumption [149, 150].
Because active enzymes necessarily have very flexible conformation, a spacer
between the enzyme and the carrier surface is often used to improve the enzyme activity
expression[151]. The obtained immobilized enzymes with spacers could have higher
activity due to the favorable environment created by the hydrophilic spacer. Most
probably, the presence of the spacer is able to endow the enzyme more conformational
flexibility, which is usually a prerequisite for higher activity. Controlled modification of
the enzyme molecules with the use of a bi-functional cross-linker have also been recently
used to adjust the number of bonds and the spacer formed between the carrier and enzyme
28
.To preserve the activity of biomolecules upon immobilization, several techniques have
been directed towards site-specific covalent immobilization [152, 153]. For antibodies, the
carbohydrate moieties located in the Fc region were targeted to form aldehyde groups,
using periodate for site-specific coupling to solid supports [154, 155].
2.4.3. Bioaffinity Immobilization
Biochemical affinity reactions are also used for immobilization of proteins,
providing an important advantage over other surface immobilization techniques.
Moreover, not only oriented and homogeneous attachment is obtained, but it is also
possible to detach proteins and make repeated use of the same surface. An example is
described below.
Avidin-Biotin interaction
This approach takes advantage of one of the strongest noncovalent bonds (Kd =
1015 M-1) and allowing the use of harsh conditions during biochemical assays [156]. The
specificity of the interaction permits uniformly oriented protein immobilization [157].
Avidin is a tetrameric glycoprotein soluble in aqueous solutions and stable over wide pH
and temperature ranges. It can bind up to four molecules of biotin. This bond formation is
very rapid and unaffected by pH, temperature, organic solvents, enzymatic proteolysis,
and other denaturing agents [157].
Streptavidin is a closely related tetrameric protein, with similar affinity to biotin,
but differing in other aspects, such as molecular weight, amino acid composition, and pI.
The properties of both avidin and streptavidin have been improved using chemical and
recombinant methods providing enhanced stability and/or controlled biotin binding [157].
Biotin or vitamin H is a naturally occurring vitamin found in all living cells. Only the
29
bicyclic ring is required to be intact for the interaction with avidin; the carboxyl group on
the valeric acid side chain is not involved and can be modified to generate biotinylation
reagents used for conjugation with proteins [158]. Since biotin is a small molecule, its
conjugation to macromolecules does not affect conformation, size, or functionality [158].
Biotinylation reagents can be classified depending on their reactivity toward different
functional groups. The NHS ester of biotin is the most commonly used biotinylation
reagent to target amine groups [159].
Both biotin and avidin/streptavidin may be attached to a variety of substrates.
Direct immobilization of avidin occurs either via adsorption or via covalent coupling. It
has been reported that covalent attachment of avidin onto carboxyl-derivatized polymers
produces a higher coating density than immobilization via a biotinylated interlayer [160].
However, coupling via carbodiimide chemistry may affect its binding activity. A typical
multilayer is composed by biotin directly immobilized and avidin creating a secondary
layer for binding biotinylated molecules. This approach is generally preferred due to the
higher organization (biotin/avidin/biotin) obtained in comparison to that of the direct
immobilization of avidin [161].
30
Figure 2.4. Schematic of the Streptavidin-biotin system: Binding of biomolecules
to the particle surface: (Source: http://www.microparticles.de/news_2005.htmlaccessed - 20/03/2011).
2.4.4. Surface modification
Spontaneous adsorption of proteins from biological fluids onto synthetic materials
such as biomaterials and biomedical devices may induce undesirable reactions of the body
to the foreign materials, such as immune responses, blood coagulation, or bacterial
adhesion [162, 163]. Due to the diversity of the interactions between proteins and
surfaces, a preferred strategy for blocking the adsorption of proteins is to immobilize
polymers in the form of well-solvated brushes (e.g., poly (ethylene glycol), PEG) on the
surface.
Pegylation of a surface can be achieved by adsorbing PEG-containing block
copolymers such as polyethylene oxide/polypropylene oxide [164] or poly(d,l-lactide)/
polyethylene oxide [160] on the surface, surface grafting using established strategies
[165]depositing PEG-like coatings using a radiofrequency glow discharge technique [166]
or forming self-assembled monolayers. The polymer layer shields the surface, introducing
a high activation barrier for the proteins to adsorb. Poly (ethylene glycol) (PEG) has been
shown to successfully confer protein resistance to a variety of surfaces [167, 168]. Both
PEG chain length and surface coverage have been demonstrated to play a key role in
imparting protein resistance, with PEG chains of typically 1-10 kDa molecular weight
providing protein resistance. The protein resistance by PEG chains has been associated
with two main mechanisms, steric repulsion and a hydration or water structuring layer
[167].
31
Attachment via PEG spacers also minimizes lateral interactions of immobilized
protein with the surface [169, 170]. The spacer groups are thought to permit a degree of
freedom to the reagent moiety separating it from the particle surface, thereby lending
enhanced specificity [167]. In particular, nanoparticles whose surfaces were modified by
the incorporation of poly(ethylene glycol) (PEG) during nanoparticle formulation either
through covalent attachment of PEG to surface functional groups or through physical
adsorption of PEG to the surface, have show a decreased uptake by phagocytic cells and
an increased circulation time to effectively target diseased cells [171, 172]. PEG
molecules on the surface of a polymeric nanoparticle can reduce the adsorption of
opsonins and other serum proteins [172]. PEGylated polycyanoacrylate particles
transferred more effectively across the blood brain barrier than with poloxamine coating
[173].PEG density and configuration were shown to be important for this [174].
Additives such as polyethylene glycol (PEG), polyvinyl alcohol and albumin, can
have a stabilizing effect even on sol-gel entrapped enzymes. For example, Zanin and coworkers [175] compared three different methods – physical binding, covalent attachment
and gel entrapment, in the presence and absence of PEG 1450 – for the immobilization of
Candida rugosa lipase. Immobilization yields varied from 3 to 32%, the most active
biocatalyst resulting from the encapsulation in the presence of PEG.
2.5. Nanotechnology for enzyme immoblization
Nanotechnology also seems to be as a promising alternative to overcome the
problems of the administration of peptides and proteins and also stabilizing these
biomolecules before targeted delivery. Various nanostructures, generally providing a large
32
surface area for the immobilization of enzyme molecules, have been actively developed
for enzyme stabilization. The coupling of biomolecules and materials at the nanoscale has
the potential to revolutionize many fields of science and technology potentially having a
very significant impact on biomedical technology. Both nanomaterials and biomolecules
lie in the same nanometer length scale and can thus complement each other in hybrid
systems. Protein based nanostructures are expected to offer some additional advantages
and play a key role in the development of multifunctional materials and devices for
nanotechnological applications. [176-179].
Many methods have been developed to incorporate enzymes into nanostructures.
Surface attachment was employed in many of the earlier studies and is often the method of
choice [180-182]. Another way to develop nanoscale biocatalysts is to entrap enzymes
within nanopores. Mesoporous silica gels have been used to host enzymes through
physical adsorption [183] or chemical binding. Using carefully designed synthetic routes,
enzymes were also entrapped within the cores of discrete polymeric articles. More
sophisticated structures, such as porous materials hosting enzyme-carrying nanoparticles
and cross-linked enzyme aggregates [185], have been developed by applying enzyme
modification and fabrication procedures. Thus the functionalization and modification of
nanomaterials using enzymes have led towards research and development for biomedical
applications.
However there are drawbacks to the handling of nanomaterials, which presents
certain health and environmental concerns. Carbon nanotubes, which exist in powder form
when in a pure state, share these concerns when used as enzyme supports due to toxicity
[180-182]. Some of these problems can be overcome by using one-dimensional
33
nanomaterials, such as polymeric nanofibers [186]. The surface area: volume ratio of
nanofibers is also high, representing two-thirds that of nanoparticles of the same diameter
when considering equivalent amounts of material. Nanofibers, however, have the benefit
of being much easier to produce and handle. They can be applied in the form of coils,
sheets or dispersed fibers and can also be attached to the surface of other materials or
blended with them, thus offering very flexible design of reactors [186].
In the case of surface attachment, smaller particles can provide a larger surface
area for the attachment of enzymes, leading to higher enzyme loading per unit mass of
particles [187]. A growing interest has been shown in using nanoparticles as carriers for
enzyme immobilization [187-190]. The effective enzyme loading on nanoparticles could
be achieved up to 10% wt due to a large surface area per unit mass of nanoparticles [188].
Figure 2.5. Nanotechnology for enzyme immobilization (a) Nanoparticles with
surface-attached enzymes. (b) Nanofibers carrying enzymes. (c) Nanoporous
matrix with entrapped enzymes. (d) Carbon nanotube–enzyme hybrid materials –
Figure taken from Wang et al; 2006 [191].
34
2.6. Nanoparticles
The use of nanoparticles as enzyme supports was first reported in the late 1980s
[192, 193]. Therefore, nanoparticles may appear as alternative carriers considering the
design of more sophisticated systems including multifunctional types of device.
Nanoparticles are of great scientific interest as they are effectively a bridge between bulk
materials and atomic or molecular structures. Size-dependent properties are observed in
case of nanoparticles when compared to the bulk material. Since they have a very high
surface area to volume ratio it provides a tremendous driving force for diffusion,
especially at elevated temperatures.
Advantages of using nanoparticles over other „nanomaterials‟ and formulations are
that they (e.g. polymer nanopaticles) are cheaper materials than lipids (e.g liposomes), and
that polymers offer wider chemical engineering solutions. In addition, the stability of
polymer particles can be much better controlled than the stability of liposomes upon slight
modifications of the formulation. Intrinsic properties of nanoparticles, such as size,
surface charge density, surface chemistry etc can also be used to add functionality to their
conjugates with biomolecules.
For example, drugs, proteins and peptides can be incorporated into nanospheres
that are composed of a biodegradable polymer, and this allows for the timed release of the
drug as the nanospheres degrade [194] The circumstances that cause the particle to
degrade can be adjusted by varying the nature of the chemical bonding within the particle.
For example, when acid-labile bonds are used, the particles degrade in acidic
microenvironments, such as would exist in tumour cells or around a site of inflammation
[195], and so this approach allows site-specific delivery. In other recent studies, polymeric
35
nanoparticles were labeled on their outer surfaces with a viral peptide sequence that
promotes the permeation of substances through cell membranes [196]. These peptidederivatized nanoparticles passed through cell membranes, and were incorporated into
living cells at much higher levels than nanoparticles without the surface-bound peptide.
Similarly Gold nanoparticles (20 nm) modified with shells of bovine serum
albumin (BSA) were conjugated to various cellular targeting peptides to provide
functional nanoparticles that penetrate the biological membrane and target the nuclei [197,
198]. Hybrids of silver nanoparticles with amphiphilic macromolecules exhibit
antimicrobial properties and are used to treat bacterial infections [198]. Covalent coupling
of proteins to metal nanoparticles is performed using appropriate capping agents such as
citrate and thiol groups. This method has been applied in coating of colloidal gold with
thiol containing proteins such as immunoglobulins and serine albumins which have
cysteine residues that are accessible for coupling [199].
2.6.1. Polymer nanoparticles
The term `polymer colloid' refers to a suspension or dispersion of polymeric
microspheres having a diameter in the order of sub-micron to several microns. The
dispersion medium is generally water. A dispersion whose medium is not water is referred
to as a nonaqueous dispersion. `Latex' originally means a dispersion of microspheres from
natural rubber. The stability of the polymer latex itself is decided by the contribution of
three factors: electrostatic repulsive forces, van der Waals' attractive forces and steric
repulsive forces among the particles [200].
Polymer colloids have a low viscosity and high fluidity compared to solutions
containing the same amount of solid. The viscosity of polymer colloids is a universal
36
function of apparent volume fraction of the microspheres. The apparent volume fraction of
microspheres can be changed by environmental conditions such as pH and temperature for
some polymer colloids [201]. In dispersion, the microspheres can move macroscopically
through the medium by gravity, electric field, etc and microscopically via Brownian
motion. These movements keep a fresh interface between the microspheres and the
medium. Particles having soft layers on the surface allow water to penetrate the layer and
meet less resistance when they move through water [202]. Polymer particle dispersions are
an alternative to be able to offer a large surface and they can be used as the carrier if easily
recovered from the medium.
Polymer latexes, usually with antibodies or antigens attached to their surfaces, can
be used in the diagnostic tests. There are many factors that dictate the use of latex particles
than the use of any other solid support for agglutination reactions. These include
monodispersity (for detection of agglutination using light scattering techniques), colloidal
stability and hydrophobicity of the particles. Biological molecules adsorb strongly to
hydrophobic surfaces and this can be controlled by time, temperature, pH and protein
concentration [149]. An antibody is an asymmetric molecule and its antigen-binding
fragment (Fab) is located in two arms of Y-shaped molecules. Antibodies must be
immobilized in a manner in which the Fab fragment does not have any steric hindrance and
can bind to the directed antigen. When an antibody is immobilized in the opposite
orientation, it causes not only lowering of reactivity but also nonspecific agglutination,
since the fragment Fc of an antibody adsorbs to certain proteins [149]. Furthermore,
surface activity could be enhanced if the antibodies are covalently coupled, by reducing
the rearrangement of the protein molecules during and after adsorption [203-205]. Specific
37
covalency between protein and functionalized latex is achieved by treatment of protein–
latex system with surfactants under appropriate conditions able to remove all the
physically adsorbed protein from the particle surface [150].
2.6.2. Protein-nanoparticle conjugates
In recent years, much attention has been paid to the use of enzymes conjugated to
polymer colloids in biomedical applications [206-208]. Such colloids are easy to produce
and can be prepared with a high degree of monodispersity [209-211]. Colloidal
nanoparticles provide an almost ideal remedy to the issues encountered in the optimization
of immobilized enzymes: minimum diffusional limitation, maximum surface area per unit
mass, and high enzyme loading [187]. The attachment of molecules to latex particles can
be achieved through physical adsorption or covalent coupling.
Polymer engineering has facilitated the synthesis of latex particles with surface
reactive groups such as carboxylate, aldehyde, chloromethyl, and hydroxyl groups to
enable covalent coupling of protein molecules to the particles[166, 212, 213] Some of
these groups must be activated prior to protein immobilization. A number of special
linkers can be used to convert one surface functional group on a microsphere to another.
For example, amino microspheres can be converted to carboxylic particles by reacting
with succinic anhydride [214]. Carboxylic particles can be converted to amino
microspheres through water-soluble carbodiimide-mediated attachment of a diamine
[215].
38
Fig 2.6. Commonly used covalent coupling procedures for peptides and proteins
to the surface of nanoparticles with reactive surface groups.
There are also major drawbacks of any immobilization protocol based on the use
of highly reactive and unstable intermediates. The balance between covalent coupling and
unproductive side reactions depends to some extent on the activation reaction conditions.
The ionic composition of the reaction, the pH, and the buffer type can greatly enhance or
inhibit the total binding of protein to particles. Sufficient amounts of functional groups
should be present to provide adequate coupling of the protein. The covalent attachment of
the IgG molecule to carboxylated particles improves the immunoreactivity of antibodies
when compared with physical adsorption and also maintaining its immunoreactivity after
long periods of storage [150].
2.6.3. Stabilization using blocking agents and surfactants
There also other techniques in order to improve stability and reduce non specificity
of the protein-nanoparticle conjugates. The colloidal suspension can be treated with either
a second protein or a surfactant that can act as a stabilizer by covering all the unoccupied
39
parts of the surface and thereby reducing non specificity [216]. Bovine serum albumin
(BSA), a globular protein has been commonly used to improve the colloidal stability of
protein coated microspheres. The BSA molecule is a highly-charged protein at
physiological pH. It helps in the electrostatic stabilization of the protein-nanoparticle
conjugate [217, 218]. Moreover, BSA is easily obtainable in significant amounts, which is
why it is commercially available at reasonably low prices.
There are two different methods for preparing latex–antibody complexes with
coadsorbed BSA: (1) sequential adsorption where in a first step the antibody adsorption is
carried out and after centrifugation the conjugate is resuspended in a solution of BSA at
constant concentration [219]; (2) competitive adsorption whereby the adsorption of both
proteins (including BSA) occurs in a single step [217, 218]. For competitive adsorption,
the method becomes more complicated because of the two types of proteins in the
dispersion medium. Peula et al. observed that the colloidal stabilization of IgG-covered
particles appears when the coverage of co-adsorbed BSA is high and at pH 7 and 9, and is
not stable at pH 5 (pI of albumin) [217]. It was also indicated from the same study that it
is necessary to find the adequate equilibrium between the amount of IgG that produces a
good immunological response and the amount of BSA responsible for the colloidal
stability. These ways of stabilizing antibody–latex particles may have some disadvantages,
e.g., only antibody–latex complexes with low antibody coverage can be stabilized or coadsorption with inactive proteins may involve partial or complete displacement of the
preadsorbed molecules of antibody.
The use of surfactants (e.g. Tween) as stabilizer molecules can also have also have
desired effects on nonspecific agglutination. They mainly stabilize protein-coated particles
40
by means of steric forces, although electrostatic repulsions can also be generated by ionic
surfactants. The surfactant concentration requires careful optimization because excess
surfactant could also inhibit interaction with the desired antigen/substrate. However, this
strategy to preserve the colloidal stability of particles presents some disadvantages, since
such molecules can also desorb the previously adsorbed protein [220].
2.7. Enzyme-nanoparticle conjugates as antibacterials – Targeted delivery
The stratum corneum layer of the epidermis is a strict barrier allowing limited
penetration of particulate materials. This aspect has potential to serve as a novel route for
drug delivery and has attracted enormous pharmaceutical research interests [221]. While
intact skin is known to act as a barrier to nanoparticles, skin wounds may give rise to
easier translocations. Dressings and bandages embedded or coated with antibacterial
protein-nanoparticle conjugates could be potentially used for prevention of sepsis of
severe skin wounds like burns. Close contact may allow nanoparticle-conjugates to
penetrate through compromised skin barrier and gain access to the dermal capillaries.
Recently Solid lipid nanoparticles (SLN) are gaining increasing importance as
pharmaceutical formulations [222, 223]. Due to their small particle size, chemistry and
consequent high surface area, they possess strong adhesive properties. Incorporation of
antibacterials into the solid lipid matrix offers protection against chemical degradation of
the active compound, as well as allowing either immediate or sustained release, depending
upon the application [224]. Sustained release is important with antibacterials that are
irritating at high concentrations or when a supply to the skin over a prolonged period of
time is desired, whereas immediate release can be useful to improve penetration.
41
Antibacterial
activity
of
nisin-loaded
polylactide
nanoparticles
against
Lactobacillus delbrueckeii has also been evaluated [225]. In most of these cases, the role
of nanoparticles was to provide a sustained release of the loaded drug; however, targeted
delivery of antibiotics to infected macrophages and to liver has been utilized using
nanoparticles and liposomes [226] . The above targeting was made possible because of the
increased uptake of polymeric nanoparticles by reticuloendothelial system.
More recently, specific targeted delivery of polymeric nanoparticles either
unloaded or loaded by an antimicrobial drug, to bacteria had been studied. The
antimicrobial effect of insoluble cross-linked quaternary ammonium polyethylenimine
(PEI) nanoparticles incorporated at 1% w/w in a resin composite was assayed and resulted
in complete inhibition of growth S. aureus, S. epidermidis, P. aeruginosa, E. faecalis, and
E. coli (p < 0.0001) [227]. Mannosylated or galactosylated polystyrene latex nanoparticles
were shown to result in agglutination of mutant strains of E. Coli overexpressing mannose
and galactose surface receptors, respectively [228, 229]. These studies have been thus far
restricted to model E. coli mutants; however, the authors envision development of
nanoparticles coated by oligosaccharides that will specifically recognize desired bacterial
strains and kill them by sticking to bacterial cell walls and causing bacterial agglutination.
Effect of antibody-directed targeting to Gram-positive Listeria Monocytogenes has
recently been studied by the same group for lysozyme attached to polystyrene latex
nanoparticles simultaneously with anti- L. Monocytogenes antibody [230]. The authors
observed significantly (p<0.05) higher antibacterial activity for nanoparticle conjugates
containing both lysozyme and the antibody, than for free lysozyme and enzymenanoparticle conjugates in the absence of the antibody at higher concentrations.
42
2.8. Enzymes as antibacterial coatings on medical devices and implants
Bacterial adherence to an implant is a known precursor to prosthetic infection
[231, 232]. Antimicrobial coating of medical devices/implants has recently emerged as a
potentially effective method for preventing infections [233-236]. Several attempts have
been undertaken to immobilize enzymes and peptides on biomedical devices and implants
to prevent device-related infections. For a catheter surface, the coating may be retained by
either physical adsorption or covalent attachment of the enzyme.
Wilcox et.al immobilized synthetic antimicrobial peptide melimin on contact
lenses by physical adsorption, and observed significantly decreased bacterial colonization
[237]. In another embodiment, antimicrobial protein lactoferrin was also adsorbed on
contact lenses by the same group of authors and was also found to prevent bacterial
colonization [238]. Balaban et.al found that adsorbed dermaseptin to Dacron vascular
grafts showed resistant to bacterial infection and demonstrated in vivo efficacy in a rat
model [239]. Langer et al covalently attached a hybrid synthetic AMP, Cecropin-Melittin,
to model surfaces in a highly ordered manner, and reported enhanced antibacterial activity
in comparison to physically adsorbed and randomly immobilized peptides. Shah et.al
performed demonstrated significant antimicrobial activity against S. aureus for
intravenous catheters coated by physically adsorbed lysostaphin, and observed high
efficacy [240]. DispersinB, a naturally occurring enzyme in bacteria, Actinobacillus
actinomycetemcomitans, coated on polymer well plates, rods and catheters has been
shown to prevent the formation of Staphylococcus epidermis biofilms [241].
Despite such growing interest, a combination of enzymes coated on implant
surfaces to prevent bacterial colonization and subsequent biofilm formation has not been
43
studied. DispersinB, is an enzyme that is specifically active against Staph species,
however when used alone it can only inhibit and disperse biofilms, but not in killing of the
bacteria, which could still persist in the blood-stream to cause infection. So, it would be
necessary to combine enzymes such as DispersinB (antibiofilm) with antimicrobial
enzymes and peptides such as lysozyme, lysostaphin, LL-37 etc. to not only prevent
colonization but also lyse bacteria embedded in biofilms. The application of enzymes and
peptides has been shown to work synergistically against a broad spectrum of bacteria from
various studies [102-104]. But a systematic study on combination of different enzymes
and/or peptides coated on surfaces at different coating concentrations working
synergistically has not been performed. Also, aspects of enzyme leaching, cytotoxicity and
long term durability on devices for antimicrobial applications have not been studied in
detail. With this in mind, the final goal of our project aimed to address through
nanotechnology, all issues involved in coating of enzymes (alone or in combination) on
catheters by evaluating their in vitro performance through simple model studies.
44
CHAPTER 3
SPECIFIC AIMS AND SIGNIFICANCE
3.1. Clinical significance
Central venous catheters are commonly used modality in hospitals and clinics for
administration of fluids, chemotherapeutic agents, central venous pressure monitoring,
extracorporeal treatment regimen etc. thus serving a vital role as both a therapeutic and
diagnostic tool for monitoring of health in critically ill patients. Nearly 5 million CVCs are
inserted annually [242]. However, central venous catheterizations are associated with
nearly 90% of nosocomial infections and are responsible for nearly 400,000 cases per year
in the United States [242]. Although the incidence of local or bloodstream infections
(BSIs) associated with peripheral venous catheters is usually low, serious infectious
complications produce considerable annual morbidity because of the frequency with
which such catheters are used. The magnitude of the potential for CVCs to cause
morbidity and mortality resulting from infectious complications has been estimated in
several studies [243]. The attributable cost per infection is estimated to be ~ $45000 [244]
and the annual cost of patient treatment with CVC-associated BSIs has gone up to $2.3
billion [245].
Finding an effective treatment is also difficult as S. aureus has the ability to form
biofilms on devices, which makes it even more impenetrable for antimicrobial agents. The
antibiotic concentrations required to kill bacteria in the biofilm is a thousandfold higher
than that needed to kill the planktonic cells of the same species [246]. Recent prophylactic
strategies have included precoating devices with different antibiotics [247], however the
45
occurrence of a number of multi-drug resistant strains as spurred the need for alternatives
to antibiotic treatment. [248].
Use of enzymes and peptides as antimicrobial coatings on surfaces has prevented
device-associated infections [79-81]. The use of nanoparticles (NPs) as surfaces for
enzyme immobilization offers several important advantages over the use of purely enzyme
based coatings.
Inherent physiochemical properties of NPs. can be used to improve
activity of the enzymes immobilized on them [249]. Although coating medical devices and
implants with plain antimicrobial enzymes could work just as effectively, coating catheter
surfaces with enzyme-NP conjugates may be advantageous due to larger surface area for
effective enzyme coating, higher stability, durability and lower enzyme leaching. However
it is important to characterize enzyme-NP conjugates by studying individually whether
aspects of nanoparticle size, surface charge density etc. could help improve efficacy of
enzymes for antibacterial applications.
Our studies involve simple models for studying efficacy of enzyme-NP conjugates
to planktonic bacteria or cell suspensions, simultaneously against that of free enzymes.
Also, these studies will help optimize and standardize a protocol that can be used for the
preparation and synthesis of enzyme-NP conjugates for multiple enzymes for different
applications. For e.g. by studying the binding yields of enzymes at different NP: enzyme
ratios, we can understand the binding affinities for each of the enzymes to the surface of
nanoparticles. Specific antibacterial activity assays can quickly be used to measure the
activity of conjugated enzymes to NPs. Here it is important to clarify the difference in
using the terms „activity‟ and efficacy of enzyme-NP conjugates. Activity of the enzyme
refers to a specific assay that characterizes the catalytic function of the enzyme against its
46
substrate. Efficacy of enzyme-NP conjugates refers specifically to the targeting and
antibacterial activity of the conjugates against bacterial cell suspensions.
High antibacterial efficacy of the conjugates against planktonic cells does not
imply high efficacy against the same bacteria in colonies, as different factors may control
activities in these cases. Therefore, it is necessary to perform in vitro experiments in order
to evaluate the inhibition and disruption of bacteria grown on a surface.
Various
combinations of enzymes have been studied for the disruption and destruction of bacterial
biofilms [106]. This requires a minimum of two enzymes, one enzyme for disruption of
the biofilm matrix and a second enzyme with bactericidal activity. Conjugation of these
enzymes to the surface of the same catheter could address the problems associated with
this application. We chose to attach enzymes to the surfaces of PLA nanoparticles,
characterize enzyme-NP conjugates and coat catheter segments at different concentrations,
simultaneously coating catheters with free enzymes and then evaluating the inhibition of
S. aureus biofilms. Thus adsorption of two enzymes to the same surface could result in a
synergistic effect that could cause inhibition and lysis of the biofilm-infected device and
subsequently avoid the need for surgical removal of the infected device.
Since our system will function as a blood contacting device in vivo, the sample
coated catheters will encounter various blood plasma proteins [235]. If binding yield and
activity of enzyme-NP conjugates coated on catheters is significantly lower than free
enzyme coatings or enzyme leaching from the surface of the NPs is a problem, then an
alternative strategy will be precoating the catheter segments with NPs before enzyme
adsorption . Then, the goal of our final study will be comparison of in vitro performance
for plain and NP coated catheter segments coated by antimicrobial enzymes.
47
3.2. Specific aims
Specific aim I. Effect of charge-directed targeting on antibacterial activity using
enzyme-nanoparticle conjugates of lysozyme to Micrococcus lysodeikticus
Hypothesis: Use of antimicrobial enzymes attached to nanoparticles is of special
interest alternative to antibiotics to treat microbial infections. Special properties of the
carrier can be used to improve targeting and activity of the conjugated enzyme. Here, we
decided to show in a model system that nanoparticle charge can be used to enhance
delivery of lysozyme to the cells of its natural substrate, Micrococcus lysodeikticus and
significantly improve its lytic activity. Hen egg lysozyme was covalently attached to two
types of polystyrene latex nanoparticles: positively charged, containing aliphatic amine
surface groups, and negatively charged, containing sulfate and chloromethyl surface
groups. Covalent conjugation of lysozyme to each of these groups was achieved either by
direct conjugation or by using PEG spacers. Our study aimed at comparing the lytic
activity of these enzyme-nanoparticle conjugates with that of free enzyme using either
bacterial or low molecular weight substrates. The effect of attaching PEG spacers and
method of enzyme purification from nanoparticles (through dialysis and centrifugation) on
enzyme activity was also studied [250].
Specific aim II. Effect of antibody-directed targeting on antibacterial activity using
enzyme-nanoparticle conjugates of lysostaphin to Staphylococcus aureus
Hypothesis: Our previous study clearly illustrated that physiochemical properties
of nanoparticles can be used to help improve targeting of enzymes to bacterial cells.
Lysozyme shows little or no antibacterial activity by itself against S .aureus [251].
48
However, lysostaphin is an antibacterial enzyme which specifically cleaves cross-linked
pentaglycine bridges in the peptidoglycan of S. aureus, thereby hydrolyzing the cell wall
and lysing the bacteria [252]. We decided to study and compare the antimicrobial activity
of lysostaphin adsorbed on the surface of PLA (poly (lactic acid)) nanoparticles and
compare the activity of immobilized lysostaphin to that of the free enzyme. Enhanced
antimicrobial efficacy against Gram-positive Listeria monocytogenes has been previously
demonstrated for lysozyme co-immobilized with anti- L. monocytogenes antibody on
polystyrene latex NPs for applications in storage and food packaging [230]. However, a
systematic study of activity of targeted enzyme-NP conjugates with different enzyme and
antibody loadings against clinically relevant pathogens has not been performed previously.
With this in mind, the goal of the present work was a detailed study of antimicrobial
activity of targeted (lysostaphin-antibody-NPs) and untargeted (lysostaphin-NPs)
conjugates against S. aureus at different enzyme and antibody ratios.
Specific aim III. Evaluation of antimicrobial activity of lysostaphin coated hernia repair
meshes
Hypothesis: The goal of the earlier proposed work was to initially show a proof of
concept where we could achieve enhanced antibacterial efficacy through targeted delivery
of enzymes using polymer nanoparticles to cell suspensions (model for planktonic or free
floating bacteria) in vitro (Specific aim I and II). Bacteria can either colonize the implant
surface or exist in a planktonic free floating form. Hence we had initially proposed to use
enzymes and enzyme-NP conjugates as surface coatings for indwelling medical devices
and implants to inhibit bacterial adhesion and subsequent colonization. But as mentioned
earlier, high activity of enzymes of conjugates enzymes to bacterial cell suspensions does
49
not imply high activity against colonized bacteria, as different factors may control
activities in these cases.
Therefore, it was necessary to perform in vitro experiments in order to evaluate the
inhibition bacteria adhering to a surface and then expand these studies to test and compare
the antimicrobial activity and antibiofilm activity of devices coated by enzymes and
enzyme-NP conjugates. However, the studies required to characterize and evaluate
enzymes and enzyme-NP conjugates coated on implant surfaces are slightly different
compared to studies characterizing enzymes conjugated to nanoparticles. Therefore,
before embarking on this study, we decided to study and characterize antimicrobial
activity of enzyme adsorption to implants and also evaluate inhibition of bacterial
colonization.
The objective of this study was to evaluate the in vitro antimicrobial antimicrobial
susceptibility of S. aureus to lysostaphin-coated hernia repair polymer meshes and study
the effect of enzyme coating concentration on antimicrobial activity. A pilot in vivo study
was also performed by our collaborators at Carolinas Medical Center to study dosedependence on antimicrobial activity of lysostaphin-coated meshes using in a rat hernia
repair model.
Specific aim IV. Application of enzymes as antibacterial coatings on plain and
nanoparticle coated catheter segments
Hypothesis: We had initially proposed to study and characterize the in vitro
antimicrobial and antibiofilm efficacy of Central venous catheters segments coated with
enzyme-NP conjugates and free enzymes. Different coating concentrations were estimated
in order to optimize the bound weight of enzyme-NP per specific surface area to the bound
50
concentration of plain enzyme for the catheter segment from the same batch. Synthesis of
enzyme-NP conjugates were done via coadsorption of two model enzymes, DispersinB
(antibiofilm) and lysostaphin (antimicrobial) on the surface of Poly ((lactic
acid))nanoparticles. However, we were not able to achieve higher loading via adsorption
of enzyme-NP conjugates on the surface of 1 cm catheter segments compared to plain
enzyme adsorption. We hypothesized that coating by enzymes interfered with nanoparticle
attachment and lead to reduced adsorption.
We therefore proposed an alternative strategy, which involved: a) Initial coating of
the catheter segments by nanoparticles and b) Coating of a series of different initial
enzyme concentrations (same set) on the surface of both plain and NP coated catheter
segments. Through this method, we also did not had to optimize enzyme adsorption
protocols for NPs separately, achieve high monolayer coatings individually and overall
economically it was a more feasible option.
We performed a series of experiments to study and characterize enzyme adsorption
to plain and Poly ((lactic acid))nanoparticles coated Polyurethane catheter segments. PLA
nanoparticle coating on Polyurethane catheter segments were achieved via adsorption and
surface area for plain and NP coated catheter segments was characterized using BET
surface are analysis and SEM imaging. The enzymes used for adsorption were the same
set of enzymes used for characterization earlier, lysostaphin (antimicrobial and
antibiofilm) and DispersinB (antibiofilm). Binding yield of lysostaphin and DispersinB to
plain and NP coated segments either alone or coadsorption (in the presence of the other)
was studied for a series of different coating concentrations through BCA and fluorescence
analysis. Adsorbed lysostaphin and DispersinB activity on catheter segments was done
51
through turbidimetric assay (antibacterial activity) and colorimetric assay respectively.
Enzyme leaching studies were performed using 2% BSA in phosphate buffered saline
(PBS) as a model system. Cytotoxicity of the catheter segments were evaluated using
MTS assay with 3T3 Mouse fibroblasts as a cell model for wound healing. Antibiofilm
efficacy of the coated catheter segments was evaluated using a methylene blue staining
method to quantify the amount of biomass (biofilm) adhered to the surface of the catheter.
Finally we studied long term durability testing of the coated catheter segments to evaluate
its shelf life stability (equivalent of 4 months) for both lysostaphin and DispersinB
adsorbed on plain and NP coated segments. Activity of enzymes was monitored similarly
using a turbidity assay (lysosyaphin) and colorimetric assay (DispersinB) as described
previously.
We expect to prevent bacterial colonization and eventual biofilm formation
through surface coating of the catheters with enzyme-nanoparticle conjugates and plain
enzymes but not expected to treat S. aureus associated-blood stream infections. Long term
durability of NP coated catheters is expected due to enhanced stability of adsorbed
enzyme compared to free enzyme at same coating concentrations. The approach to be
developed here is universal: it can potentially be used for treatment of other medical
device-associated infections.
52
CHAPTER 4
CHARGE-DIRECTED TARGETING OF ANTIMICROBIAL ENZYMENANOPARTICLE CONJUGATES
4.1. Introduction
Antibiotics once were regarded as a universal antimicrobial weapon. However,
many bacteria have developed antibiotic-resistant strains, and the number of such resistant
species is growing quickly [253]. Alternative approaches to treat bacterial infections are
urgently needed in healthcare facilities worldwide. Use of enzymes as antimicrobial
agents is nature-inspired and has recently attracted much attention as an antibiotic-free
approach to treat bacterial infections [254]. The use of antimicrobial enzymes covalently
attached to nanoparticles is of special interest because of enhanced stability of proteinnanoparticle conjugates and the possibility of targeted delivery.
In recent years, much attention has been paid to the use of proteins conjugated to
polymer colloids in biomedical applications [206-208, 255, 256] . Such colloids are easy
to produce and can be prepared with a high degree of monodispersity [210, 211, 257, 258].
Colloidal nanoparticles provide an almost ideal remedy to the usually contradictory issues
encountered in the optimization of immobilized enzymes: minimum diffusional limitation,
maximum surface area per unit mass, and high enzyme loading [187]. Different properties
of nanoparticles, such as size, surface charge density or density of the surface reactive
groups can also be used to add functionality to their conjugates with biomolecules. In spite
of tremendous interest in potential applications of protein-nanoparticle conjugates,
systematic studies of the effect of the properties of nanoparticles on the function of the
attached proteins have rarely been conducted.
53
Here, we study the effect of nanoparticle charge on bactericidal activity of an
antimicrobial enzyme, lysozyme. Hen egg white lysozyme (HEWL) belongs to a
subfamily of c-type lysozymes (chicken or conventional). HEWL has molecular weight of
14.3 kDa and consists of 129 amino acid residues. Among these, 17 are positively charged
(6 lysines containing free amine groups and 11 arginins) and 9 are negatively charged (2
glutamate and 7 asparatate residues). HEWL natural substrate is peptidoglycan murein, the
major component of the bacterial cell wall. Hydrolysis of murein results in bacterial cell
lysis [259]. Gram-positive bacteria are generally more sensitive to lysozyme than Gramnegative bacteria because the latter possess a protective lipopolysaccharide outer
membrane that prevents access of lysozyme to its substrate, murein.
In this study, we covalently attached hen egg lysozyme to two types of polystyrene
latex nanoparticles: positively charged, containing aliphatic amine surface groups, and
negatively charged, containing sulfate and chloromethyl surface groups. Covalent
coupling to each of these groups was achieved either by direct conjugation or by using
polyethyleneglycol (PEG) spacers. Our study aimed at comparing the activity of these
protein-nanoparticle conjugates with that of free enzyme using either bacterial or low
molecular weight substrates.
4.2. Materials and methods
4.2.1. Materials
Hen egg lysozyme and lyophilized Micrococcus lysodeikticus were purchased from
Sigma Aldrich (St. Lous, MO). Bis-butyraldehyde PEG cross-linker and NH2-PEG-COOH
54
cross-linker were purchased from Nektar Therapeutics, Inc. (San Carlos, CA). All other
chemicals were purchased from Sigma Aldrich.
Protein concentrations for binding efficiency studies were determined using BCA
and Micro BCA assay kits purchased from Pierce Biotechnology (Rockford, IL).
Colorimetric and turbidometric activity assays were performed using a Synergy
microplate reader (Bio-Tek Instruments, Inc., Winooski, VT). Spectrum/Por Biotech
Cellulose ester membranes with 5 kDa and 100 kDa (MCWO) from Spectrum laboratories
Inc. (Rancho Dominguez, CA) were used for dialysis.
Protein-nanoparticle conjugates were characterized by zeta potential measurements
using a ZetaPlus Analyzer (Brookhaven Instruments Co, Holtsville, NY). The samples
containing protein-nanoparticle conjugates were diluted 1:100 in 6 mM potassium
phosphate buffer to obtain the appropriate particle concentration. Data presented are the
average of the measurements of three sample replicates each reproduced four times. The
protein-nanoparticle conjugates were characterized using Dimension III Atomic Force
Microscope (AFM) with Nanoscope 4 controller (Vecco Instruments, Santa Barbara, CA).
AFM experiments were conducted in tapping mode in ambient conditions using
rectangular cantilevers backside-coated with aluminum with a spring constant of ~40 N/m
(Mikromasch Inc., Chapel Hill, NC). For sample preparation, a silicon substrate surface
was sonicated in reagent-grade ethanol and then rinsed copiously with HPLC-grade water.
A 10-15 L drop of nanoparticle suspension was placed on the substrate surface,
incubated for 3 min, and blown off with blast of compressed nitrogen.
55
4.2.2. Methods
4.2.2.1. Preparation of protein-nanoparticle conjugates.
The polystyrene nanoparticles used in all experiments were purchased from
IDC.(Interfacial dynamics Corporation, Eugene, OR) and contained monodisperse
particles with a diameter close to 20 nm. Positively-charged nanoparticles surfacemodified with aliphatic amine groups and negatively-charged nanoparticles surfacemodified with chloromethyl and sulfate groups were used in this study.
Sample
Aliphatic
amine
particles
R-CH2Cl
particles
Mean
Solids, Particles
diameter
%
per mL of
of
suspension
particles
(nm)
Surface area
per functional
group
Surface
No. of
density of functional
functional
groups
groups,
per
cm-2
particle
20
2.4
3.1·1015
65 Å2/NH2
15.3·1012
2850
20
4.1
9.3·1015
4828 Å2/RCH2Cl
2194 Å2/ R-SO3-
2.1·1012
4.5·10 12
26
56
The properties of the nanoparticles, as provided by the manufacturer, are highlighted in
Table 4.1. above.
Polystyrene nanoparticles are supplied as a stable suspension in solution
containing a proprietary surfactant. To remove the surfactant, this suspension was purified
by dialysis using a membrane with the 5 kDa molecular weight cutoff. Approximately 1
mL of the suspension was diluted two-fold by 2 mM MES buffer and loaded into the
dialysis tubing. The tubing was then clamped on both ends using dialysis membrane
closures and placed in a 5 L reservoir containing 2 mM MES buffer. The dialysis was
56
performed overnight at room temperature; the entire dialysate was stirred continuously
and changed after 2-4 hours, 6-8 hours and 10-14 hours. Sample recovery was done by
removing it from the tubing with a micropipette. The volume of dialysate did not change
significantly after dialysis. Dialyzed sample was then sonicated for 30 min to destroy
aggregates.
4.2.2.2. Protein conjugation to aminated nanoparticles
In the case of direct conjugation of lysozyme to the amine-modified nanoparticles
(Figure 4.1.A), dialyzed nanoparticles were again diluted two-fold in 2 mM MES buffer
(pH=5.0) to the final concentration of ~7.8·1014 particles per mL. An 8 wt. % solution of
glutaralehyde in 2 mM MES buffer was then added to the diluted nanoparticle suspension
to achieve a final glutaraldehyde concentration of 1 wt. %. A 100 mM solution of sodium
cyanoborohydride, Na[BH3(CN)] (Caution: foul odor; should be used in a chemical hood
only), in 2 mM MES was added to the sample immediately after glutaraldehyde. Final
concentration of sodium cyanoborohydride in the suspension was 10 mM. The amine
groups on the surface of the particles react with the aldehyde moieties to form Schiff‟s
base intermediates, which are then selectively reduced by sodium cyanoborohydride.
After 7 h incubation at room temperature, the unreacted glutaraldehyde and
Na[BH3(CN)] were separated by dialysis using a procedure similar to that described
above. A cellulose ester membrane with a molecular weight cutoff of 5 kDa was used for
dialysis. Six lysozyme solutions with concentrations of 2.94, 29.4, 73.6, 147.2, 220.8 and
294.4 g/mL were prepared in 2 mM MES buffer at pH 5.0 from a 5 mg/mL lysozyme
stock solution. These concentrations were chosen so that to achieve initial lysozyme-to-
57
nanoparticle ratios of 1:1, 10:1, 25:1, 50:1, 75:1, and 100:1 enzyme molecules per
nanoparticle, respectively.
100 L of purified nanoparticle suspension (~6.2·1013 nanoparticles) was added to
0.5 mL of each of the protein solutions. The mixtures were incubated overnight at room
temperature with gentle stirring. The protein-nanoparticle mixture was then treated with
0.1% Tween 20 for 2 h to remove any physically adsorbed enzyme. The unbound protein
along with the excess Tween 20 was then separated by either centrifugation or dialysis. In
the case of centrifugation, the suspensions were centrifuged at 13,000 g for 75 minutes.
The pellet was resuspended in 2mM MES buffer and centrifuged again. The supernatant
was collected in both steps to quantify the unbound protein. Separation by dialysis was
done similarly to dialysis of free nanoparticles described above. A cellulose ester
membrane with a MWCO of 100 kDa was used for dialysis to separate any unbound
lysozyme (molecular weight 14.3 kDa).
Bis-butyraldehyde PEG spacers (O=CH-CH2-CH2-CH2-(PEG)n-CH2-CH2-CH2CH=O, MW 3,700) were also used for conjugation of lysozyme to aliphatic amine
particles (Figure 4.1.B). Since attachment via PEG spacers minimizes interactions of
immobilized protein with the support [170, 260], lysozyme conjugated to nanoparticles via
PEG spacers was expected to show higher enzymatic activity. The conjugation protocol,
enzyme and nanoparticle concentrations, and purification methods were similar to those
described above for the glutaraldehyde-based attachment. Final concentration of bisbutyraldehyde PEG spacers was 25 mg/mL in all experiments. It should be noted that the
use of butyraldehyde chemistry enables protein conjugation specifically via its N-terminus
if the reaction is performed at pH 5.0 [261].
58
4.2.2.3. Protein conjugation to chloromethylated nanoparticles
Direct coupling of lysozyme to chlomethylated latex nanoparticles occurs via the
reaction of the latex CH2Cl- groups with the amine groups of lysozyme to give a
secondary amine (Figure 4.1.C). Conjugation in 2 mM MES buffer (pH = 5.0) was
performed overnight at room temperature. Six lysozyme solutions with the concentrations
of 8.82, 88.2, 220.5, 441, 661.5 and 882 g/mL were prepared in 2 mM MES buffer at pH
5.0 from a 5 mg/mL lysozyme stock solution. The dialyzed chloromethylated particles
suspension was diluted two-fold in 2 mM MES buffer, and 80 L of this suspension
(~1.9·1014 particles) was added to 0.5 mL of each of the protein solutions. Lysozyme
concentrations were chosen to achieve initial lysozyme-to-nanoparticle ratios of 1:1, 10:1,
25:1, 50:1, 75:1, and 100:1 protein molecules per nanoparticle, respectively. To remove
any physically adsorbed lysozyme, samples were treated with 0.1% Tween 20, and the
protein-nanoparticle conjugates were separated by either centrifugation or dialysis.
For conjugation via a spacer, a bifunctional NH 2-(PEG)n-COOH (MW 3,400) spacer
was employed. Dialyzed nanoparticle suspension was mixed with the spacer solution in
2 mM MES buffer. Final concentration of nanoparticles was ~1.9·10 15 mL-1 ; and final
concentration of the NH2-(PEG)n-COOH spacer was 5 mg/mL. The nanoparticle/spacer
mixture was incubated overnight at room temperature with gentle stirring. The amine
group of the spacer reacts with the chloromethyl group on the nanoparticles (Figure 4D).
The excess spacer was then separated by dialysis using 5 kDa MWCO membrane.
1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDAC) chemistry
(Hermanson. 1996) was used to conjugate lysozyme to the PEGylated nanoparticles. First,
25 mg/mL EDAC solution in 2 mM MES buffer was added to the dialysed nanoparticles
59
suspension (final EDAC concentration was 5 mg/mL). After 10 min incubation,
nanoparticles with activated carboxyl groups were added to lysozyme solutions.
Lysozyme and nanoparticle concentrations were the same as used for direct attachment to
chloromethylated particles. After treatment with Tween 20, unbound lysozyme, as well as
excessive reagents and reaction products, were separated by either centrifugation or
dialysis with a 100 kDa MWCO membrane.
Figure 4.1. Schematic of protein conjugation to nanoparticles. (A) Direct
conjugation to aminated nanoparticles. (B) Conjugation to aminated nanoparticles
via bis-butyraldehyde PEG spacer. (C) Direct conjugation to chloromethylated
nanoparticles. (D) Conjugation to chloromethylated nanoparticles via NH 2-PEGCOOH spacer.
60
4.2.2.4. Binding yield study
The number of bound lysozyme molecules per nanoparticle was determined as follows.
Samples with initial lysozyme concentrations corresponding to 1, 10, 25, 50, 75, and 100
molecules per nanoparticle were prepared. After the conjugation step, samples were
treated with Tween 20 to remove any non-covalently adsorbed protein and centrifuged at
13,000 g for 75 min. The clear supernatant was then collected and BCA protein assay was
used to determine the concentration of the unbound protein in the supernatant. Lysozyme
standards were prepared by diluting 5 mg/mL lysozyme stock solution in 2 mM MES
buffer containing 0.1% of Tween 20. The amount of bound protein was found from the
difference between the initial and unbound protein concentrations. Each of the
experiments on determination of binding efficiency was reproduced three times with
independently synthesized samples.
4.2.2.5.Enzyme activity assay with bacterial substrate
Protein-nanoparticle conjugates prepared from the suspension with the initial 25:1
lysozyme:nanoparticle ratio were used for all enzyme activity assays and for zeta-potential
measurements. A turbidometric enzyme activity assay was performed by monitoring the
decrease in optical density of an aqueous suspension of a natural substrate for lysozyme,
Micrococcus lysodeikticus, in potassium phosphate buffer. Micrococcus lysodeikticus was
preferred to other Gram-positive species because it is routinely used as a substrate for
assaying lysozyme activity (Shugar 1952). A 6 mM solution of potassium phosphate was
prepared by dissolving 0.898 g of potassium dihydrophosphate in 1 L of deionized water.
The pH of the solution was adjusted to 6.2 using a 0.1 M solution of potassium hydroxide.
A 0.018 wt. % suspension of Micrococcus lysodeikticus was prepared by suspending 3.75
61
mg of lyophilized cells in 20 mL of 6 mM potassium phosphate buffer solution. Proteinnanoparticle conjugates were then added to the bacterial suspension; net concentration of
lysozyme in all final mixtures was adjusted to 8.3 g/mL. Samples were incubated in a 12well microplate at room temperature, and the decrease in optical density at 405 nm was
monitored as a function of time. Initial rates of the reactions were calculated from the
slopes of the linear initial parts of the kinetic curves. Two independently synthesized
samples were studied for each type of nanoparticles. Control experiments were performed
with free lysozyme at the same enzyme concentration as in the samples. All kinetic
experiments were reproduced in triplicate.
4.2.2.6. Enzyme activity assay with a low molecular weight substrate
Lysozyme activity was also determined colorimetrically using a neutral oligosaccharide
substrate for lysozyme, p-nitrophenyl penta-N-acetyl-ß-chitopentaoside, PNP-(GlcNAc)5,
and was based on a coupled reaction with NAHase [262]. A 10 mM solution of PNP(GlcNAc)5 at pH 6.2 was prepared by dissolving 1.5mg/mL PNP-(GlcNAc)5 in 6 mM
potassium phosphate buffer. Lysozyme cleaves the first glycosidic bond of PNP(GlcNAc)5, giving the monochitoside, PNP-GlcNAc. NAHase [purchased as a suspension
in 2.5 M (NH4)2SO4 containing 1.2 mg protein/mL and diluted 1:100 in 6 mM phosphate
buffer] then catalyzes the release of the chromogen, p-nitrophenol, from the PNP-GlcNAc.
Addition of 50 mM Na2CO3 stops the reaction and yields a yellow color from the released
p-nitrophenolate. For activity measurements, 50 L sample aliquots were incubated with
150 L of PNP-(GlcNAc)5 and 10 µL of NAHase at 37C in a 96 microplate well. Six
reaction wells were prepared for each sample. Every 30 minutes, 100 µL of a 50 mM
62
Na2CO3 was added to one of the wells, and the absorbance at 405 nm was measured using
a microplate reader, while the reaction was allowed to continue in the other wells. Six time
points were taken for each sample. From these points, initial rates were determined and
used as a measure of enzyme activity. Two independently synthesized samples were
studied for each type of nanoparticles. Control experiments were performed with free
lysozyme at the same enzyme concentration as in the samples. All kinetic experiments
were reproduced in triplicate.
4.3. Results
4.3.1. Yield of binding.
Figure 4.2 shows efficiency of binding of lysozyme to polystyrene latex nanoparticles
without PEG spacers. In the case of attachment via PEG spacers, binding yields are
similar to those shown in Fig. 2. Binding yield in the case of aminated particles is in both
cases lower than that for their negatively charged counterparts. We speculate that this
difference is primarily due to electrostatic effects. The initial step of covalent attachment
of lysozyme to nanoparticles involves its physical adsorption (Suen and Morawetz 1985;
Sarobe et al. 1998; Molina-Bolivar and Galisteo-Gonzales 2005; Giacomelli et. al 2000).
This adsorption is primarily determined by the interactions of the hydrophobic parts of
lysozyme molecules with the hydrophobic surface of polystyrene latex. However, it can be
expected that more positively charged lysozyme molecules (pI=10.6) can be adsorbed to a
negatively charged surface, than to a positively charged surface. This difference in the
initial physical adsorption results in different binding efficiency. Notably, a theoretical
monolayer of lysozyme on a 20 nm spherical particle calculated from the dimensions of
63
the lysozyme molecule (2.5 nm X 3.5 nm X 3.5 nm) should consist of ~120 molecules per
nanoparticle [263].
Figure 4.2. Efficiency of covalent attachment of lysozyme to polystyrene latex
nanoparticles for samples prepared by (A) direct conjugation and (B) conjugation
via PEG spacers. Black squares correspond to aminated nanoparticles and red
circles correspond to chloromethylated nanoparticles, respectively.
At high initial lysozyme concentrations, the number of attached lysozyme
molecules exceeds the number of chloromethyl groups available for attachment (see Table
4.1). However, it should be noted that lysozyme tends to form oligomers (dimers and
tetramers) at pH>4.0 [264]. The degree of oligomerization can also be affected by
interaction with polystyrene latex particles. The observed excessive conjugation is
consistent with the attachment of oligomers. Possible presence of physically adsorbed
lysozyme that cannot be removed by treatment with 0.1% Tween 20 also cannot be
completely ruled out. It should be noted, however, that according to the literature data
(Peula et al. 1995), treatment by Tween 20 removes 75 - 85 % of physically adsorbed
64
protein from polystyrene latex microspheres. We observed binding yields of ~80% in the
case of lysozyme attachment to chloromethylated nanoparticles after treatment with
Tween 20. It is thus unlikely that non-covalent adsorption of lysozyme plays an important
role in the observed phenomenon.
4.3.2. Bacterial turbidometric activity assay
Protein-nanoparticle conjugates prepared from the suspension with the initial 25:1
lysozyme: nanoparticle ratio were used for all enzyme activity assays. A bacterial
turbidometric activity assay was performed by monitoring the decrease in optical density
of an aqueous suspension of Micrococcus lysodeikticus. Lysozyme cleaves the β (1-4)glycosidic bond between N-acetylmuramic acid (MurNAc) and the N-acetyl-Dglucosamine (GlcNAc) in the polysaccharide that forms the backbone of the bacterial cell
walls. This cleavage is the first step in the lysis of bacterial cells. The degree of lysis is
measured as reduction in turbidity of the bacterial suspension (Shugar 1952). We
monitored the decrease in optical density at 405 nm for two sets of samples: a) samples
purified by centrifugation and b) samples purified by dialysis. To take into account
possible precipitation of bacteria from the suspension, the turbidity of the bacterial
suspension alone was monitored in a control experiment.
65
B
A
C
D
Figure 4.3. Results of bacterial lysis assay. (A) Direct attachment of lysozyme to
nanoparticles; samples purified by centrifugation, (B) attachment via PEG spacer;
samples purified by centrifugation, (C) Direct attachment of lysozyme to
nanoparticles; samples purified by dialysis, and (D) attachment via PEG spacer;
samples purified by dialysis. Black squares – lysozyme attched to positively
charged nanoparticles; red circles – lysozyme attached to negatively charged
nanoparticles, blue triangles – free lysozyme.
In the first set of samples, the final step of purification of protein-nanoparticle
conjugates was centrifugation. Figure 4.3 compares the rates of bacterial lysis for
66
lysozyme directly conjugated to nanoparticles to that of free lysozyme. The initial rate of
the lysis reaction was calculated from the slopes of the linearized kinetic curves. We
observed significantly faster bacterial degradation for lysozyme conjugated to aminated
nanoparticles (Table 4.2). Bacterial lysis rates for lysozyme conjugated via PEG spacer
were not significantly different from those observed for lysozyme directly attached to the
same type of nanoparticles.
Initial rate from linearized slopes for
sample purified by
Sample
dialysis
centrifugation
4.90.1 · 10-4
Free lysozyme
Lysozyme directly conjugated to aminated
nanoparticles
8.90.2 · 10-4
7.00.1 · 10-4
Lysozyme conjugated to aminated
nanoparticles via PEG spacer
9.30.2 · 10-4
6.50.1 · 10-4
Lysozyme directly conjugated to
chloromethylated nanoparticles
2.70.1 · 10-4
2.40.1 · 10-4
Lysozyme conjugated to chloromethylated
nanoparticles via PEG spacer
2.40.1 · 10-4
2.50.1 · 10-4
Table 4.2. Initial rates of the bacterial lysis study for samples purified by dialysis and
centrifugation
In an another study, we also studied the effect of controls, positively charged aliphatic
amine and negatively charged chloromethylated nanoparticles on lytic activity of
Micrococcus lysodeikticus.. No detectable lytic activity was observed for any of the
controls. Figure 4.4 compares the rates of bacterial lysis for both types of polystyrene
latex nanoparticles along with control bacterial cell suspension in the same study
67
Figure 4.4. Results of bacterial lysis assay. Pink circles – Aliphatic amine
nanoparticles; Light Blue squares – Chloromethyl nanoparticles, Yellow squares –
Control bacterial cells.
We also studied the effect of purification method on the activity of the conjugates.
The unbound protein was separated from protein-nanoparticle conjugates by either
dialysis or centrifugation. Figure 4.3 (B) shows results of bacterial lysis assay for samples
purified by centrifugation, and Figure 4.3 (A) shows the data for samples purified by
dialysis. Table 4.2 summarizes results of bacterial lysis assay. As can be seen from Table
4.2, dialyzed samples always show slightly higher bacterial lysis rates than those purified
by centrifuging. A possible explanation of the effect of purification method is higher
degree of nanoparticle aggregation in the case of centrifugation. As a result of
aggregation, some lysozyme molecules can be trapped inside the aggregates and blocked
from the interaction with their substrate. Figure 4.5 compares representative AFM images
of lysozyme conjugated to aminated nanoparticles and purified by dialysis (Fig. 4.3A) or
68
centrifugation (Fig. 4.3B). As can be seen from Fig.4.5, much larger aggregates are
observed in the case of samples purified by centrifugation. Since samples purified by
dialysis showed higher activity than those purified by centrifugation, dialysis has been
used as the primary purification method in all further experiments.
A
B
Figure 4.5. Tapping mode AFM images comparing the degree of aggregation for
samples purified by dialysis and centrifugation. Scan size is 5 m x 5 m and Z
scale is 100 nm. (A) Lysozyme conjugated to aliphatic amine nanoparticles after
purification by dialysis and (B) lysozyme conjugated to aliphatic amine
nanoparticles after purification by centrifugation.
4.3.3. Enzyme activity assay with a low molecular weight substrate
PNP-(GlcNAc)5 is a chromogenic pentachiteoside that serves as an alternative substrate
for lysozyme [265, 266]. The color is produced by free p-nitrophenol (PNP), liberated
from PNP-(GlcNAc)5 through a coupled reaction of lysozyme and NAHase. First,
PNP-(GlcNAc)5 is cleaved by lysozyme at the first glycosidic linkage, releasing
monochiteoside PNP-GlcNAc [266]. The presence of NAHase is essential for the assay to
promote the liberation of free p-nitrophenol. Addition of Na2CO3 at the final step
69
increases the pH of the reaction mixture, stops the reaction and reveals the yellow color of
PNP.
Figure 4.6. Lysozyme activity assay with low molecular weight substrate
PNP-(GlnAc)5. (A) Direct attachment and (B) attachment via PEG spacer. Black
squares – lysozyme attached to positively charged nanoparticles; red circles –
lysozyme attached to negatively charged nanoparticles, blue triangles – free
lysozyme.
Figure 4.6 shows the results of the assay with PNP-(GlcNAc)5. In this case, free
lysozyme shows higher activity than lysozyme conjugated to both types of nanoparticles.
The difference between the activities of lysozyme attached to aminated and
chloromethylated nanoparticles is much less pronounced than in the case of bacterial lysis
assay. To obtain numerical values of kinetic constants, kinetic curves were empirically
linearized in coordinates Log [Optical Density] – Time.
70
Sample
Free enzyme
Initial rate from linearized
slopes
5.10.5 · 10-3
Lysozyme directly conjugated to aminated nanoparticles
4.40.3 · 10 -3
Lysozyme conjugated to aminated nanoparticles via PEG
spacer
Lysozyme directly conjugated to chloromethylated
nanoparticles
Lysozyme conjugated to chloromethylated nanoparticles
via PEG spacer
4.90.3 · 10-3
4.30.4 · 10-3
4.4 0.3 · 10 -3
Table 4.3. Initial rates of enzyme activity with low molecular weight substrate for all
samples
4.3.4. Zeta potential analysis
Zeta potential studies were performed to characterize electrostatic properties of
colloidal suspensions used in this study. Table 4.4 shows zeta potential values averaged
over 4 independent experiments for each of the samples. As expected, Micrococcus
lysodeikticus in aqueous suspension has highly negative zeta potential. Highly positive
zeta potentials are observed for aminated nanoparticles; zeta potential values slightly
decrease upon direct coupling of lysozyme and further decrease upon protein conjugation
via bis-butyraldehyde spacers.
It is currently not clear why conjugation of positively charged lysozyme to positively
charged nanoparticles reduces their zeta-potential. One possible explanation of this fact is
increase of hydrodynamic resistance of the conjugates. Presence of a “fluffy” lysozyme
layer may decrease electrophoretic mobility of nanoparticles. Since the absolute value of
zeta-potential is proportional to the electrophoretic mobility, this will result in the decrease
of zeta-potential. This explanation is in agreement with the observed higher drop of zeta71
potential for the case of lysozyme attached to nanoparticles via PEG spacers, in which
lysozyme has even more spatial freedom. Notably, even in the latter case zeta potential
remains highly positive and significantly exceeds that of free lysozyme. Stronger
electrostatic interactions resulting in more effective delivery can thus be expected between
enzyme-nanoparticle conjugates and Micrococcus lysodeikticus.
Sample
Bare aminated nano-particles (control)
Lysozyme directly conjugated to
aminated nanoparticles
Lysozyme conjugated to aminated
nanoparticles via PEG spacer
Bare chloromethylated nanoparticles
(control)
Lysozyme directly conjugated to
chloromethylated nanoparticles
Lysozyme conjugated to
chloromethylated nanoparticles via PEG
spacer
Bacterial substrate
Lysozyme
Zeta potential,
mV
+ 37.3  1.1
+ 31.5  1.7
+ 27.2  1.7
- 43.3  1.5
- 32.0  1.6
- 25.0  1.4
- 27.8  1.1
+ 8.84  1.2
Table 4.4 - Results of zeta potential measurements.
Bare chloromethylated nanoparticles have highly negative zeta potential. Its absolute
value decreases upon direct coupling of lysozyme and further decreases upon coupling of
lysozyme via PEG spacers, consistent with the hypothesis of decreased electrophoretic
mobility of the particles upon protein attachment. Yet for all protein conjugates with
chloromethylated particles zeta potential remains highly negative hindering their
interactions with the bacteria.
72
4.4. Discussion
Electrostatic interactions play an important role in targeting lysozyme to its
substrate bacteria [267-271]. Lysozyme has pI of 10.6 [271] and therefore bears
significant positive charge (+8 or +9 per molecule) at neutral pH [272]. Most bacterial cell
walls, on the other hand, are negatively charged due to the presence of teichoic,
lipoteichoic, and teichuronic acids in the cell membrane [273, 274].
Previous results indicate that positive charge is essential for lysozyme‟s
antibacterial function but not to activity towards low molecular weight substrates.
Acetylation of all six free lysine residues of lysozyme abolished action towards
Micrococcus lysodeikticus but did not affect the cleavage of the tetramer of Nacetylglucosamine, (GlnAc)4, from chitin [275]. T4 bacteriophage lysozyme lost > 90% of
its native activity in bacterials lysis assay with Micrococcus lysodeikticus when adsorbed
onto 9 nm silica nanoparticles, which are strongly negatively charged [276]. At the same
time, lysozyme adsorbed onto similarly negatively charged silica nanoparticles (4, 20, and
100 nm in diameter) retained ~ 80% of its native activity when assayed with a neutral
substrate, PNP-(GlnAc)5 [263].
The results of this study indicate that nanoparticle charge can be used to control
bacteriolytic activity of lysozyme. Lysozyme attached to positively charged nanoparticles
shows significantly higher activity in bacterial lysis assay than free lysozyme. The
observed effect is probably due to the differences in targeting of the enzyme to negatively
charged bacterial cell walls (Figure 6) because similar activities are observed for lysozyme
attached to both positively and negatively charged nanoparticles when assayed with a
neutral low molecular weight substrate, PNP-(GlnAc)5. These activities are somewhat
73
lower than that of free enzyme, consistent with the previous data [276-279] on decreased
activity of immobilized enzymes due to conformational changes arising from the
interactions with nanoparticle surface and partial screening of the active site by the
nanoparticle.
Figure 4.7 shows charge-directed targeting of lysozyme to bacterial cell walls
The hypothesis of electrostatic contribution to targeting nanoparticles to bacteria is
confirmed by zeta-potential measurements, which show negative zeta potential for
Micrococcus lysodeikticus suspension and lysozyme conjugated to negatively-charged
nanoparticles, but a positive zeta potential for free lysozyme and lysozyme attached to
positively-charged nanoparticles. Notably, zeta potential for lysozyme attached to
positively charged nanoparticles is more positive than that of free lysozyme, in good
agreement with the observed higher activity of the former. On the other hand, electrostatic
interaction is probably not the only factor that mediates targeting of lysozyme to bacterial
74
cells because lysozyme attached to negatively-charged particles also shows some
bacteriolytic activity in spite of the fact that these nanoparticles should be electrostatically
repelled from bacterial cell walls.
Another important result of this study is the development of a protein attachment
protocol that minimizes nanoparticle aggregation. Aggregation often has been observed
previously [280-282]and presents a serious problem for the use of nanoparticles in
biomedical applications because of the uncontrollable size of the aggregates and
broadening of the distributions of their functional properties. Here, we observed that
samples with higher degree of aggregation show decreased enzymatic activity of the
immobilized protein. The protocol developed in this work uses small amount of surfactant,
Tween 20, to remove non-covalently bound protein and employs dialysis for purification
from unbound proteins and small molecules. Optimization of the concentrations of the
components enabled us to significantly reduce degree of aggregation, as can be seen from
representative AFM images in Figure 4.5.
We also studied the effect of attachment via PEG spacers on enzymatic activity of
lysozyme. No significant difference in activity has been observed for lysozyme attached
via a spacer compared to lysozyme directly attached to nanoparticle surface. This result
was somewhat contradictory to our expectations because previous research [170, 260]
demonstrated a beneficial effect of attachment via spacer on activity of immobilized
enzymes. This effect was attributed to reduced interaction of the enzyme with the surface
of its support due to the presence of the spacer. However, lysozyme is known to be a
“rigid” enzyme [283], which reluctantly changes its structure. In addition, we use Tween
20, which blocks the hydrophobic polystyrene surface and produces oligoethyleneglycol-
75
terminated surface. Thus, interaction of lysozyme with the nanoparticle surface is likely to
be reduced even in the case of direct conjugation due to the presence of Tween 20. This
agrees well with the small drop in enzymatic activity of the protein-nanoparticle
conjugates, as compared to free lysozyme, in the assay with PNP-(GlnAc)5. Therefore, use
of PEG spacers does not have a considerable effect on enzymatic activity of lysozyme.
In conclusion, nanoparticle charge has been used to controllably change activity of a
covalently immobilized protein. Enhanced charge-directed targeting of lysozymenanoparticle conjugates to bacteria was achieved using positively charged aliphatic amine
nanoparticles as the protein carriers. This result can be useful for development of
antibacterial, antibiotic-free protein-nanoparticle conjugates.
76
CHAPTER 5
ANTIBODY-DIRECTED TARGETING OF LYSOSTAPHIN ADSORBED TO
POLYLACTIDE NANOPARTICLES INCREASES ITS ANTIMICROBIAL
ACTIVITY AGAINST S.AUREUS IN VITRO
5.1. Introduction
Staphylococcus aureus is a versatile and virulent Gram-positive pathogen that is
frequently found among clinical isolates from patients in the United States. It is capable of
causing a wide range of infections, being one of the major causes of bacteremia and is
associated with significant mortality [39]. It produces a variety of virulence factors, with
both structural and secreted products playing a role in the pathogenesis of infection. The
rates of infections caused by both community and hospital-acquired strains are increasing
steadily and treatment of these infections through conventional antibiotic therapy is
becoming more and more difficult because of the increasing prevalence of multidrugresistant strains [44]. The decreased effectiveness of current antibiotic treatments due to
resistance has spurred the search for alternative approaches to antimicrobial therapy.
Currently, much attention is paid towards the use of natural antimicrobial proteins
and peptides as antibacterials to overcome the problems associated with antibiotic
resistance [284]. Naturally occurring antimicrobial peptides and enzymes are characterized
by high activity, broad antimicrobial spectrum, and low rates of resistance [285, 286].
Lysostaphin is an antimicrobial enzyme produced by Staphylococcus staphylolyticus. It
belongs to the class of Endopeptidases, which specifically cleave cross-linked
pentaglycine bridges in the peptidoglycan of S. aureus, thereby hydrolyzing the cell wall
and lysing the bacteria [287].
77
In spite of the great enthusiasm regarding the prospect of developing antibacterial
enzymes and peptides as a new generation of antibacterial agents, they are still at an early
stage of technologic maturation, and many hurdles have yet to be overcome [288].
Because of their peptidic nature, they could present the following problems: 1) short half
life in vivo, 2) loss of activity in physiological conditions (especially at high ionic
strength, which may interfere with their charge-directed targeting and mechanism of
action), 3) toxicity and immunogenicity, 4) poor storage stability, and 5) interference with
normal bacterial flora that may arise when attempts are made to use those peptides as
antibacterial agents (3). Availability of a technology for targeted delivery of antimicrobial
peptides could provide a valuable contribution towards addressing most if not all of these
problems. Most importantly, targeted delivery can improve efficacy and reduce systemic
toxicity of antimicrobial peptides by reducing their action against non-specific targets.
Although much work has been done recently on design of drug carriers for sitespecific delivery, studies of targeted drug delivery have been thus far primarily focused on
anticancer drugs [289] and certain other therapeutics targeting the host cells and tissues
[290, 291]. Much less attention has been paid up to date to studies of targeted delivery of
antimicrobial agents. Recently, mannosylated or galactosylated polystyrene latex
nanoparticles were shown to result in agglutination of mutant strains of E. Coli
overexpressing mannose and galactose surface receptors, respectively [228, 229]. Targeted
photothermal lysis of pathogenic bacteria such as Salmonella and Pseudomonas
aergusinosa has been studied using antibody-conjugated gold nanoparticles and nanorods,
respectively [292, 293]. Our group has previously studied charge-directed targeting of
lysozyme conjugated to nanoparticles and demonstrated enhanced antimicrobial activity
78
against Micrococcus lysodeikticus for enzyme conjugated to positively charged
nanoparticles
[250,
294].
Enhanced
efficacy
against
Gram-positive
Listeria
Monocytogenes has also been demonstrated for lysozyme co-immobilized with anti- L.
Monocytogenes antibody on polystyrene latex NPs [295, 296]. However, a systematic
study of activity of targeted enzyme-NP conjugates with different enzyme and antibody
loadings against clinically relevant pathogens has not been performed previously. With
this in mind, the goal of the present work was a detailed study of antimicrobial activity of
targeted (lysostaphin-antibody-NPs) and untargeted (lysostaphin-NPs) conjugates against
S. aureus at different enzyme and antibody ratios.
5.2. Materials and methods
Staphylococcus aureus (ATCC 27660) was purchased from American Type Culture
Collection (Manassas, VA). Lysostaphin (26 kDa) isolated from Staphylococcus
staphylolyticus was purchased from AMBI products LLC (Lawerence, NY). Poly (DLlactide) and Bovine Serum Albumin (BSA) were purchased from Sigma Aldrich (St.
Louis, MO). Immunogen-specific Rabbit Polyclonal antibody to Staphylococcus aureus
(ATCC 27660) formulated in a phosphate saline buffer (0.01M, pH 7.2) was purchased
from ViroStat (Portland, ME). Alexa Fluor 350 and Alexa Fluor 594 Carboxylic acid,
Succinimidyl ester dyes used for fluorescent labeling were purchased from Invitrogen,
Molecular Probes (Carlsbad, CA).
Microsep Centrifugal Devices with a modified
Polyethersulfone membrane (low protein binding; MWCO - 3K and 30K) used for protein
purification were purchased from Pall Life Sciences (Ann Arbor, MI). Fluorescence and
turbidimetric activity assays were performed using a Synergy microplate reader (Bio-Tek
79
Instruments, Inc., Winooski, VT). Particle sizing and Zeta potential analysis were done
using a 90plus Particle Size Analyzer (Brookhaven Instruments Co, Holtsville, NY).
5.2.1. Preparation of nanoparticles
Polylactide nanoparticles were prepared using a nanoprecipitation method based on
solvent diffusion. Briefly, Poly(DL-lactide) (Mw 97,500) was dissolved in 25 ml of
acetone at a concentration of 8 mg/ml and this solution was then dispersed in an aqueous
phase (200 ml) containing Pluronic F68 (5 mg/ml) under sonication for 15 min. The
acetone solution was added into the aqueous phase dropwise at a uniform rate. The
resulting suspension containing the nanoparticles was then purified from excess of
polymer and Pluronic by triple centrifugation followed by redispersion in DI water.
Centrifugation was performed at 3,000 rcf for 1 hr at 25°C using Allegra™ 64R
Centrifuge (Beckman Coulter, USA). Nanoparticle suspension in DI water was stored in
refrigerator at 4°C. Concentration of PLA in the suspension was determined using
gravimetric analysis. The number of nanoparticles per ml of the suspension was then
calculated using the size of nanoparticles determined from DLS experiments (see below)
and assuming PLA density of 1.25 g/cm3 [297].
5.2.2. Fluorescent labeling of proteins
Alexa Fluor succinimidyl ester dyes (Alexa Fluor 350 and Alexa Fluor 594) were used for
fluorescent labeling of the antibody and lysostaphin, respectively. The dyes react with the
primary amines of proteins at pH 7.5–8.5 yielding stable conjugates [298]. Approximately
1 μg of Alexa Fluor 350 (ε 19,000) was dissolved in 100 μl of dimethylformamide and 25
μl of the solution was then added to 2 ml of a 1 mg/ml solution of S. aureus antibody in 10
80
mM PBS (10 mM phosphate; 140 mM NaCl, 3 mM KCl). A similar procedure was performed
for labeling of lysostaphin using Alexa Fluor 594 (ε 73,000). The samples were incubated
at 25°C for 2 h, diluted two fold in 10 mM PBS and then centrifuged at 3,000 rcf for 1 h
using a Nanosep® membrane centrifugal device (MWCO 3K and 30K for lysostaphin and
anti-S. aureus antibody, respectively) to remove any unbound dye. As per the
manufacturer‟s instructions, the molecular weight cutoff (MWCO) of approximately 6
times less than the molecular weight of the protein to be retained was chosen for
purification to achieve maximum sample recovery.
5.2.3. Protein adsorption to PLA nanoparticles
Adsorption of Alexa 594 labeled lysostaphin with or without Alexa 350 labeled S.
aureus antibody to PLA nanoparticles was studied at different NP: lysostaphin and NP:
lysostaphin: antibody ratios. The ratios used for the preparation of lysostaphinnanoparticle conjugates corresponded to 1300, 2700 and 5400 lysostaphin molecules per
nanoparticle. For protein adsorption, 900 μl each of lysostaphin solutions with
concentrations of 5.68, 11.79 and 23.59 μg/ml were prepared in 1 mM PBS buffer from a
1.25 mg/mL lysostaphin stock solution and combined with 100 μl of the NP stock solution
containing 9.8*1011 particles per ml.
The ratios for targeted lysostaphin-antibody-nanoparticle conjugates consisted of
1300: 450; 2700: 900 and 5400: 1800 lysostaphin and antibody molecules per
nanoparticle, respectively. Three protein solutions containing both lysostaphin and
antibody (900 μl each) were prepared in 1 mM PBS buffer from lysostaphin and antibody
stock solutions. Lysostaphin and antibody concentrations for each of the ratios consisted
of 5.68 : 10.9 μg/ml for sample 1300: 450; 11.79 : 21.9 μg/ml for sample 2700: 900; and
81
23.59 : 43.8 μg/ml and 87.6 μg/ml antibody for sample 5400: 1800. 100 μl of the
nanoparticle stock solution (containing 9.8*1011 particles per ml) was added to each of the
protein solutions to make up the final volume of 1 ml.
All sample mixtures were then incubated for 6 h in a shaker at 150 rpm at 20°C.
The samples were then centrifuged at 3,000 rcf for 1 h to remove any unbound enzyme
and antibody. The supernatant was collected and the pellet was resuspended in 1 ml of 10
mM PBS and stored at 4 °C. The concentration of unbound enzyme and antibody in each
of the supernatants was determined from fluorescence measurements. Enzyme and
antibody solutions with known concentrations were used as the standards. The
concentrations of bound enzyme and antibody in the resuspended pellets were found from
the difference between the initial concentration and concentration in the supernatant. The
signal-to-background ratio was ~ 500: 1 for the fluorescently labeled lysostaphin (with or
without antibody) and ~ 1000:1 for fluorescently labeled antibody. Protein adsorption
experiments were reproduced in triplicates.
A second batch of protein-NP conjugates was also synthesized using newly
prepared NPs to verify batch-to-batch reproducibility of antibacterial activity. Unlabeled
proteins were used for the preparation of the second batch; otherwise all synthesis
procedures were identical to those described above. Particle sizing and zeta potential
measurements were also performed for the second batch.
5.2.4. Particle Size Measurements and Zeta potential analysis
Stock solutions containing nanoparticles or protein-nanoparticle conjugates were diluted
1:1000 in 10 mM PBS in a disposable cell and particle size was measured using dynamic
light scattering with a 90Plus particle size /zeta potential analyzer (BrookHaven
82
Instruments Corporation, Holtsville, NY). Before particle sizing, the samples were
sonicated for 20 min to break up any loosely-held agglomerates. Particle size was
expressed as a combined mean diameter in nm (+/- SD). For zeta potential analysis, stock
solutions were diluted 1:100 in 1 mM PBS in a disposable cuvette and zeta potential was
measured using the 90 Plus analyzer. Zeta potential was expressed in millivolts (mV) as a
combined mean of (+/- SD).
5.2.5. Bacterial turbidimetric activity assay
A cell suspension of S. aureus was prepared by inoculating 10 ml of 30% (w/v)
tryptic soy with 100 µl of S. aureus culture prepared according to ATCC instructions. The
procedure used was similar to the one employed by Schindler and Schuhardt [299]. The
cells were incubated at 37C for 18 - 24 h under gentle shaking to grow to a mid-log
phase. The cells were then pelleted at 2,000 rpm for 10 min and washed twice with 10 mM
PBS. The cells were then resuspended in PBS to prepare bacterial suspension with optical
density (OD600) of 0.56 at 600 nm (1 cm light path). The turbidity assay was performed
by adding 100 µl of the diluted sample solution in a 96 well microplate followed by 100µl
of thus prepared S. aureus cell suspension. Lysostaphin concentration was fixed for all
samples and consisted of 0.83 g/ml. The decrease in OD600 was monitored at 37°C in 1
min intervals for a total period of 3 h. These kinetic data were fitted exponentially, in
assumption of the first order kinetics with respect to bacteria. The mean kinetic constants
determined from these exponential fits were used as the measure of lysostaphin activity in
a particular sample. Control experiments were performed with free lysostaphin at the same
83
enzyme concentration. All kinetic experiments were reproduced in triplicate for two
different batches of samples synthesized independently (n=6).
5.2.6. Fractional analysis of enzyme activity in leached samples
To determine whether antibacterial activity of the enzyme-NP conjugates comes
primarily from adsorbed or leached enzyme, we performed fractional analysis of enzyme
activity. Two samples corresponding to the highest lysostaphin and antibody coatings
(1:5400:0 and 1:5400:1800 NP: lysostaphin: antibody ratios) were prepared using the
procedure described above. The samples were then incubated in PBS buffer at 37C with
gentle shaking. 100 l of each sample was taken at each of the different time points (1, 6
and 24 h) and centrifuged at 3,000 rcf for 1 h followed by collection of the supernatants. A
cell suspension of S. aureus was grown to a mid-log phase similarly to the procedure
described previously, centrifuged at 2,000 rpm for 10 min, washed twice and resuspended
in 10 mM PBS. 100 µl of the cell suspension was added to wells of a 96 well plate
containing either 100 µl of the total sample or 100 µl of the supernatant collected at a
particular time point (leached aliquot). The samples were incubated with the cell
suspension at 37ºC under continuous shaking, and the rate of bacterial lysis was monitored
for 3 h, similarly to the procedure described in the previous section. From these
experiments kinetic constants for bacterial lysis were determined using the exponential fits
and used as the measure of leached lysostaphin activity at different time points. All kinetic
experiments were reproduced in triplicates.
5.2.7. Statistical analysis
84
Data are presented as means±SD. Paired t-test and one-way analysis of variance
(ANOVA) was performed for the binding efficiency data of proteins on NPs and kinetics
constants in the bacterial lysis assay. The null hypothesis was that the means are not
significantly different. Tukey‟s test was used to compare different samples in the event
that the ANOVA null hypothesis was not true. Data were analyzed using Origin 6 Data
Analysis and Graphing Software (OriginLab Corporation, MA, USA). A p value of <0.05
was considered statistically significant.
5.3. Results and Discussion
5.3.1. Characterization of enzyme-NP conjugates
Lysostaphin and anti S. aureus antibody was adsorbed onto biodegradable PLA
NPs (Figure 5.1). We found that adsorption was preferred method of immobilization in
this case because it resulted in strong protein attachment and high activity without the
need for additional chemical modification steps. Also, adsorption can be carried out in
milder chemical conditions, generally resulting in better protein stability and is more cost
effective. Adsorption has been successfully used previously to co-immoblize two proteins
on PLA NPs for in vivo applications [300]. Leaching studies discussed in detail in the
following section were performed to demonstrate that major part of antibacterial activity is
associated with the enzyme conjugated to NPs. Samples with several enzyme : antibody :
nanoparticle ratios were prepared (Table 5.1). The initial protein concentrations were
chosen to correspond to approximately 0.5, 1, and 2 monolayers for lysostaphin and
antibody on the NPs, respectively.
85
Figure 5.1. Schematic of the synthesis of protein-NP conjugates.
The following assumptions were used when calculating amount of a protein
required for a monolayer coating. The average dimensions of the lysostaphin molecule are
4.5 nm x 5.5 nm x 8.5 nm [301]. The average surface area occupied by an adsorbed
lysostaphin molecule based on the smallest dimensions was therefore assumed to be 25
nm2. The size of uncoated PLA NPs measured using dynamic light scattering was found to
be 145 nm (Table 5.2). A theoretical monolayer of lysostaphin on the surface of a 145 nm
PLA nanoparticle calculated from the smallest dimensions of the lysostaphin molecule
should consist of ~2,700 molecules per NP. The average dimensions of an IgG antibody
molecule reported in the literature vary greatly from 14 nm x 8.5 nm x 4 nm to 23 nm x 23
86
nm x 4.4 nm [302]. We therefore used hydrodynamic radius, Rh, of IgG molecule for
estimation of its theoretical monolayer coating. Rh can be determined more accurately and
its value reported for IgG molecule in aqueous solution, Rh=5.3 nm, is consistent in the
literature [303]. A theoretical monolayer of antibody calculated in the assumption of dense
hexagonal packing of the spheres with the radius of 5.3 nm would consist of ~900
molecules per NP.
A
87
B
Figure 5.2. Binding yield of adsorption on PLA NPs for (A) AlexaFluor 594-labeled
lysostaphin (Black squares – Plain lysostaphin and Red circles – lysostaphin
coadsorbed in the presence of antibody (n = 3)), and (B) Alexa Fluor 350 labeled
anti-S. aureus antibody (n = 3)
Binding yields for lysostaphin consisted of 40-44% for plain lysostaphin-coated
samples and 29-38% when lysostaphin was co-adsorbed with the antibody (Table 5.1).
Binding yield for the antibody consisted of 15-23% for different ratios. Overall, amount of
adsorbed lysostaphin and the antibody increased linearly with the increase of the
concentration of the corresponding protein (Figure 5.2 A, B). For lysostaphin adsorption at
the highest initial enzyme concentrations (which corresponded to two monolayers of
lysostaphin), coating by the adsorbed enzyme approached theoretically predicted
monolayer and consisted of 85% and 77 % of a monolayer for plain and antibody-coated
samples, respectively (Table 5.1). At the same time, antibody coating at its highest initial
concentration consisted of ~30% of theoretical monolayer. This is an assumption that is
based on theoretical dimensions of lysostaphin and antibody molecules adsorbed on
88
nanoparticles. However, the actual monolayer coating could be higher as enzymes are
known to unfold upon conjugation resulting in a slightly higher specific surface area
occupied by each enzyme molecule.
Initial Ratio, protein
Number of theoretical
Binding yield
Fraction of protein
molecules per
monolayers assuming
of protein, %
monolayer adsorbed,
nanoparticle
all added protein
%
were adsorbed
1NP : 1300 LS
0.5 LS
402 LS
201 LS
1NP : 2700 LS
1 LS
445 LS
455 LS
1NP : 5400 LS
2 LS
423 LS
856 LS
1NP :1300 LS : 450 Ab
0.5 LS, 0.5 Ab
293 LS; 234 Ab
151 LS; 122 Ab
1NP :2700 LS : 900 Ab
1 LS, 1 Ab
383 LS; 177 Ab
391 LS; 177 Ab
1NP :5400 LS : 1800
Ab
2 LS, 2 Ab
392 LS; 151 Ab
763 LS; 302 Ab
Table 5.1. Characteristics of samples with different nanoparticle (NP): lysostaphin (LS):
antibody (Ab) ratios. Binding yield of protein on NPs. Data are presented as meanSD
(n=3).
Overall lower binding yields for antibody compared to lysostaphin could be
attributed to stronger electrostatic interaction between the highly positively charged
lysostaphin molecules (pI = 10.6) and negatively charged PLA NPs (zeta potential -48
mV). For comparison, pI of IgG lies in the range 6.4-9.0 and because of the polyclonal
89
nature of antibody, molecules with different pIs from this range are expected to be present
in the solution [304].
Size was determined by light scattering for bare NPs and protein-NP conjugates
(Table 5.2). The mean diameter of the enzyme-NP conjugates increased significantly
(p<0.05) upon adsorption of lysostaphin with or without antibody to the surface of NPs.
Zeta potentials for all conjugates were highly negative (<-30 mV) indicating that the
conjugates, similarly to bare NPs, were negatively charged. The mean values from two
different batches are tabulated in Table 5.2 below.
Ratio, protein
molecules per
nanoparticle
Number of
theoretical
monolayers if all
added protein
molecules were
adsorbed
Size (nm)
(Mean  SD)
Zeta potential (mV)
(Mean  SD)
1NP : 1300 LS
0.5 LS
177.4 ± 6.8
-39.7 ± 11.6
1NP : 2700 LS
1 LS
182.6 ± 6.8
-39.6 ± 13.2
1NP : 5400 LS
2 LS
187.3 ± 4.7
-42.6 ± 10.3
1NP :1300 LS : 450 Ab
0.5 LS, 0.5 Ab
182.7 ± 1.8
-45.3 ± 1.9
1NP :2700 LS : 900 Ab
1 LS, 1 Ab
193.3. ± 7.8
-41.1 ± 4.1
1NP :5400 LS : 1800 Ab
2 LS, 2 Ab
180.9 ± 9.8
-33. ± 1.4
148.1 ± 4.1
-49.5 ± 2.1
Uncoated NPs
Table 5.2. Characteristics of samples with different nanoparticle (NP):lysostaphin
(LS):antibody (Ab) ratios - Size and zeta potentials for all samples.
90
5.3.2. Fractional enzyme activity from leaching study
It is important to know whether activity of enzyme coated NPs is due to
immobilized or leached lysostaphin. Fluorescence measurements were found to be not
sensitive enough to detect enzyme leached from the NPs into the supernatant. Therefore,
turbidimetric bacterial lysis assay was employed to determine the fraction of total activity
contributed by leached lysostaphin. This method was found to be more sensitive because
even extremely small concentrations of lysostaphin can cause significant lysis of bacteria
in S. aureus suspension.
Figure 5.3 shows the percent of leached enzyme activity for the aliquots collected
at different time points compared against total activity for two samples with highest
lysostaphin loading. These results indicate that less than 10% of enzyme was desorbed
from the NPs after 24 h incubation in PBS buffer. Shah et al. studied the antimicrobial
activity of lysostaphin adsorbed on two different plastic surfaces, polystyrene (well plates)
and fluorinated ethylene-propylene polymer (a Teflon-like material used in Angiocath
catheters) [240]. Similarly to our findings, they observed that the antibacterial activity was
primarily due to immobilized and not leached enzyme.
91
Figure 5.3. Fraction of leached lysostaphin activity for aliquots collected at
different time intervals compared against total enzyme-NP activity for samples
with highest lysostaphin and antibody coatings. Black squares – NP: lysostaphin
(1:5400:0) and red circles - NP: lysostaphin: antibody (1:5400:1800); n=3
5.3.3. Bacterial lysis assay – Enzyme activity
Enzyme activity was measured as reduction in turbidity of S. aureus suspension in
PBS buffer by monitoring the decrease in optical density at 600 nm. Lysostaphin
concentration was 0.83 µg/ml in all experiments. Enzyme concentration was thus fixed in
all kinetic experiments; however the number of NPs (and, where applicable, the antibody
concentration) was different because of different number of lysostaphin molecules
adsorbed per NP for different ratios.
92
Sample
Final
Lysostaphin
concentration
( µg/ml)
Final
antibody
concentr
ation in
assay
(µg/ml)
Kinetic
constant of
bacterial lysis
(n=6; 3
replicates from
2 different
batches)
NP:lyso (1:1300)
0.83
-
NP:lyso (1:2700)
0.83
-
0.0136 ± 1E-03
p < 0.05
NP:lyso (1:5400)
0.83
-
0.0147 ± 2E-03
p < 0.05
NP:lyso:Ab (1:1300:450)
0.83
1.27
0.0088 ± 1E-03
p > 0.05
p > 0.05
NP:lyso:Ab (1:2700:900)
0.83
0.68
0.0175 ± 6E-03
p < 0.05
p > 0.05
NP:lyso:Ab (1:5400:1800)
0.83
0.6
0.032 ± 7E-03
p < 0.05
p < 0.05
Free lysostaphin
0.83
-
0.009 ± 1E-03
-
0.007 ± 2E-03
Significance
compared
against free
lysostaphin;
n=6;
paired ttest)
Significance
compared
against
enzyme:NP
analogue
lacking the
antibody; n
= 6;
paired t-test
p < 0.05
Table 5.3 - Bacterial turbdimetric assay results for all the samples.
Figure 5.4 below shows results of the activity assay for two batches of enzyme-NP
conjugates and free enzyme (n=6). Note: The high error bars in NP-enzyme samples are
also indicative of interference of NPs during absorbance reading at 600 nm. Assuming
first order kinetics with respect to bacteria, the time dependencies were fitted
exponentially and the mean kinetic constants of the first order reaction were calculated
from the exponential fits and used as the measure of antimicrobial activity. Table 5.3
shows results of the analysis of kinetic data obtained from two different batches (n=6; 3
replicates from each batch). Comparison of the activities for two sets of the samples with
the lysostaphin loadings (1:1300:0 and 1:1300:450; 1:2700:0 and 1:2700:900) showed no
significant difference (p>0.05, paired t-test; n=6, see Table 5.3), despite a higher mean rate
of bacterial lysis for antibody coated samples. Activity of the sample with the lowest
93
lysostaphin loading that lacked the antibody (1:1300:0) was found to be significantly
lower than that of the free enzyme (p<0.05, paired t-test; n=6, see Table 5.3). For its
corresponding analogue that contained the co-adsorbed antibody (1:1300:450), activity
was not found to be significantly different from that of the free enzyme (p>0.05, paired ttest; n=6, see Table 5.3).
However, enhanced activity compared to the free enzyme was observed for both
enzyme-coated and enzyme-antibody coated nanoparticles for all other ratios (p<0.05,
paired t-tests; n=6, Figs. 5.4B and 5.4C, Table 5.3). The rate of bacterial lysis for the
highest antibody coating was (corresponding to an initial bilayer of molecules to NP) was
the highest amongst all samples and significantly different compared to its corresponding
analogue without antibody (Table 5.3).
It had a nearly fourfold faster mean rate of
bacterial lysis compared to free enzyme at the same concentration. These experiments
demonstrate the possibility of antibody-directed targeting for samples with the highest
antibody coating concentration. We hypothesize that co-immobilization of lysostaphin and
antibody on nanoparticles leads to enhanced binding to bacteria through antibody-antigen
interactions resulting in higher local concentrations of lysostaphin and therefore leading to
increased bacterial lysis rates. However, enhanced potency can only be achieved at higher
antibody and lysostaphin loadings warranting further studies of the mechanism of the
observed phenomenon. Results of this model study suggest the possibility of regulating
properties of enzyme-NP conjugates by optimizing the ratio between the functional
enzyme and the targeting antibody attached to a nanoparticle. Targeting can be enhanced
by increasing the number of antibody molecules per NP (Figure 5.5B), even though total
antibody concentration in the assay decreases for samples with higher lysostaphin and
94
antibody loading (Table 5.3). Overall, bacterial lysis rates increased with the increased
number of antibody molecules per NP (Figure 5.5B). This increase was found significant
for all samples with different antibody: NP ratios (p<0.05, One way ANOVA).
An unexpected result was the observation of enhanced activity of lysostaphincoated nanoparticles compared to the free enzyme. In spite of the fact that lysostaphin
concentration in the assay was kept constant, bacterial lysis rates increased with the
increased number of lysostaphin molecules per NP (Figure 5.5A). This increase was found
to be statistically significant for all samples with different enzyme:NP ratios (p<0.05, One
way ANOVA). For two samples with higher lysostaphin loading (1:2700:0 and 1:5400:0),
activity was significantly higher than that of the free enzyme (p<0.05, paired t-test, n=6),
while for sample with the lowest lysostaphin loading activity was significantly lower than
that of the free enzyme (p<0.05, n=6).
95
A
B
C
Figure 5.4. Results of bacterial lysis assay. The data are the means of
absorbance for six replicates from two batches along with the standard deviations
for each time point. Blue triangles correspond to untreated bacterial suspension
(negative control). Black squares – lysostaphin adsorbed on PLA nanoparticles;
red circles – lysostaphin and antibody co-adsorbed on PLA nanoparticles, and
green triangles – free lysostaphin. (A) – NPs with low lysostaphin and antibody
coatings (1:1300:0 and 1:1300:450); (B) – NPs with intermediate lysostaphin and
antibody coatings (1:2700:0 and 1:2700:900); and (C) – NPs with high lysostaphin
and antibody coatings (1:5400:0 and 1:5400:1800). High error bars in NP-enzyme
samples are also indicative of interference of NPs during absorbance reading at
600 nm.
96
In general, partial loss of enzymatic activity is expected for surface-immobilized
enzymes including enzyme-nanoparticle conjugates [263, 277]. In our case, decrease of
enzymatic activity was only observed for the conjugates with the lowest lysostaphin
loading, while enhanced activity was observed for the other ratios. We have previously
observed enhanced enzymatic activity for lysozyme conjugated to positively-charged
polystyrene latex NPs. This effect was attributed to charge-directed targeting to negatively
charged bacteria (M. Lysodeikticus).
We considered the possibility that enhanced activity of lysostaphin-coated NPs
observed in the present work was also due to charge-directed targeting since lysostaphin is
positively charged and its attachment to nanoparticles could have lead to the formation of
positively charged conjugates. However, all conjugates were found to have highly
negative zeta-potential (Table 5.2). Also of note is that charge-directed targeting reported
by our group [250] was only observed in an environment with low ionic strength (6 mM
phosphate buffer, no saline) and was not noticeable at physiologically relevant ionic
strength (PBS buffer, ~150 mM saline). On the contrary, lysostaphin-coated NPs prepared
in the present work were robust and showed enhanced activity compared to the free
enzyme in PBS buffer containing 150 mM saline. Our hypothesis is therefore that
enhanced activity of lysostaphin-coated NPs in the absence of co-immobilized antibody
occurs due to multiple-ligand targeting. The mature form of lysostaphin consists of two
domains, the endopeptidase domain responsible for its catalytic activity, and the Cterminal cell wall-targeting domain (CWT) [305]. Lysostaphin-NP conjugates studied here
carry from several hundred to several thousand lysostaphin molecules per nanoparticle
(Figure 5.5A). Therefore, each particle presents a large number of CWT domains available
97
for binding to bacteria, which can lead to considerably higher binding constant for
interaction between the NP conjugates and bacteria, compared to that for interaction
between free lysostaphin and bacteria.
A
B
98
Figure 5.5. (A) Kinetic constant of bacterial lysis plotted as function of adsorbed
lysostaphin molecules per NP for lysostaphin:NP conjugates lacking the antibody
(mean±SD; n=6). (B) Kinetic constant of bacterial lysis plotted as function of
adsorbed antibody molecules per NP (mean±SD; n=6). Note that total antibody
concentration in the assay decreases with the increase of the number of antibody
molecules per NP (see Table 5.3).
Interestingly, enzyme and enzyme/antibody- coated nanoparticles remain
negatively charged. This finding can have important implications because toxicity of
many antimicrobial peptides to eukaryotic cells can potentially be reduced by their
conjugation to negatively charged carriers. High toxicity of antimicrobial peptides remains
one of the most important concerns in their in vivo applications [306]. Use of negatively
charged conjugates is expected to reduce non-specific attachment of antimicrobial
peptides to slightly negatively charged surface of eukaryotic cells therefore reducing their
toxicity.
In conclusion, we demonstrated that lysostaphin-coated nanoparticles show
enhanced antimicrobial activity against S. aureus at physiologically relevant conditions.
We also showed that co-adsorption of anti-S. aureus antibody to lysostaphin-coated
nanoparticles leads to further increase of their antimicrobial activity. Results of this study
indicate that enhanced antimicrobial activity of enzyme-nanoparticle conjugates is due to
targeted delivery to bacteria.
99
CHAPTER 6
EVALUATION OF ANTIMICROBIAL ACTIVITY OF LYSOSTAPHIN-COATED
HERNIA REPAIR MESHES
6.1. Introduction
The development of an incisional hernia is a common complication after
abdominal surgery which results in 90, 000 ventral hernia repair surgeries per year in the
United States [307]. The implantation of a prosthetic mesh is a well-established procedure
for reconstructing or reinforcing the abdominal wall and has been shown to decrease the
rate of hernia recurrence [308-310]. However, one of the most common problems
associated with the use of prosthetic meshes is bacterial infection. The incidence rate for
mesh-related infection has been reported to vary between 1% and 18% in different clinical
studies [311-313].
Mesh infection is associated with significant morbidity and is increased
considerably in patients with diabetes, immunosuppression, and obesity [314]. Infection
usually occurs in the early postoperative period and while its incidence can be decreased
by the use of antibiotics [315], no significant improvement has been made in the reduction
of mesh infection for more than 20 years. Current conventional therapy has proven to be
largely unsuccessful due to implantation of a foreign body, wide tissue dissection and the
development of a number of multi-drug resistant strains. In a study of mesh-related
infections following incisional herniorrhaphy, 63% of the microorganisms isolated were
Methicillin-resistant Staphylococcus aureus (MRSA)[313].
Bacterial adhesion and proliferation to implanted biomaterial surfaces is a key step
in the pathogenesis of infection [316-318]. Multiple experimental and clinical studies have
100
been focusing on the use of different antimicrobial agents immobilized on the surface or
released from the bulk of an implant [319]. Increasingly, attention is being paid to surface
coatings on implants that contain active compounds that may either kill directly on contact
or via leaching over time. Naturally occurring antimicrobial peptides and enzymes have
recently attracted much attention because of their high activity, broad antimicrobial
spectrum, and the low rate of antimicrobial resistance.
Balaban et.al found that adsorption of an antibacterial peptide, dermaseptin to
Dacron vascular grafts resulted in increased resistance to bacterial colonization and
demonstrated its in vivo efficacy in a rat model [239]. Shah et.al demonstrated significant
antimicrobial activity against S. aureus for intravenous catheters coated by physically
adsorbed lysostaphin, and observed high efficacy in vitro [240]. Lysostaphin possesses
extremely high activity against a variety of staphylococcal strains including MRSA [240].
Based on literature data lysostaphin has a reported minimum inhibitory concentration at
which 90% of the strains are inhibited [MIC90] of 0.001 to 0.064 µg/ml [320]. This is
comparatively much lower than MIC90 for most broad spectrum antibiotics and drugs that
are antimicrobial against S. aureus. For comparison, MIC90 for vancomycin, which is
currently recommended for treatment of methicillin-resistant S. aureus infections, is 2
µg/ml [321]. Lysostaphin could therefore be superior to other antibacterial agents in hernia
repair-related infections given that up to 90% of mesh-related infections result from S.
aureus. Here, we evaluate the in vitro antimicrobial susceptibility of S. aureus to
lysostaphin-coated hernia repair polymer meshes and study the effect of enzyme coating
concentration on antimicrobial activity
101
6.2. Materials and methods
Staphylococcus aureus (ATCC 32370) was purchased from American Type Culture
Collection
(Manassas,
VA).
Lysostaphin
(25
kDa;
LSPN-50)
isolated
from
Staphylococcus staphylolyticus was bought from Ambi Products LLC (Lawerence, NY).
Bovine Serum Albumin (BSA) was bought from Calbiochem (San Diego, CA). Ultrapro
(Lightweight Polypropylene mesh with Monocryl weave) from Ethicon Inc, Johnson and
Johnson (Langhorne, PA) and PariatexTM Composite mesh (Polyester mesh with
resorbable film) from Covidien (Mansfield, MA) was donated by Carolinas Medical
Center (Charlotte, NC). Alexa Fluor 594 Carboxylic acid, Succinimidyl ester dyes used
for fluorescent labeling were purchased from Invitrogen, Molecular Probes (Carlsbad,
CA). Microsep Centrifugal Devices with a modified Polyethersulfone membrane (low
protein binding; MWCO - 3K) used for enzyme purification were purchased from Pall
Life Sciences (Ann Arbor, MI). All other reagents were purchased from Fisher Scientific
(Pittsburgh, PA), and used without further purification unless otherwise specified. Type 1
Clear Borosilicate glass vial (25 mL) was with attached screw caps was bought from
VWR (West Chester, PA).
6.2.1 Fluorescent labeling of lysostaphin
Alexa Fluor 594 dye was used for fluorescent labeling of lysostaphin due to its enhanced
resistance to photobleaching [298]. Fluorescent labeling is considered to be an acceptable
method for monitoring protein concentration, which is assumed to be proportional to the
fluorescent signal [322]. Labeling of lysostaphin by Alexa Fluor 594 was performed
according to the manufacturer‟s instructions. Approximately 5 μg of Alexa Fluor 594 (ε
102
73,000) was dissolved in 500 μL of dimethylformamide and 100 μL of this solution was
added to 2 mL of a 1 mg/mL solution of lysostaphin in 10mM PBS buffer (10 mM
phosphate; 140 mM NaCl, 3 mM KCl). The enzyme was incubated with dye at 30°C for 2
h, then diluted two-fold in 10 mM PBS buffer and centrifuged at 3,500 g for 1 h using a
Microsep membrane centrifugal device (MWCO 3K) to remove any unbound dye
followed by resuspension of the concentrated labeled enzyme in PBS buffer.
6.2.2. Measurement of the specific surface area of the mesh samples
The specific surface area determined for one of the pieces by Brunauer-Emmett-Teller
(BET) analysis of N2 adsorption at 77 K in a Micromeritics ASAP 2020 (Micromeritics
Instrument Corporation, Norcross, GA) was found to be 5.5 m2/g. Prior to N2 adsorption,
the mesh samples were degassed overnight at 80 °C. Surface area for each of the mesh
pieces used in the following experiments was calculated based on the weight of the piece.
The average weight of a 1 × 1 cm2 Ultrapro mesh piece was 6.9±1.0 mg.
6.2.3. Preparation of lysostaphin-coated meshes
Ultarpro, Ethicon Inc., mesh (Lightweight Polypropylene mesh with Monocryl weave)
was cut into ~ 1 × 1 cm pieces in a laminar flow hood under sterile conditions prior to
physical adsorption of enzyme. Initial enzyme concentrations of 0.01, 0.025, 0.05, 0.1,
0.25, and 0.5 mg/ml in PBS buffer were prepared from a 1 mg/mL stock solution of Alexa
Fluor 594-labeled lysostaphin. The initial fluorescence intensities of the enzyme sample
solutions (1 mL) were measured in a 12 well plate (Nunc Inc.) using a microplate reader
(Ex 594 nm; Em 625 nm) and then added to 25 mL sterile glass vials. Using a pair of
sterile tweezers, mesh pieces were gently placed into each of the vials containing the
103
enzyme solutions and incubated overnight at room temperature with gentle shaking (100
rpm). The enzyme solution over the mesh was then collected and stored for fluorescence
measurements, and the mesh was gently washed 2 times with 1 mL of PBS buffer. The
wash solution was also collected and used in determination of enzyme binding yield. In
order to remove any loosely adsorbed enzyme, 1 mL of 0.1 (v/v %) Tween 20 solution
(non ionic surfactant) was then added to each of the glass vials followed by incubation for
3 h. This surfactant solution was also collected and used in the determination of the
amount of desorbed enzyme and the mesh samples were then washed with copious amount
of PBS buffer.
The concentration of unbound enzyme in each of the supernatants and wash
solutions was determined from fluorescence measurements. Initial enzyme solutions with
known concentrations were used as the standards. The concentration of the
unbound/desorbed enzyme at each step was then calculated and subtracted from initial
concentration of enzyme present in the initial solution. The difference corresponded to the
bound enzyme concentration on the mesh. Adsorption experiments were reproduced in
triplicates. Note: Unlabeled lysostaphin was used in all other experiments except binding
efficiency studies.
6.2.4. Leaching studies
Leaching of non-specifically bound lysostaphin from the surface can occur both in
vitro and in vivo. To model in vivo conditions, we studied enzyme leaching from
lysostaphin-coated meshes in the presence of BSA. The samples were incubated with 1
mL of 2 % (w/v) BSA solution in PBS buffer for 24 h at 37C. We found that fluorescence
measurements lacked sensitivity to determine concentration of leached enzyme in the
104
samples. We therefore estimated enzyme concentration based on its activity. For kinetic
measurements, 0.1 ml aliquots from each of the samples were taken at different time
points (5, 15 and 24 h) and added to a 96 well plate. Lysostaphin solutions with known
enzyme concentrations of 0.0122 – 25 µg/ml were used as standards for each of the
different time points. A 100 µl cell suspension of S. aureus in PBS buffer with optical
density (OD600) of 0.3 at 600 nm (1 cm light path) was then added to each of the different
wells and the rate of bacterial lysis was monitored continuously for 4 h at 37ºC.
Enzymatic activity for standards and unknown samples was measured simultaneously in
the same microplate. The initial rate of the reaction for each of the standards and
unknowns was calculated from the linearized slopes. The standard curve was obtained by
plotting the initial rate against the enzyme concentration. From the standard curve, the
unknown enzyme concentrations were determined and the amount of leached enzyme for
each of the samples was plotted as a function of time. All kinetic experiments were
reproduced in triplicates.
6.2.5. Turbidimetric assay
A cell suspension of S. aureus was prepared by inoculating 10 mL of 30% (w/v) tryptic
soy with 100 µL of S. aureus culture prepared according to ATCC instructions. The
procedure used was similar to the one employed by Schindler and Schuhardt [299]. The
cells were incubated at 37C for 18 - 24 h under gentle shaking to grow to a mid-log
phase. The cells were then pelleted at 3,000 g for 10 min and washed twice with 10 mM
PBS. The cells were then resuspended in PBS to prepare bacterial suspension with optical
density (OD600) of 0.55 at 600 nm (1 cm light path). 1 mL of the suspension containing
~107 CFU (colony forming units) was added to 25 mL glass vials containing the mesh
105
samples. The samples were incubated with the suspension at 37ºC under continuous
shaking, and the rate of bacterial lysis was monitored for 5 h by taking 0.2 mL aliquots
and measuring the OD600 in a 96 well plate at different time intervals. Enzyme activity
was expressed in units (one unit of lysostaphin activity is defined as a decrease of 0.01
absorbance units at 600 nm, according to the manufacturer‟s specifications). The vials
containing the cell suspension were continuously shaken in order to maintain contact with
the meshes. All kinetic experiments were reproduced in triplicates.
6.2.6. Fractional analysis of enzyme activity in leached samples
To determine whether antibacterial activity of the coated meshes comes primarily from
immobilized or leached enzyme, we performed fractional analysis of enzyme activity. A
set of mesh samples with the same initial coating concentrations was prepared in
duplicates using the procedure as described previously. All the samples were incubated in
25 ml glass vials containing 0.5 mL of 2 % (w/v) BSA solution in PBS buffer for 24 h at
37C. From one set of the samples, the meshes were removed at different time intervals
(1, 4 and 24 h) and a corresponding set containing mesh samples of identical coating
concentrations were left intact.
A cell suspension of S. aureus was grown to a mid-log phase similar to the
procedure described previously, centrifuged at 3,000 g for 10 min, washed twice with 10
mM PBS and resuspended in PBS to a final optical density (OD 600) of ~0.9 at 600 nm. 0.5
ml of this cell suspension was added to the 25 ml glass vials containing each set of the
samples and the final OD of this mixture measured was 0.55 at 600 nm. The samples were
incubated with the cell suspension at 37ºC under continuous shaking, and the rate of
bacterial lysis was monitored for 2 h by taking 0.2 mL aliquots from the reaction mixture
106
and measuring the OD600 in a 96 well plate at different time intervals. This procedure was
done for identical sets of the samples at the different time intervals to compare the enzyme
activity of the leached sample against total activity of the mesh sample incubated in BSA.
6.2.7. Colony counting
The sample meshes were placed in sterile glass vials and challenged with an inoculum of 1
ml of S. aureus suspension in tryptic soy broth containing 6 x 107 CFU for 24 h at 37 ºC.
After the incubation, the mesh pieces were retrieved and the broth supernatant was
collected for subsequent analysis. Each mesh piece was then washed once vigorously with
1 mL of PBS buffer using a pipette followed by vortex washing for 5 minutes and
collected for subsequent colony counting analysis. Serial dilutions and spot plating were
then performed for the broth supernatant (broth count) and the pooled wash (wash count)
on agar nutrient plates. The wash count was used to dislodge and quantify the loosely
attached bacteria. Bacterial counts were quantified as the number of CFU per mL. All
counts were performed in triplicates and results were calculated as mean log reduction
±SD.
6.2.8. In vivo Rat Model
All in vivo studies were performed by our collaborators, Dr. Yuliya Yurko, Dr. Amy
Lincourt, and Dr. Todd Heniford at Carolinas Medical Center along with Dr. John Shipp
at ViMedrx. A total of 40 male Lewis rats (225-275g, Charles River Laboratories,
Raleigh, NC) were used. All animal experiments were approved by the Institutional
Animal Care and Use Committee (CMC) and performed in accordance with NIH
guidelines. Rats were randomly assigned to one of the ten groups (n=4 each group) as
107
follows. Groups A-F had polypropylene mesh implanted as an onlay to the anterior
abdominal wall fascia. Group A received mesh with no lysostaphin and no S. aureus
inoculum. Group B received no lysostaphin and S. aureus inoculum of 5x105 CFU. Group
C received mesh-bound lysostaphin (Initial coating concentration - 50 µg/ml) and no S.
aureus inoculum
Group D received mesh-bound lysostaphin (Initial coating
concentration – 50 µg/ml) and S. aureus inoculum of 5x105 CFU. Group E received
mesh-bound lysostaphin (Initial coating concentration – 100 µg/ml) and no S. aureus
inoculum. Group F received mesh-bound lysostaphin (Initial coating concentration – 100
µg/ml) and S. aureus inoculum of 5x105 CFU. Groups G-J had polyester mesh implanted
as an onlay to the anterior abdominal wall fascia. Group G received mesh with no
Lysostaphin and no S. aureus inoculum. Group H received no Lysostaphin and S. aureus
inoculum of 5x105 CFU. Group I received mesh-bound Lysostaphin (Initial coating
concentration – 100 µg/ml) and no S. aureus inoculum. Group J received mesh-bound
Lysostaphin (Initial coating concentration – 100 µg/ml) and S. aureus inoculum of 5x105
CFU.
6.2.9. Surgical Procedures
Surgical anesthesia was induced and maintained with inhaled isofluorane. The abdominal
wall was shaved, prepared first with Betadine®, then isopropyl alcohol 70% (v/v), and
draped in sterile fashion. A 1 cm midline vertical incision was made through dermis and
subcutaneous pockets was then created bilaterally over the anterior abdominal fascia. A
3cm x 3cm square piece of sterile polypropylene mesh was placed on the fascia and
secured with eight stitches of 4-0 Prolene suture (Ethicon Inc.). For groups assigned to
receive the bacterial innoculum, a 1 cc suspension of 5 x 10 5 CFU S. aureus was applied
108
directly on the mesh using a sterile syringe. The incision was closed using vicryl suture
and reinforced with skin staples. Topical Bitter Orange (ARC Laboratories, Atlanta, GA)
was applied over the closure to dissuade wound disruption.
All animals received
bupranorphine (0.03mg/kg) immediately after surgery and every 12 hours thereafter for
the next 48 hours.
Mesh explantation was performed on day 7 after implantation. General anesthesia
was induced with isofluorane and the animals were euthanased by intracardiac
pentobarbitol injection. The skin around the original incision was prepared in the same
fashion as above and opened sharply and widely to allow full exposure of the graft
implant. The entire abdominal wall including the mesh interface was excised in en-block
fashion. For quantitative bacterial culture analysis the mesh was carefully separated from
the muscle, agitated and washed five times with sterile phosphate buffer solution. This
washings were serially diluted up to the final concentration of 10-8 and each dilution was
plated on tryptic soy agar and allowed to incubate overnight at 37º C. Bacterial colonies
were then counted.
6.2.10. Statistical analysis
Data are presented as means±SD. One-way analysis of variance (ANOVA) was performed
for colony counts from the broth supernatant and wash to compare the samples with
different initial enzyme concentrations. The null hypothesis was that the means are not
significantly different. Tukey‟s test was used to compare different samples in the event the
ANOVA null hypothesis was not true. Data were analyzed using Origin 8 Data Analysis
and Graphing Software (OriginLab Corporation, MA, USA). A p value of <0.05 was
considered statistically significant.
109
6.3. Results
6.3.1. Binding yield of lysostaphin on Ultrapro
Binding yield was calculated based on the amount of fluorescently labeled enzyme
adsorbed on mesh and corresponded to the difference in fluorescence intensities of initial
enzyme and the supernatant solutions as described above. Figure 6.1 shows the binding
yield for Alexa Fluor-labeled lysostaphin on a 1×1 cm Ultrapro mesh. The amount of
adsorbed enzyme per unit BET surface area (µg/cm2) is plotted as a function of the initial
enzyme concentration. Based on the above described experimental conditions, 18 to 40%
of lysostaphin initially present in the solution remained bound to the mesh after treatment
by Tween 20 solution.
0.40
2
Bound (ug/cm )
0.35
0.30
0.25
0.20
0.15
0.10
0.05
0.00
0
100
200
300
400
500
600
Initial lysostaphin concentration (ug/ml)
Figure 6.1. Binding yield for Alexa 594 labeled lysostaphin on Ultrapro (1 × 1 cm)
after overnight adsorption at room temperature (n = 3).
110
The degree of surface coverage on the mesh can be predicted from the binding
yield of enzyme. Average dimensions of the lysostaphin molecule are 4.5 nm X 5.5 nm X
8.5 nm. The average surface area occupied by an adsorbed protein molecule based on the
smallest dimensions is 25.02 nm2. The maximum theoretical monolayer of lysostaphin
adsorbed on a 1×1 cm piece of Ultrapro mesh calculated using the above surface area per
molecule and the BET specific surface area of the mesh (3.8 x 10 16 nm2) consisted of
1.5x1015 molecules. We observed approaching a theoretical 50%, 100%, and 150%
monolayer coating of adsorbed lysotaphin at initial enzyme concentrations of 100, 250,
and 500 g/mL, respectively. Typically, the amount of adsorbed enzyme on a surface
increases sharply at low initial enzyme concentration, and reaches a plateau at higher
concentrations, thereby approaching a certain limiting value for adsorption [323]. In our
case, adsorption did not reach saturation even at the highest coating concentration of 0.5
mg/mL suggesting the possibility of multilayer adsorption at higher concentrations.
6.3.2. Lysostaphin leaching
Enzyme leaching could be a potential problem in the case of non-covalent
adsorption as the functional protein may be replaced on the surface by more abundant
nonfunctional ones in vivo. Also, the Ultrapro mesh is constructed of a monofilament
lightweight large porous polypropylene with pores larger than 3 mm which could result in
initial confinement of the enzyme followed by release in vivo [324]. In our study, we
assessed lysostaphin leaching in the presence of 2 % (w/v) BSA as a model for the most
abundant protein in the abdominal fluid. Figure 6.2 shows the standard curve representing
the rate of lysis for the standards with known enzyme concentrations. From this standard
111
curve, the unknown enzyme concentrations were determined and the amount of leached
enzyme for each of the samples was plotted as a function of time.
Figure 6.2. Standard curve plotting the mean rate of lysis (OD per min) against
the different known lysostaphin concentrations (µg/ml) (n=3).
The samples designated based on their initial lysostaphin concentrations of
10µg/ml, 25µg/ml, 50 µg/ml, 100µg/ml, 250µg/ml and 500µg/ml were L-10, L-25, L-50,
L-100, L-250 and L-500 respectively. We observed an overall (0.01- 3) % leaching of
enzyme based on the initial coating concentration from all the studied samples after 24 h
incubation in BSA. Bacterial lysis was observed in all mesh samples including those that
leached very low amounts of lysostaphin at the different time points, suggesting that small
concentrations of enzyme could be effective in inhibiting growth of cells. In vitro
antimicrobial activity against S. aureus was evaluated for all of the above samples (L-10
through L-500) using turbidity assay and colony counting method. Figure 6.3 below
shows leaching for all different coating concentrations after 24 h.
112
Figure 6.3. Leaching of adsorbed lysostaphin as a function of time for mesh
samples treated by different initial enzyme concentrations. Black squares 10µg/ml, Red circles -25µg/ml, Green triangles - 50µg/ml, Blue inverted triangles 100µg/ml, Maroon diamonds - 250µg/ml and Pink arrows - 500µg/ml. Leaching
studies were performed in the presence of 2 % (w/v) BSA in PBS buffer (pH = 7.4)
at 37C.
6.3.3.Turbidimetric antibacterial activity assay
Activity assay provides the most direct measure of an enzyme‟s functionality. Qualitative
evaluation of lysostaphin activity was done by monitoring the cell lysis of a S. aureus cell
suspension [325]. The degree of cell lysis is directly proportional to the decrease in OD 600
of the bacterial suspension.
113
Figure 6.4. Turbidimetric enzyme activity assay for the different lysostaphincoated hernia mesh samples. Black squares - 10µg/ml, Red circles -100µg/ml,
Blue triangles - 500µg/ml and Green inverted triangles - Unocoated mesh; (n=3);
p < 0.05 (one way ANOVA; n=3).
We monitored the decrease in optical density of the bacterial suspension and compared the
rate of lysis for the samples with different enzyme coatings. Figure 6.4 shows the time
course of bacterial degradation for three of the samples along with the appropriate
controls. The vials containing the cell suspension were continuously shaken in order to
maintain constant contact with the meshes.
114
Sample
Initial coating
concentration
(µg/ml)
Enzyme activity
(Units/ml) 1
(mean±SD)
L - 10
10
2.97 ± 0.8
L - 25
25
3.08 ± 0.8
L - 50
50
3.78 ± 1.1
L - 100
100
4.02 ± 0.01
L - 250
250
5.92 ± 0.3
L - 500
500
6.98 ± 0.3
P - 100
100
6.98 ± 0.3
Table 6.1 - Specific enzyme activity of samples with different enzyme coating concentrations
6.3.4. Leached enzyme activity assay
It is important to know whether activity of coated meshes is due to immobilized or leached
enzyme. A turbidity assay that compared the rates of leached sample against that of the
total activity of enzyme-coated mesh at different time points (incubated in bovine serum
albumin) was studied to calculate the fraction of activity contributed by lysostaphin
leaching from the mesh surface into the cell suspension. Enzyme leaching during the first
5 h (which was the duration of the turbidity assay) never exceeded 1%. However,
extremely small concentrations of lysostaphin leaching from the mesh can cause
1
Enzyme activity was expressed in units (one unit of lysostaphin activity is defined as a decrease of 0.01
absorbance units at 600 nm, according to the manufacturer‟s specifications).
115
significant lysis to its natural substrate, i.e. cell walls of S. aureus. Fractional analysis of
enzyme activity was calculated for the aliquots collected at different time intervals and
compared against total mesh activity for different coating concentrations for 24 h in 2 %
BSA. Figure 6.5 below shows the results for all different coating concentrations.
Figure 6.5. Fraction of leached sample activity for the aliquots collected at
different time intervals compared against total mesh activity for different coating
concentrations for 24 h in 2 % BSA. Black squares – 10 µg/ml, Red circles -50
µg/ml, Green triangles – 50 µg/ml, Blue inverted triangles – 100 µg/ml, Light blue
diamonds - 250µg/ml and Pink arrows – 500 µg/ml
It was found that lytic activity for these samples is almost entirely due to enzyme
leaching from the mesh and lysing bacteria in the surrounding supernatant rather than
adsorbed lysostaphin (data not shown). Interestingly, Shah et al. observed that the
antimicrobial activity of lysostaphin coated on two different plastic surfaces, polystyrene
(well plates) and FEP (fluorinated ethylene-propylene) polymer, a Teflon-like material
used in Angiocath catheters was primarily due to immobilized but not leached enzyme
116
[240]. These authors used a methodology similar to that reported here. One possible
explanation is difference in materials used in our and Shah‟s studies. Ultrapro, a
lightweight polypropylene macroporous mesh (with a monocryl weave) containing an
absorbable component of poliglecaprone is different in density, weave, porosity (3 mm)
and relative hydrophobicity. It has more „yarn-like‟ properties with its higher specific
surface area compared to „flat‟ surfaces such as polystyrene well plates and the lumenal
sides of catheters made up of FEP polymer. Therefore, larger amounts of lysostaphin can
be adsorbed to and consequently leached from the Ultrapro mesh leading to increased
fraction of activity coming from the leached enzyme.
6.3.5. Colony counting
Log reduction in bacteria is estimated as logarithm of ratio of initial bacterial colonies to
average final surviving colonies after treatment. This has been used for both broth
supernatant and wash counts. Table 6.2 shows the log reduction in colonies recovered
from the broth supernatant and wash counts for all the different initial coating
concentrations. Broth count is a model for activity against the bacteria present in the
wound fluid, while the wash count models activity against colonized bacteria present on
the surface of the mesh, as initial bacterial attachment plays an important role in infection.
An averaged 1.6 ± 0.4 log reduction in bacterial counts was observed in the case of broth
supernatant recovered from all the lysostaphin-coated mesh samples after 24 h incubation
with bacteria.
117
Sample
L - 10
L - 25
Initial coating
concentration
(µg/ml)
Log10 reduction in
Colonies for Broth
Supernatant
Log10 reduction in
Colonies for Wash
Count
(Mean ± SD)
(Mean ± SD)
1.2 ± 0.2
1.5 ± 0.4
1.3 ± 0.2
2 ± 0.3
10
25
L - 50
50
1.8 ± 0.8
2 ± 0.4
L - 100
100
1.8 ± 0.6
1.6 ± 0.2
L - 250
250
2.3 ± 0.5
2 ± 0.3
L - 500
500
1.6 ± 0.4
1.6 ± 0.2
P - 100 -
100
1.3 ± 0.6
1.5 ± 0.3
(Polyester)
Table 6.2 - Effect of lysostaphin coating concentration on log reduction of colonies for the
broth supernatant and wash count; (n =3).
There was no significant difference between samples with different initial
lysostaphin coating concentrations (p > 0.05; one way ANOVA); however, observed log
reduction was significantly higher for the all lysostaphin-coated samples when compared
against uncoated mesh samples (p<0.05; Tukey‟s test). Similarly, an average 1.8 ± 0.36
log reduction in bacterial counts was observed from the pooled wash count for all the
samples that were not significantly different from each other (p>0.05; one way ANOVA).
In contrast, uncoated mesh samples showed hardly any antimicrobial activity and the
bacteria thrived in their presence as indicated by a comparable surviving bacterial count
with plain control cells. Bacterial adherence to the mesh is a known precursor to prosthetic
118
infection [326, 327]. A significantly less number of CFU were recovered from the wash
counts for lysostaphin-coated samples compared to the uncoated samples (p < 0.05). It is
thus evident from the observed data that lysostaphin-coated meshes are able to both
efficiently kill bacteria in solution and possibly inhibit surface colonization of bacteria.
6.3.6. In vivo rat model trial
All twenty rats survived to the time point of mesh extraction at 7 days. Groups that did
not have S. aureus inoculum (Groups A, C, E, G, I) had sterile cultures at the time of
extraction. None of lysostaphin treated controls had impaired healing. Two of four rats
with polyester and no lysostaphin and one of four with polypropylene and no lysostaphin
had wound complications. All of the rats receiving a S. aureus inoculum and no
lysostaphin had positive mesh cultures at seven days. Animals treated with L-50 and
inoculated with 5 x 105 CFU of S. aureus had positive mesh cultures. The L-100 group
with 5 x 105 CFU S. aureus inoculum (Groups F and J) had negative cultures (Table 6.3).
119
Group
Mesh Type
Lysostaphin
Dose
A
B
C
D
E
F
G
H
I
J
Polypropylene
Polypropylene
Polypropylene
Polypropylene
Polypropylene
Polypropylene
Polyester
Polyester
Polyester
Polyester
None
None
L-50
L-50
L-100
L-100
None
None
L-100
L-100
S. aureus
Inoculum
(CFU)
Time at
Extraction
None
5 x 105
None
5 x 105
None
5 x 105
None
5 x 105
None
5 x 105
7 days
7 days
7 days
7 days
7 days
7 days
7 days
7 days
7 days
7 days
Positive
Cultures
0/4
4/4
0/4
4/4
0/4
0/4
0/4
4/4
0/4
0/4
Table 3: In vivo rat study results
6.4. Discussion
Bacterial infection at the site of implanted medical devices presents a serious and
ongoing problem in the biomedical arena. Mesh implantation has reduced the rate of
recurrent hernia but has led to an increased rate of bacterial infections. The incidence is
reported to be as high as 8% following repair of incisonal hernias [311, 314, 328].
Although mesh-related infections occur relatively infrequently compared to other devicerelated infections, it can cause significant morbidity resulting in non-wound healing,
recurrent hernia, and need for reoperation and mesh excision. Considering that nearly one
million inguinal and incisional hernia repairs are done every year, this is a real and
significant medical issue.
In our current study, we evaluate the in vitro antimicrobial susceptibility of S.
aureus to lysostaphin-coated hernia repair meshes and study the effect of different enzyme
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coating concentrations on antimicrobial activity. Lysostaphin is a glycylglycine
endopeptidase isolated from Staphylococcus staphylolyticus which specifically cleaves
pentaglycine cross bridges found in the staphylococcal peptidoglycan [329]. It kills S.
aureus within minutes (MIC at which 90% of the strains are inhibited [MIC90], 0.001 to
0.064 µg/ml) [320]. We observed that the antimicrobial activity of the meshes occurs due
to extremely low concentrations of lysostaphin leaching from the surface into the
supernatant. These concentrations are higher than its minimum inhibitory concentration
(MIC) and enzyme release from the mesh into surrounding tissue or fluids can be expected
to counter any initial elevated infection risk immediately post-surgery or implantation.
A 6-h post implantation “decisive period” has been identified during which
prevention of bacterial colonization mediated by adhesion is critical to the long term
success of an implant [330]. Concentrations of lysostaphin that leached from the mesh in
the first 5 h incubation in BSA varied from 0.0026 to 0.5 µg/ml depending upon the initial
coating concentration. This is followed by a sustained release of enzyme at the same
concentration that could be effective against occurrence of a latent infection. Passive
coatings that aim to reduce bacterial adhesion are not as effective as “active” coatings that
are designed to allow release of antibacterial agents immediately following the
implantation [330]. The antimicrobial activity of the lysostaphin-coated meshes suggests
that such enzyme releasing surfaces could be efficient at actively resisting bacterial
adhesion and preventing subsequent colonization of implant.
From the colony counting data, the antimicrobial activity was not significantly
concentration dependent in the range of L-25 to L- 500 (25 to 500µg/ml) as all the six
samples reduced bacterial titers to the same level. This agrees well with the previously
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reported data [240]. They found that lysostaphin adsorbed onto two different plastic
surfaces i.e. polystyrene and FEP polymer showed killing that was not concentration
dependent where all coating concentrations (0.1, 1 and 10 mg/ml) reduced the bacterial
count to the same level. The overall 1.6 (± 0.421) log reduction in the broth count versus
1.8 (± 0.36) log reduction in the wash count for the coated meshes emphasized the slightly
higher activity against adherent bacteria on the mesh surface, possibly due to a higher
local concentration of lysostaphin near the mesh surface. Bacterial adhesion after 24 h of
incubation in a 107 CFU/ml of S. aureus suspension is greatly reduced by enzyme-coated
meshes compared to uncoated sample.
The results of the in vivo study show that none of the controls had bacterial
contamination at 7 days. Treatment with L-50 was insufficient to clear all of bacterial
innoculum, as evidenced by positive cultures in all rats in Group D. Interestingly, when
doubling the treatment concentration to L-100, despite the presence of a foreign body, the
entire bacterial load was cleared. Although the colony count data in the in vitro studies
did not demonstrate significant difference in bacteriocidal activity of lysostaphin at
different concentrations, the antibacterial efficacy of lysostaphin appears to be doserelated in vivo. The dramatic antimicrobial activity in L-100 treated group demonstrates
great potential for lysostaphin in the clinical setting for preventing Staphylococcal related
prosthesis infections.
Based on our in vivo findings, we decided to characterize and test the P-100 Polyester
mesh (PariatexTM) sample with the same set of in vitro experiments as the Ultrapro
Polypropylene mesh.
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Our preliminary binding yield study showed a twofold decrease in the total amount
of adsorbed lysostaphin per mg of a 1 x 1 cm piece of Polyester mesh compared to its
Polypropylene counterpart with the same dimensions (data not shown).
The P-100
Polyester mesh sample showed significantly higher leaching (~ 4%) in 24 h compared to
the L-100 Ultrapro sample which showed negligible leaching. This was further confirmed
with an activity assay of the P-100 Polyester sample that showed an almost twofold higher
enzyme activity (7 ± 0.3 units per ml) calculated from the initial rate of the reactions from
linearized slopes compared to the L-100 Polypropylene sample (Table 1). This can be
attributed to the difference in rigidity, composition and hydrophobicity between the two
mesh types. PariatexTM is a composite mesh made up of multifilament Polyester mesh and
absorbable hydrophilic film made up of Collagen, Polyethylene Glycol and Glycerol while
Ultrapro is a lightweight polypropylene mesh (with a monocryl weave) containing an
absorbable component of poliglecaprone with pores between 0.1 and 3 mm [331].
Hydrophilicity and low surface tension due to Polyethylene Glycol in the film could result
in minimizing and weakening enzyme adsorption to Polyester meshes. We observed an
average of 1.3 ± 0.6 log reduction for broth supernatant and 1. 46 ± 0.3 log reduction in
the bacterial wash count for the P-100 Polyester sample after 24 h incubation. The L-100
Polypropylene sample also had a similar (1.56 ± 0.2 log reduction) in the wash count
despite higher amount of lysostaphin adsorbed to the mesh surface (twofold higher) and
negligible leaching of enzyme in vitro.
In conclusion we have demonstrated the antimicrobial activity of lysostaphincoated hernia repair mesh against S. aureus at different coating concentrations. Our early
in vivo trial did not show impaired wound healing in the lysostaphin treated groups;
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furthermore these data demonstrate that lysostaphin has significant promise in preventing
prosthetic infection when used at dose of 100 µg/ml. Utilizing such an antibiotic-free
approach, one can control the release of enzyme locally, in the area of implant infection,
by using an appropriate coating concentration that would result in lower systemic toxicity
and higher antibacterial efficacy. Further work is on the way to confirm our findings in the
clinical arena. Surface-coatings using antibacterial enzymes could be a ground breaking
addition to the field of hernia repair and other areas of surgery in which prosthetic
materials, or even biologic materials, are implanted.
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CHAPTER 7
APPLICATION OF ENZYMES AS ANTIBACTERIAL COATINGS ON PLAIN
AND NANOPARTICLE COATED CATHETER SEGMENTS
7.1. Introduction
Medical devices such as intravascular and urinary catheters are routinely employed
in healthcare sector for hemodialysis, drainage of urine, administration of IV fluids,
medication, oxygen, anesthetic agents and monitoring of blood and intracranial
pressure[332, 333]. Unfortunately, these catheters are also a major cause of nosocomial
infections that occur in more than two million hospitalizations in the US annually, with
major medical remediation costs Consequently, catheter-related infections are notoriously
difficult to treat via conventional antibiotic therapy, with associated mortality rates
ranging from 12% to 25% [334]. Extended hospital stays therefore calls for an active
intervention on the part of the healthcare personnel, and has driven the estimated annual
healthcare cost arising from these catheter-related biofilm infections to more than nine
billion dollars [334].
Catheters are generally infected by microorganisms that are typically present in the
surrounding site of insertion, usually as part of natural flora in the skin; contaminate the
catheter along the outer surface [335]. They can also be contaminated in their lumenal
compartments through fluid flow from contaminated infusate or from parts of the implant
that are improperly sterilized [335]. Sometimes, percutaneous devices such as catheters
sometimes are „marsupialized‟, where basal cells from the epidermis near the edge of the
wound may migrate down the edge of the dermis to surround the implant forming a pouch
without any surrounding tissue integration [336]. The space between the implant and the
125
epidermis then becomes filled with cell debris which then is a permanent source for onset
of infection because of the lack of a natural mechanism for rejuvenation or cleaning [336].
In addition, upon insertion in the host, the surface of the catheter becomes „passivated‟ by
a layer of blood proteins (due to contact with device) that assists attachment of planktonic
bacteria. Eventually, the final stage in bacterial colonization leads to the formation of
biofilms on the surface of the device [3, 6, 41, 231, 232]. Various clinical studies have
been focusing on the use of different antibiotics immobilized on the surface or released
from the bulk of an implant for prevention of biofilm formation [337, 338]. However,
treatment with conventional antibiotics frequently fails because bacteria develop multidrug resistance and bacteria growing in biofilms are more resistant than planktonic cells.
Staphylococcus aureus is a Gram-positive, biofilm-forming bacterium that is regarded as a
leading cause of nosocomial infections and accounting (together with Staphylococcus
epidermidis) for more than one-half of prosthetic device-associated infections[339].
Thus, there is an urgent need for development of alternative treatment strategies that
could overcome the problems associated with the use of conventional antibiotic therapy.
Increasingly, attention has been paid specifically to enzymes and peptides as surface
coatings to prevent device-associated infections [106, 109, 240, 340, 341]. For a catheter
surface, the coating may be retained by either physical adsorption or covalent attachment
of the enzyme. The use of nanoparticles as enzyme supports since the 80s has thereafter
sustained a continued interest for its application modern biotechnology [187-190]. Higher
surface area for enzyme loading, better stability and long term durability are some of the
expected outcomes of using nanoparticles as coatings on catheter segments for enzyme
immobilization. Hence we propose to use poly (lactic acid) nanoparticles as „surrogate‟
126
surface coatings for enzyme adsorption on indwelling medical devices such as Central
venous catheters (CVCs). In this preliminary study, we propose to coat DispersinB and
lysostaphin to the surface of Polyurethane catheter segments at different coating
concentrations through single adsorption and coadsorption (two enzymes), characterize
and evaluate the in vitro performance of plain and nanoparticle coated catheter segments.
7.2. Materials and methods
Materials: Staphylococcus aureus (ATCC 27660) was purchased from American Type
Culture Collection (Manassas, VA). Lysostaphin (25 kDa; LSPN-50) isolated from
Staphylococcus staphylolyticus was bought from Ambi Products LLC (Lawerence, NY).
DispersinB (41 kDa) produced by Aggregatibacter actinomycetemcomitans was bought
from Kane Biotech Inc. (Manitoba, Canada). Bovine Serum Albumin (BSA) was bought
from Calbiochem (San Diego, CA). Polyurethane Catheter tubing (Single lumen) was
purchased A.P. Extrusion (Salem, NH). Antibiotic–antimycotic solution, heat inactivated
fetal bovine serum, Dulbecco‟s modified eagle medium and Alexa Fluor 594 Carboxylic
acid, Succinimidyl ester dyes used for fluorescent labeling were purchased from
Invitrogen, Molecular Probes (Carlsbad, CA). PBS Tablets (#524650) were purchased
from Cabiochem (La Jolla, CA). BCA and micro BCA reagent kits were purchased from
Pierce Biotech Inc (Rockford, IL). BBLTM Trypticase Soy Broth was purchased from
Becto, Dickinson and Company (Sparks, MD). Methylene Blue was obtained from the
Department of Biological Sciences at Clemson University courtesy of John Abercombie.
Poly (DL-lactide; average Mw (97500), Pluronic F68 (MW = 8.4 kDa) and 4-nitrophenylN-acetyl- D-glucosaminide (substrate for DispersinB) was purchased from Sigma Aldrich
127
(St. Lous, MO). CellTiter 96® AQueous One Solution Cell Proliferation Assay (MTS) for
cell viability studies was purchased from Promega Corporation (Madison, WI). Type 1
Clear Borosilicate glass vial (8 mL) was with attached screw caps was bought from VWR
(West Chester, PA). A microsep Centrifugal Devices with a modified Polyethersulfone
membrane (low protein binding; MWCO - 3K) used for enzyme purification were
purchased from Pall Life Sciences (Ann Arbor, MI). All other reagents were purchased
from VWR (West Chester, PA) and Fisher Scientific (Pittsburgh, PA), and used without
further purification unless otherwise specified. Binding efficiency, Turbidimetric activity
and Colorimetric assays were performed using a Synergy microplate reader (Bio-Tek
Instruments, Inc., Winooski, VT). Particle sizing and Zeta potential analysis of
nanoparticles (NPs) was done using a 90plus Particle Size Analyzer (Brookhaven
Instruments Co, Holtsville, NY). Scanning electron microscopy was done at the Electron
microscopy facility at the Advanced Materials Research Laboratory (Pendleton, SC).
7.2.1. Preparation of nanoparticles
Polylactide nanoparticles were prepared using a nanoprecipitation method based on
solvent diffusion. Briefly, Poly(DL-lactide) (average Mw (97500) was dissolved in 30 ml
of acetone at a concentration of 8 mg/ml and this solution was then dispersed in an
aqueous phase (200 ml) containing Pluronic F68 (5 mg/ml) under sonication ((Bransonic
5510, Branson, MO) for 30 min. The acetone solution was added dropwise at a uniform
rate into the aqueous phase. The resulting suspension containing the nanoparticles was
then purified three times in DI water by ultracentrifugation (Allegra™ 64R Centrifuge,
Beckman Coulter, USA) at 3000g for 1 hr at 25°C to remove any excess polymer and
Pluronic. Nanoparticle suspensions in DI water were stored in refrigerator at 4°C. The
128
number of nanoparticles in the final suspension was then determined using gravimetric
analysis.
7.2.2. Particle sizing
Stock solutions containing nanoparticles were diluted 1:1000 in DI water in a disposable
cell and particle size was measured using dynamic light scattering with a 90Plus particle
size /zeta potential analyzer (BrookHaven Instruments Corporation, Holtsville, NY).
Before particle sizing, the samples were sonicated for 30 min to break up any loosely-held
agglomerates. Particle size was expressed as a combined mean diameter in nm (+/- SD).
7.2.3. Preparation of NP coated catheter segments
Single lumen Polyurethane catheter tubing (Outer diameter (0.125 inches) and Inner
diameter (0.63 inches) purchased from A.P.Extrusion was used for all the adsorption
studies. 1 cm segments were cut off from the main tubing in a laminar flow hood under
sterile conditions and autoclaved at 121 °C. Using a pair of sterile tweezers, the catheter
segments were gently placed in 8 ml glass vials containing 0.5 ml of the concentrated
nanoparticle solution and incubated overnight at room temperature with gentle shaking (50
rpm). Control catheter segments were placed in glass vials containing 0.5 ml of DI water.
The samples were then washed gently with DI water three times and air dried at RT for 6
h before protein adsorption experiments. The average weight of a 1 cm catheter segment
was 77.5 mg calculated from combined weight of 6 catheter segments weighed together.
7.2.4. Surface characterization of catheter segments with scanning electron
microscopy
129
NP coated catheter and uncoated samples were cut longitudinally to 0.2 cm segments and
gently adhered horizontally to aluminum specimen support stubs using double-sided
carbon tape. Specimens were then sputter-coated with Platinum and examined using a SU
6600 field emission SEM equipped with variable pressure. The instrument was operated at
10 kV accelerating voltage with a 10-12 mm working distance and the images were
captured digitally.
7.2.5. Enzyme adsorption on plain and NP coated catheter segments
Initial enzyme concentrations of 0.025, 0.05, 0.1, 0.25, and 0.5 mg/ml in 1mM PBS buffer
were each prepared from a 1 mg/mL stock solution for lysostaphin and DispersinB
respectively. Using a pair of sterile tweezers, catheter segments (plain and NP coated)
were gently placed into each of the vials containing 0.5 ml of enzyme solutions and
incubated overnight at room temperature with gentle shaking (100 rpm). The sample
solutions were removed and the catheter segments were gently flushed with 5 ml of PBS
buffer five times and air dried for 1 h. The enzyme solution above the catheter segment
was then collected and stored for binding efficiency measurements. The concentration of
unbound enzyme in the supernatants was determined using BCA assay. Initial enzyme
solutions with known concentrations were used as the standards. The concentration of the
unbound enzyme was calculated and subtracted from initial concentration of enzyme
present in the initial solution. The difference corresponded to the bound enzyme
concentration on the catheter segment. Adsorption experiments were reproduced in
triplicates.
7.2.6. Enzyme coadsorption on plain and NP coated catheter segments
130
For coadsorption studies, we used fluorescently labeled lysostaphin (AlexaFluor 594) of enzyme
concentrations - 0.025, 0.05, 0.1, 0.25, and 0.5 mg/ml in 1mM PBS buffer and correspondingly
mixed with 0.025, 0.05, 0.1, 0.25, and 0.5 mg/ml of DispersinB to make up the final volume of 0.5
ml. Alexa Fluor 594 dye was used for fluorescent labeling of lysostaphin as described in
previous sections [298] . Catheter segments (plain and NP coated) were placed into each of the
vials containing 0.5 ml of enzyme mixture solutions and incubated overnight at room temperature
with gentle shaking (100 rpm). The sample solutions were removed and the catheter segments
were gently flushed with 5 ml of PBS buffer five times and air dried for 1 h. The enzyme solution
above the catheter segment was similarly collected and stored for binding efficiency
measurements. The concentration of total unbound enzyme in the supernatants was determined
using BCA assay. Initial enzyme mixture solutions with known concentrations were used as the
standards. The concentration of unbound enzyme was calculated and subtracted from initial
concentration of total enzyme present in the initial mixture solution. The difference corresponded
to the total bound enzyme concentration on the catheter segment for both lysostaphin and
DispersinB.
The concentration of unbound lysostaphin in the supernatant was determined from
fluorescence measurements. Fluorescence intensities of the supernatant and standards (100
µl) were measured in a 96 well plate using a microplate reader (Ex 594 nm; Em 625 nm).
The concentration of unbound lysostaphin was calculated and subtracted from initial
concentration of enzyme present in the initial solution. The difference corresponded to the
bound lysostaphin concentration on the catheter segment. The difference of total bound
concentration of both enzymes (BCA assay) and bound lysostaphin (fluorescence)
corresponded to the bound concentration of DispersinB on the catheter segment.
Coadsorption experiments were also reproduced in triplicates.
131
7.2.7. Activity assay for adsorbed enzymes
Lysostaphin: Turbidity assay was used to assess the activity of lysostaphin adsorbed to
Polyurethane segments. A cell suspension of S. aureus was prepared by inoculating be 10
mL of 30% (w/v) tryptic soy with 100 µL of S. aureus culture prepared according to
ATCC instructions as described previously. The cells were incubated at 37C for 18 - 24 h
under gentle shaking to grow to a mid-log phase. The cells were centrifuged at 2,000 g for
10 min and washed twice with 10 mM PBS. The cells were then resuspended in PBS to
prepare a bacterial suspension with optical density (OD600) of ~ 0.4 at 600 nm (1 cm light
path). 0.5 mL of the bacterial suspension was be added to 8 mL glass vials containing the
catheter segments. The samples were be incubated with the suspension at 37ºC under
continuous shaking, and the rate of bacterial lysis was be monitored for 1 h by taking 0.1
mL aliquots and measuring the OD600 in a 96 well plate. The vials containing the cell
suspension will be continuously shaken in order to maintain contact with the catheter
segments. All kinetic experiments were reproduced in duplicates.
DispersinB: Activity of adsorbed DispersinB was measured using a colorimetric assay
[107]. DispersinB activity was quantitatively assessed using UV-visible (UV-VIS)
spectroscopy by measuring the absorbance at 400 nm of the p-nitrophenolate reaction
product resulting from enzymatic hydrolysis of the substrate 4-nitrophenyl-N-acetyl- Dglucosaminide. 0.5 ml of 5 mM substrate in 50mM Sodium Phosphate buffer was added to
the vials containing the catheter segments and the enzymatic reaction was carried out for 1
h at 37°C. After 1 h, the reaction was quenched by adding 50 µl of 10 N NaOH and 100 µl
aliquots were collected from each sample and the absorbance was measured at 400 nm.
All experiments were reproduced in duplicates.
132
7.2.8. Enzyme leaching studies
Lysostaphin: Leaching of non-specifically bound enzymes from the catheter surface can
occur both in vitro and in vivo. To model in vivo conditions, we studied enzyme leaching
from enzyme coated catheter segments in the presence of BSA. The samples were
incubated with 0.5 mL of 2 % (w/v) BSA solution in PBS buffer for 72 h at 37C. We
found that fluorescence measurements lacked sensitivity to determine concentration of
leached enzyme in the samples. We therefore estimated enzyme concentration based on its
activity. For kinetic measurements, 50 µl aliquots from each of the samples were taken at
different time points (6, 24, 48 and 72 h) and added to a 96 well plate.
A 125 µl cell suspension of S. aureus in PBS buffer with optical density (OD600) of
0.4 at 600 nm (1 cm light path) was then added to each of the different wells and the rate
of bacterial lysis was monitored continuously for 2 h at 37ºC. Lysostaphin solutions with
known enzyme concentrations of 0.0071 – 7.14 µg/ml were used as standards for the
assay. Enzymatic activity for standards and unknown samples was measured
simultaneously in the same microplate. The initial rate of the reaction for each of the
standards and unknowns was calculated from the linearized slopes. The standard curve
was obtained by plotting the initial rate against the enzyme concentration. From the
standard curve, the unknown enzyme concentrations were determined and the amount of
leached enzyme for each of the samples was plotted as a function of time. All kinetic
experiments were reproduced in duplicates.
DispersinB:
Similarly, leaching of DispersinB was studied modeling in vivo conditions, in the
presence of 2 % (w/v) BSA solution in PBS buffer for 72 h at 37C. For kinetic
133
measurements, 50 µl aliquots from each of the samples were taken at different time points
(6, 24, 48 and 72 h) and added to a 96 well plate. A 100 µl solution of 4-nitrophenyl-Nacetyl- D-glucosaminide (5 mM) was added to each of the different wells and the
absorbance at 400 nm of the p-nitrophenolate reaction product resulting from enzymatic
hydrolysis of the substrate was monitored continuously for 1 h at 37ºC.
DispersinB solutions with known enzyme concentrations of 0.083 – 8.3 µg/ml
were used as standards for the assay. Enzymatic activity for standards and unknown
samples was measured simultaneously in the same microplate. The initial rate of the
reaction for each of the standards and unknowns was calculated from the slopes. The
standard curve was obtained by plotting the initial rate against the enzyme concentration.
From the standard curve, the unknown enzyme concentrations were determined and the
amount of leached enzyme for each of the samples was plotted as a function of time. All
kinetic experiments were reproduced in duplicates.
The different samples designated based on their initial coating concentrations of
10µg/ml, 25µg/ml, 50 µg/ml, 100µg/ml, 250µg/ml and 500µg/ml were L-25, L-50, L-100,
L-250, L-500 for lysostaphin and D-25, D-50, D-100, D-250 and D-500 for DispersinB
coated on uncoated catheter segments. Similarly, samples were designated as NPL-25,
NPL-50, NPL-100, NPL-250, NPL-500 and NPD-25, NPD-50, NPD-100, NPD-250,
NPD-500 for lysostaphin and DispersinB adsorbed on NP coated catheter segments
respectively.
7.2.9. Cell viability using MTS assay
The CellTiter 96® AQueous Assay uses the novel tetrazolium compound (3-(4,5dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium,
134
inner salt; MTS) and the electron coupling reagent, phenazine methosulfate (PMS). MTS
is chemically reduced by cells into formazan, which is soluble in tissue culture medium
[342]. The measurement of the absorbance of the formazan can be carried out using 96
well microplates at 492nm. The assay measures dehydrogenase enzyme activity found in
metabolically active cells. Since the production of formazan is proportional to the number
of living cells, the intensity of the produced color is a good indication of the viability of
the cells. MTS solutions were prepared according to the manufacturer‟s instructions.
Using the methods outlined in ISO standard 10993-5 as a guideline, mouse 3T3
fibroblasts (donated by Cassie Gregory; Department of Bioengineering, Clemson
University) were previously cultured in Cell culture flasks in DMEM supplemented with
1% Fetal Bovine Serum (FBS) and 1% antibiotic (Penicillin/Streptomycin). The cells were
trypsinized, centrifuged for 5 minutes at 1000 rpm with the medium aspirated and the cell
pellet resuspended in fresh media at a concentration ~10,000 cells/mL. Aliquots (100 µL)
were then pipetted into individual wells of a 24-well plate with fresh media. Cells were
allowed to grow to confluence for 24 h at 37°C in a humidified 5% CO 2 environment.
Cell viability was checked under a microscope and the media replaced every 12 h. The
samples were then cut into 0.3 cm pieces to ensure specimen coverage of approximately
one-tenth of the cell layer surface (ISO 10993-5). Samples were then gently placed on the
cell layer in the center of 24-well plates using a pair of sterile forceps. Subsequently,
plates were incubated for 48 h at 37°C in a humidified 5% CO 2 environment with media
being changed every 12 h. Care was taken during mesh placement in the wells to prevent
unnecessary movement, which can lead to physical trauma to the cells and dislodgement
135
of cells from the mesh. Sterile Latex pieces were used as positive controls for assessment
of cytotoxicity (ISO 10993-5) while cells in media alone were used as negative controls.
After 48 h, samples were gently removed using a pair of sterile forceps and 50 µl
of MTS reagent was added to every well in one of the 24-well plates in a ratio of 1:5
(MTS reagent : volume of media in the well). The plate was incubated at 37 °C for 4 h in
a humidified, 5% CO2 atmosphere. 100 µl from each well was then transferred to wells of
a 96-well plate and the absorbance was recorded at 490 nm (OD490) and 650 nm (OD650)
using a plate reader. Raw quantitative data generated by the microplate reader were
analyzed by first subtracting the difference of OD490 and OD650 reading to remove any
background interference. The resultant mean OD readings from each well were calculated
as an average of two replicates for each sample and expressed as a percentage of the OD
values observed relative to OD values measured in the negative control wells (negative
control – cells in media).
7.2.10 Antibiofilm assay
Briefly, S. aureus culture was grown overnight in 10 ml of TSB (Bidifico) supplemented
with 1% D-glucose (Tokyo Kasei, Japan). The overnight bacterial cultures was then
centrifuged twice, resuspended in TSB and spot plating on Agar was done to determine
the number of colonies in the active culture. 1 ml containing ~ 10 6 CFUs in TSB
supplemented with1% D-glucose was added to wells of a 12-well tissue culture plate
(Costar Inc., Corning, N.Y.) containing the coated catheter segments. The 12-well plate
was incubated at 37°C with shaking at 100 rpm for 12 h and then removed from the shaker
and supplemented with either 200 µl of additional fresh TSB to compensate for medium
evaporation. These plates were then be incubated at 37°C for an additional 24 h.
136
After ~36 h of growth, the media was removed from various wells, washed thrice
with phosphate-buffered saline (PBS) to remove any non-adherent bacteria. The assay
plates were allowed to dry at 37°C for approximately 2 hr. The wells containing the
segments were then stained with 0.5 ml of 1% Methylene Blue for 30 min and then gently
washed with PBS three times; air dried for 1 h and destained with 30% acetic acid
solution. The eluted dye solution was then transferred to wells of a 96 well plate and the
absorbance was recorded at 660 nm to evaluate the inhibition of S. aureus biofilms.
7.2.11. Accelerated Shelf life Study
Accelerated aging testing is performed on packaged medical devices such as
catheters and implants to ascertain its shelf life and document expiration dates. We
modeled a study based on long term stability testing of our enzyme coated catheter
segments by measuring enzyme activity after storage. Initial lysostaphin and DispersinB
concentration of 0.5 mg/ml in 1mM PBS buffer was each prepared from a 2 mg/mL stock
solution and using a pair of sterile tweezers, catheter segments (plain) were gently placed
into vials containing 0.5 ml of enzyme solutions and incubated overnight at room
temperature with gentle shaking (100 rpm). The sample solutions were removed and the
catheter segments were gently flushed with 5 ml of PBS buffer five times and air dried for
1 h. The samples were then incubated in a -80 °C freezer overnight in glass vials sealed
with parafilm. Samples were then freeze-dried for 24 h in a Freeze Dry System/Freezone
4.5 from Labconco (Kansas City, MO) courtesy of Dr. Ken Webb (Clemson University).
Samples were then hermetically sealed using a heat press in special peelable high barrier
pouches (# TPC-1475B) from Oliver-Tolas Healthcare Packaging (Grand Rapids, MI).
137
The samples were then incubated at 60 °C for 1 week in an Isotemp Oven; Model 516G
from Fisher Scientific (Ann Arbor, MI).
After 1 week, the samples were removed and placed in wells of a 24 well plate.
For lysostaphin activity, 0.5 mL of a previously purified bacterial suspension with optical
density (OD600) of ~ 0.38 at 600 nm (1 cm light path) was added to each of the wells
containing the catheter segments. The samples were incubated with the suspension at 37ºC
under continuous shaking along with appropriate controls, and the rate of bacterial lysis
was be monitored for 3 h by taking 0.1 mL aliquots every hour and measuring the OD 600
in a 96 well plate. The vials containing the cell suspension were continuously shaken in
order to maintain contact with the catheter segments. Activity of DispersinB adsorbed on
catheter segments was monitored using the colorimetric assay as described previously by
measuring the absorbance at 400 nm of the p-nitrophenolate reaction product resulting
from enzymatic hydrolysis of the substrate 4-nitrophenyl-N-acetyl-D-glucosaminide. 0.5
ml of 5 mM substrate in 50mM Sodium Phosphate buffer was added to the vials
containing the catheter segments and the enzymatic reaction was carried out for 1 h at
37°C. After 1 h, the reaction was quenched by adding 50 µl of 10 N NaOH and 100 µl
aliquots were collected from each sample and the absorbance was measured at 400 nm.
All experiments were reproduced in triplicates.
7.2.12. Statistical analysis
Data are presented as means±SD. One-way analysis of variance (ANOVA) and Paired„t‟
test was performed for comparing coated and uncoated samples with different initial
enzyme concentrations. The null hypothesis was that the means are not significantly
different. Data were analyzed using Origin 6.0 and 8.0 Data Analysis and Graphing
138
Software (OriginLab Corporation, MA, USA). A p value of <0.05 was considered
statistically significant.
7.3. Results and Discussion
7.3.1. Enzyme Binding yield
Based on previous preliminary data, which demonstrated high antibacterial activity
of enzyme-NP conjugates against planktonic cells (Chapters 4 and 5), we proposed to
compare the loading of enzymes and enzyme-NP conjugates on catheter segments and
subsequently evaluate antimicrobial and antibiofilm activity of the catheter segments. We
decided to adsorb both DispersinB and lysostaphin to the surface of PLA nanoparticles
either alone or through coadsorption. After synthesis and characterization of enzyme-NP
conjugates, we incubated them with catheter segments and compared the binding
efficiency of plain enzyme- and enzyme-NP conjugate-coated catheter segments.
Higher binding yield, better enzyme stability and long term durability were some
of the expected outcomes using this approach. However, we observed two- to five-fold
less amount of DispersinB adsorbed in the case of enzyme-NP conjugate-coated catheter
segment compared to the plain enzyme-coated catheter segment and thirty-fold less
amount of DispersinB adsorbed in case of samples coadsorbed with lysostaphin (data not
shown).
We hypothesized that coating by enzymes makes NPs more hydrophilic leading to
decreased NP attachment to the catheters. Thus, we decided to modify our approach to
achieve better coating by enzyme-NP conjugates. We opted to first coat the surface of the
catheter segment with PLA nanoparticles and then adsorb enzymes at different initial
139
concentrations either alone or through coadsorption. This approach eliminates possible
error in determination of enzyme concentration in enzyme-NP conjugates as we are using
the same initial enzyme concentrations when coating catheters by either plain enzyme or
enzyme-NP conjugates.
We made certain theoretical approximations. A greater surface area is expected
upon coating PLA nanoparticles on the surface of a Polyurethane catheter segment. The
average diameter of a PLA NP calculated from previous DLS experiments was 181 ± 29
nm (mean from 3 different batches) depending upon the batch of NPs used for coating
experiments. Based on the dimensions of a 1 cm PU catheter segment, with ID and OD of
0.317 cm and 0.158 cm, the theoretical surface area calculated assuming a hollow cylinder
is 1.614 cm2. The number of nanoparticles that could saturate the surface of a 1 cm PU
catheter segment with a diameter of 180 nm is 1.5 x 109 particles assuming that the surface
of the catheter segment is smooth and devoid of any topography. The surface area
occupied by each spherical NP adsorbed on the surface is 4πr 2. Theoretically, an enzyme
molecule that has an average hydrodynamic radius of „r‟ occupying the surface of the
catheter segment will have a surface area of r 2. However, if it were to adsorb on the
surface of a catheter segment that is completely saturated with a monolayer of NPs, it
could theoretically occupy a surface area of 4 πr 2 per NP on the catheter (Figure 7.1). This
increase in surface area and high surface energy of a nanoparticle could contribute to
better adsorption and subsequently higher effective loading based on the same initial
coating concentration of enzyme for NP coated catheter segments. Previously, Crisante et
al investigated the antibiotic release of nano-structured polymer systems in carboxylated
polyurethane [343]. They hypothesized that since nanoparticles are characterized by a high
140
surface/volume ratio; their presence in the polymer could result in a greater surface area
available for the antibiotic adsorption.
Figure 7.1. An illustration for a model single lumen catheter segment before and
after coating with nanoparticles.
Binding yield was calculated based on the total amount of enzyme adsorbed per
mg of plain or PLA NP coated PU catheter segment. Figure 7.2 shows the binding yield
for lysostaphin and DispersinB on plain or NP coated 1 cm PU catheter segment at
different initial enzyme coating concentrations. Figure 7.2A and 7.2B specifically
corresponds to enzyme adsorption for lysostaphin and DispersinB individually and Figure
7.2C and 7.2D correspond to adsorption of each enzyme in the presence of the other
(coadsorption). The amount of adsorbed enzyme (µg) per mg of catheter segment is
plotted as a function of the initial enzyme concentration.
141
A
B
C
D
Figure 7.2 A). Binding yield for lysostaphin on plain and NP coated PU catheter
segment; B). Binding yield for DispersinB on NP coated PU catheter segment C).
Binding yield for lysostaphin on plain and NP coated PU catheter segment when
coadsorbed with DispersinB at the same coating concentration. D). Binding yield
for DispersinB on plain and NP coated PU catheter segment when coadsorbed
with lysostaphin at the same coating concentration; (Mean ± SD; n = 3). Black
squares represent plain catheter segment while red circles represent NP coated
catheter segment
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Based on the above described experimental conditions, ~11 to 23% of lysostaphin
initially present in the solution remained bound to the catheter segment for plain catheter
samples and ~ 8 to 29% was bound to NP coated catheter segments depending upon the
initial coating concentration. Similarly, ~12 to 30% and ~ 20 to 35 % of DispersinB
initially present in the solution remained bound to the catheter segment for plain and NP
coated catheter segments respectively. Generally, the amount of enzyme bound to catheter
segment increased linearly on increasing the enzyme coating concentration for plain and
NP coated catheter segments (Figure 7.2A and 7.2B). Plain coated catheter segments
showed a slightly higher adsorbed lysostaphin concentration per weight than NP coated
catheter segments for the samples, while the amount of DispersinB bound to catheter
segment was not significantly different for both types of catheter segments (p > 0.05).
In case of the coadsorption study, both lysostaphin and DispersinB were
simultaneously adsorbed at different initial concentrations and the amount of each enzyme
was calculated using both BCA analysis and fluorescence. As described in the methods
section, the amount of bound lysostaphin was calculated via fluorescence labeling and the
difference between the total protein content (BCA analysis) and bound lysostaphin was
used to calculate amount of DispersinB bound on the catheter segment. Binding yields for
lysostaphin consisted of ~2 to 10% and ~8 to 13 % per total weight of the protein mixture
in the initial solution for plain and NP coated catheter segments respectively. Similarly,
the binding yield for DispersinB corresponded to ~0.1 to 68% and ~1.5 to 42% per total
protein weight for plain and NP coated catheter segments respectively when co-adsorbed
with the lysostaphin (Figure 7.2C and 7.2D).
143
We expected a higher surface area for enzyme adsorption for NP coated catheter
segments after precoating plain Polyurethane catheter segments with PLA nanoparticles.
BET specific surface analysis study comparing both plain and NP coated Polyurethane
segments was inconclusive for all sets of samples because specific surface area for all
samples was below BET detection limit (data not shown). Figure 7.3A-H below shows a
cross section of the outer surface of plain and NP coated Polyurethane catheter segments
taken at different magnifications taken on SU-6600 field emission SEM equipped with
variable pressure. These images clearly show that the sample catheter segments did not
have a monolayer of nanoparticle coating on the surface as expected. There was still a
large portion of the catheter segment that was bare and devoid of any nanoparticles.
Lysostaphin and DispersinB molecules probably bound more on the bare portion rather
than NP coated part of the polyurethane catheter segment just to due to higher available
surface area for enzyme adsorption.
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A
B
C
D
E
F
G
H
Figure 7.3. SEM images of plain (A, C, E, G) and PLA NP (B, D, F, H) coated
Polyurethane catheter segments at different magnifications taken on SU-6600
field emission SEM equipped with variable pressure..
145
The degree of surface coverage on the catheter segment can be predicted from the
binding yield of each either of the enzymes when adsorbed alone or in the presence of the
other. Average dimensions of the lysostaphin molecule are 4.5 nm X 5.5 nm X 8.5 nm
[301]. The average surface area occupied by an adsorbed protein molecule based on the
smallest dimensions is 25.02 nm2.
Similarly, the average dimensions of DispersinB
molecule are 4.1 nm X 8.6 X 18.1 nm [344]. The average surface area occupied by
adsorbed DispersinB molecule based on smallest dimensions is 35.26 nm2. The maximum
theoretical monolayer of lysostaphin adsorbed on a 1 cm catheter segment calculated
using the above surface area per molecule and the theoretical surface area of catheter
segment of 1.614 cm2 consisted of 6.5 X 1012 molecules. Similarly, for DispersinB, the
number of molecules that could occupy the surface of the catheter segment for a
theoretical monolayer is 4.6 X 1012. Based on these theoretical considerations, the binding
yield data for lysostaphin and DispersinB adsorbed alone or through coadsorption on plain
and NP coated catheter segments suggests multilayer protein adsorption, from ten to
several hundred layers of molecules, depending upon the coating concentration.
Multilayer adsorption on polymer surfaces has been observed with different
proteins interacting with a variety of surfaces [344-347]. The proteins in direct physical
contact with the adsorbing surface or the layer of water molecules present around the
surface will be strongly affected by surface properties [348, 349]. These surface properties
include wetability, topography, chemical composition etc. Due to these surface
characteristics, the interaction between a protein molecule and the surface can induce
conformational changes on the protein, resulting in a different surface that would
influence interaction with the surrounding protein molecules in the bulk solution and so
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on. The adsorption isotherms (Figure 7.2A and 7.2B) from a single protein solution do not
reach saturation upon increasing the protein concentration providing additional argument
in favor of multilayer adsorption hypothesis.
Another possible explanation is that adsorbed proteins actually form a monolayer
but real specific surface area of the catheter segment is significantly higher than that
estimated from purely geometric considerations. In fact, BET using Kr adsorption
performed in this study has a very low detection limit of 0.05 m2 (500 cm2) of the surface:
Thus, even if the catheter‟s actual specific surface area exceeds that calculated from
geometric dimensions by the factor of 200-300, it would still be undetectable using BET
method. Of note, alternative BET approach that uses N2 instead of Kr is even less sensitive
and requires 0.5-1 m2 of the surface at the lower detection limit.
Protein adsorption studies from solutions containing a mixture of proteins imply
some competitive adsorption, where different proteins with different characteristics will
compete with each other for the available surface area for adsorption [241]. Aside from
the size, isoelectric point, molecular weights, secondary structures of the protein etc
surfaces features also influences interactions between different proteins. We observed a
decrease in amount of each of individually adsorbed protein (lysostaphin and DispersinB)
when adsorption occurs from a mixture of protein solutions. At the same time, the total
amount of bound protein from a mixture of proteins is significantly higher (p < 0.05) than
total protein adsorbing from a single solution for both plain and NP coated catheter
segments.
7.3.2. Adsorbed enzyme activity assay
147
Activity of adsorbed lysostaphin on plain and NP coated catheter segments was
done by monitoring the cell lysis of a S. aureus cell suspension [325] as described
previously. We monitored the decrease in optical density of the bacterial suspension and
compared the rate of lysis for the samples with the highest and lowest initial enzyme
coating concentration (25µg/ml and 500µg/ml). Figure 7.4A shows the mean rate
constants of bacterial lysis and standard deviations for these samples (n=2) for both plain
and NP coated catheter samples.
*
Figure 7.4. Activity assay data shown as bar graphs comparing the kinetic
constant of bacterial lysis against the highest and lowest lysostaphin coating
concentrations for plain and NP coated catheter segments. Black bar – NPL-25
represnts NP coated catheter segment with initial lysostaphin concentration of 25
µg/ml, Red bar – NPL-500 represents NP coated catheter segment with initial
lysostaphin concentration 500 µg/ml; Green bar – L-25 represents plain catheter
segment with initial lysostaphin concentration of 25 µg/ml and Pink bar – L-500
148
represents plain catheter segment with initial lysostaphin concentration 500 µg/ml;
(Mean ± SD; n= 2; * - p > 0.05)
We found that the initial rate of cell wall lysis was not significantly concentration
dependent for the two sets of initial concentrations (p>0.05 –„Paired t test‟) and not
significantly different for all samples of plain and NP coated catheter segments (p>0.05 –
One way ANOVA). A similar trend in adsorbed lysostaphin activity was also seen in
samples coadsorbed with DispersinB at the same coating concentrations (data not shown).
It is well known that lysostaphin is active against S. aureus at extrememly low
concentrations [320]. The results obtained here are in strong agreement with data obtained
by Shah et al, who also evaluated lysostaphin activity after physical adsorption to catheter
segments [240]. They also found that the antibacterial activity of lysostaphin was not
dependent upon coating concentrations as they found all three concentrations used (0.1, 1
and 10 mg/ml) showed the same antistaphylococcal activity by inhibiting bacterial growth
to the same level. This is in contrast to our experiments with lysostaphin-coated hernia
repair meshes (Polypropylene weave – Ultrapro) where we observed a correlation between
lysostaphin coating concentration and lytic activity of all the samples. This can again be
attributed to differences in material composition and porosity of the two types of samples
as mentioned previously.
We used a colorimetric assay to test activity of DispersinB adsorbed to catheter
segments with a synthetic substrate, 4-nitrophenyl-N-acetyl-β-glucosaminide in a
procedure similar to the one used by Donelli et.al [107]. We monitored the optical density
(OD400) of p-nitrophenolate reaction product resulting from the enzymatic hydrolysis of
the substrate by comparing the rates of enzymatic hydrolysis for the samples with the
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highest and lowest initial enzyme coating concentration (25 µg/ml and 500 µg/ml). It is
well known that most biofilm-producing staphylococcal strains produce a linear PNAG
and it has been demonstrated previously that DispersinB is able to degrade Nacetylglucosamine-containing polysaccharides [107, 350, 351]. Figure 7.5 shows the mean
rate constants of hydrolysis and standard deviations for these samples (n=2) for both plain
and NP coated catheter samples. We found the rate of hydrolysis of substrate was
significantly concentration dependent for the two sets of initial concentrations (p<0.05 –
„Paired t test‟) for both plain and NP coated catheter segments. However, we did not see a
significant difference between plain and NP coated catheter segments (p > 0.05; one way
ANOVA). A similar trend in adsorbed DispersinB activity was also seen in samples
coadsorbed with lysostaphin at the same coating concentrations (data not shown).
*
¥
Figure 7.5. Activity assay data shown as bar graphs comparing the kinetic
constant of substrate hydrolysis via colorimetric assay against the highest and
lowest DispersinB coating concentrations for plain and NP coated catheter
segments. Black bar – NPD-25 represnts NP coated catheter segment with initial
150
DispersinB concentration of 25 µg/ml, Red bar – NPD-500 represents NP coated
catheter segment with initial DispersinB concentration 500 µg/ml; Green bar – D25 represents plain catheter segment with initial DispersinB concentration of 25
µg/ml and Pink bar – D-500 represents plain catheter segment with initial
DispersinB concentration 500 µg/ml; (Mean ± SD; n= 2; ¥ - p < 0.05; * - p > 0.05;)
7.3.3. Enzyme leaching results
We assessed lysostaphin and DispersinB leaching in the presence of 2 % (w/v)
BSA, as a model for the most abundant protein in blood plasma. Figure 7.6 shows
leaching for all samples with different coating concentrations for lysostaphin and
DispersinB adsorbed to plain and NP coated catheter segments. We chose to quantify
leaching using activity assays for the individual enzymes rather than BCA analysis or
fluorescent labeling owing to low sensitivity of the two latter techniques. The unknown
enzyme concentrations for each of the samples were determined from a standard curve
that plotted rate of the reaction (enzymatic lysis for lysostaphin and enzymatic hydrolysis
for DispersinB) against a series of known enzyme concentrations and the amount of
leached enzyme for each of the samples was plotted as a function of time.
A
B
151
D
C
Figure 7.6. Leaching of enzyme as a function of time for plain and NP coated
catheter segments treated by different initial enzyme concentrations. A.)
Lysostaphin leaching from plain catheter segment B). Lysostaphin leaching from
NP coated catheter segment C). DispersinB leaching from plain catheter segment
and D). DispersinB leaching from NP coated catheter segment. Black squares –
25 µg/ml, Red circles -50 µg/ml, Green triangles – 100 µg/ml, Blue inverted
triangles - 250µg/ml and Light Blue diamonds - 500µg/ml. Leaching studies were
152
performed in the presence of 2 % (w/v) BSA in PBS buffer (pH = 7.4) at 37C.
(Mean ± SD; n= 2).
We observed an overall < 0.2 % enzyme leaching from all the studied samples
after 72 h incubation in BSA (Figure 7.6 A-D). In case of coadsorption of lysostaphin and
DispersinB on plain and NP coated catheter segments, we observed an almost identical
trend of < 0.2% leaching of enzyme for all the samples (data not shown). Again, the data
obtained in our leaching study reflected a similar trend in the results obtained by authors at
Biosynexus Inc [240]. They observed almost negligible leaching after coating lysostaphin
on a Teflon-like FEP polymer used in angiocath catheter segments and Polystyrene well
plates.
7.3.4. Cell viability study
Of particular importance in assessing cellular response and biocompatibility is
quantifying the prospective biomaterial‟s inherent cytotoxicity, or its ability to induce cell
death [352]. In vitro models may provide answers regarding several aspects, including cell
viability (cytotoxicity of implant material or extractables), proinflammatory effects and
alteration of cellular response to different signaling molecules like cytokines [353].
Fibroblasts were chosen as an appropriate cell model as they play a crucial role in the
inflammatory response [354]. This study aimed to evaluate the cellular response to
enzyme coated catheter segments using an in vitro approach and methods in accordance
with the International Organization for Standardization‟s (ISO) standard number 10993-5.
In particular, it is important to evaluate the cytotoxic effects of the two adsorbed enzymes,
lysostaphin and DispersinB. Both enzymes are produced by different bacterial species.
Lysostaphin is isolated from culture filtrates of S. Staphylolyticus and DispersinB is
153
isolated from A. actinomycetemcomitans and could show potential cytotoxcity, especially
if small amounts of enzyme leached from the catheter surface during in vivo trials.
The results of cytotoxicity tests on extracts by MTS assay are shown in Fig.7.7.
Raw quantitative data generated by the microplate reader were analyzed by first
subtracting the difference of OD490 and OD650 reading to remove any background
interference. The resultant mean OD readings from each well were calculated as an
average of two replicates for each sample and expressed as a percentage of the OD values
observed relative to OD values measured in the negative control wells (negative control –
cells in media).
Figure 7.7. Direct cytotoxicity results of MTT assay that shows the percentage of
cell viability after 48 h of all the catheter samples coated with the highest coating
concentration that were placed in contact with 3T3 fibroblasts (n=2). Coating
concentration for all samples was 500µg/ml. Black Bar –
L-500, Red bar – D-
500, Yellow bar – LD-500, Wine Red – NPL-500, Grey Bar –NPD-500, Dark Blue
bar – NPLD-500, Pink Bar – Uncoated cathter segment, Green Bar – PLA NP
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coated catheter segment, Light Blue Bar – Negative control (Untreated
Fibroblasts) and Orange Bar - Positve control (Latex samples)
The metabolic activity of the fibroblasts is directly proportional to the cell
viability. Overall, the cell viability in the negative controls (Plain cells in media) did not
differ significantly from any of the test samples including uncoated or coated either by
lysostaphin coated or DispersinB or via coadsorption of two enzymes (p > 0.05; p =
0.31718, One way ANOVA). There was also no significant difference between coated
plain and NP coated catheter segments (p > 0.05; p = 0.14232, Paired‟t‟ test). Again, we
did not see a significant difference between plain and NP coated segments. The positive
control (Latex fragments) was shown expectedly to be highly cytotoxic showing ~ 14 %
cell viability post 48 h incubation with the cells. These results demonstrated that the
enzyme coated catheter segments were not cytotoxic in vitro in chosen experimental
conditions.
7.3.5. Antibiofilm assay
DispersinB combined with various antimicrobials has been shown to work
synergistically to inhibit the growth and proliferation of biofilm-embedded bacteria such
as Staphylococcus epidermis and Staphylococcus aureus [107, 109, 110]. Figure 7.8
shows the percentage of biomass for the samples coated with the highest enzyme
concentration – 0.5 mg/ml. We tested the in vitro antibiofilm activity of samples coated
with DispersinB or lysostaphin alone or as a combination for both plain and NP coated
catheter segments. We wanted to test synergy between lysostaphin (antibacterial) and
DispersinB (antibiofilm) and study if we could completely inhibit adhesion and growth for
samples coadsorbed with both enzymes.
155
Figure 7.8. Biofilm inhibition assay of catheter samples done by Methylene Blue
staining for catheter samples coated with highest enzyme concentration. Black
Bar – NPL-500, Red bar – NPD-500, Blue bar – NPLD-500, Green Bar – L-500,
Brown Bar –D-500, Light brown bar –LD-500 and Dark Blue Bar – Uncoated
catheter segment; (Mean ± SD ; n=3).
We chose methylene blue as our dye for biofilm staining over crystal violet
because in our preliminarily studies we observed significant non-specific binding of
crystal violet to Polyurethane catheter segments. Percentage of biomass on the catheter
segment was calculated from the following formula – ((Absorbance
(660)
of sample) /
(Absorbance (660) of control))*100
We observed an average of 26 ± 13 % biomass on NP coated catheter segments adsorbed
with lysostaphin compared to plain lysostaphin samples that showed an average of 13 ± 1
% biomass and this was not significantly different (p>0.05; Paired „t‟ test) compared to
samples coated with DispersinB for plain (12 ± 4)% and NP coated catheter samples (15 ±
4)% biomass. At coating concentrations of 0.5 mg/ml, it seems like both DispersinB and
156
lysostaphin inhibit biofilm formation on catheter surfaces and there was no significant
difference in amount of biomass on catheters even after coadsorption of the two enzymes
(p> 0.05; One way ANOVA).
Polysaccharide intercellular adhesin (PIA), composed of poly-β-1,6-linked-Nacetylglucosamine, a sticky extracellular polysaccharide, is the main constituent of S.
aureus
and S. epidermis biofilm matrix [40]. PIA is produced by the intercellular
adhesion (ica) operon present in nearly all S. aureus strains. DispersinB specifically
hydrolyses the glycosidic linkages of poly-β-1,6-N-acetylglucosamine without inhibiting
the growth of bacteria [108]. It is a stable glycoside hydrolase that functions in a narrow
pH range and effective against both Gram positive and Gram negative bacteria.
DispersinB can only inhibit or disperse Staphylococci biofilms without affecting their
growth, so it would be necessary to combine it with lysostaphin in order to inhibit the
growth and cause lysis of biofilm-embedded bacteria. Previously, authors at Kane Biotech
have shown synergy and in vitro and in vivo efficacy of DispersinB® and Triclosan
combination against S.aureus and S.epidermis biofilms [109].
However, lysostaphin by itself is known to have some biofilm inhibiting activity.
Kokai-Kun et al have previously shown inhibition of S. aureus biofilms by lysostaphin in
vitro on artificial surfaces and in vivo using a catheterized mouse model [341]. The
authors have also observed that lysostaphin not only killed the staphylococci in the
biofilm, but also appeared to prevent formation of the EPS matrix needed for biofilm
formation on the artificial surfaces. The exact mechanism by which lysosytaphin disrupts
biofilm formation remains unclear but the authors suggest that disruption occurs through
the rapid lysis of the sessile staphylococci, which may be sufficient to destabilize the
157
entire biofilm matrix in a manner that allows cell detachment from surfaces [162]. We also
similarly observed inhibition of S. aureus biofilm after coating lysostaphin on plain and
NP coated catheter segments. We hypothesize that the planktonic bacteria in the media
(during growth of biofilm) are directly interacting with the enzyme coating on our catheter
segment. Our preliminary binding yield study suggests the possibility of multiple layers of
enzyme coating on the catheter surface. These layers of lysosyaphin molecules are either
directly interacting with planktonic bacterial cells or lysostaphin molecules slowly leach
into the surrounding environment and lyse the bacteria thereby preventing subsequent
attachment and biofilm formation. The latter however seems unlikely since data from the
leaching study shows lysostaphin leaching (Fig 7.6) to be less than 1 % after 72 h of
incubation in BSA.
B
A
Figure 7.9.
Images taken on a digital camera courstey department of
Bioengineering; A). Control – Uncoated catheter PU segment. B). Lysostaphin
and DispersinB coated PU catheter segment.
7.3.6. Accelerated shelf life study
158
Biomedical devices and implants coated with antimicrobial enzymes in storage may
change as they age, but they are considered to be stable as long as they retain specific
activity as per manufacturer's specifications. The number of days that the product remains
stable at the recommended storage conditions is referred to as the shelf life. Shelf life is
commonly estimated using two types of stability testing: real-time stability tests and
accelerated stability tests [355-357] In the former, a product is stored at recommended
storage conditions and its shelf life is monitored while in the latter, a product is stored at
elevated stress conditions using temperature, humidity, pH or a combination. In practice,
evaluators use both real-time stability tests and accelerated stability tests. However, realtime stability testing can take a longer time to complete and so accelerated tests are often
used as temporary measures to expedite processes in product development [356].
Temperature is the most common acceleration factor used for chemicals, pharmaceuticals,
and biological products and accelerated aging testing is based on a thermodynamic
temperature coefficient formulated by Von't Hof stating that for every 10 C degree rise in
temperature, the rate of chemical reaction will at least double [358]. According to FDA
regulations and package testing industry, this is useful in defining and justifying
accelerated aging testing services [355]. An appropriate temperature for the accelerated
shelf life testing must be chosen to avoid failure conditions such as deformation due to
polymer melting. In our chosen study, the catheter segments are made up of Polyurethane
which has a melting temperature of 240 C.
Based on the shelf life testing of lysostaphin coated hernia-repair meshes performed by
Sriram Sankar in collaboration with CMC (data not shown), we decided to choose a
testing temperature of 60 C and a time period of 1 week for storage which is equivalent to
159
4 months of shelf life at room temperature. Before studying long term enzyme stability on
catheter segments, it is necessary to lyophilize the enzyme to improve its stability.
Lyophilization gives the opportunity to help avoid enzyme denaturation caused by
heating, by maintaining the samples frozen throughout the drying process. The first step in
the lyophilization process is to freeze all the water molecules present in the sample and
then it is placed in a lyophilizer and gradually heated in vacuum where all the water
sublimates as vapor. The samples were frozen overnight at -80 C, lyophilized for 24 h
and then sealed in air tight pouches (blasted with nitrogen) to prevent entry of air and
moisture.
We studied accelerated shelf life testing for plain and NP coated catheter samples
adsorbed with the highest coating concentration of 0.5 mg/ml of lysostaphin and
DispersinB using activity assays as described preciously. Turbidity assay was used to
quantify the activity of lysostaphin and colorimetric assay was used to test activity of
DispersinB. We also tested activity of lysostaphin samples coadsorbed with DispersinB at
the same coating concentration and vice versa. Figure 7.10 below shows the kinetic
constant for bacterial hydrolysis for lysostaphin coated samples for plain and NP coated
samples after 1 week at 60 C.
160
*
Figure 7.10. – Comparing the mean rate constants of bacterial lysis for both plain
and NP coated catheter samples after 1 day and post 1 week at 60C (equivalent
4 months at RT). Red bar – NPL-500 represents NP coated catheter segment with
initial lysostaphin concentration 500 µg/ml; Green bar – L-500 represents plain
catheter segment with initial lysostaphin concentration of 500 µg/ml; (Mean ± SD;
n= 2; * - p > 0.05)
We found that the initial rate of bacterial lysis for all the samples was almost equal to the
lytic activity assayed before shelf life testing (p>0.05 – One way ANOVA) for plain and
NP coated samples at coating concentration of 0.5 mg/ml. Again, no significant difference
between samples coated only with lysostaphin and coadsorbed with DispersinB (data not
shown). These results seem to suggest that lysostaphin retained all of its activity after 1
week at 60 C (equivalent of 4 months at RT). We had also quantified lysotaphin activity
before storage at 60 C, after a lyophilization time period of ~ 24 h and found there was no
difference in its lytic activity (data not shown). This was done in order to demonstrate if
161
the process of freeze drying the catheter samples could affect enzyme activity before
checking its stability for accelerated shelf life testing.
Figure 7.11. Comparing the mean rate constants of enzymatic hydrolysis for both
plain and NP coated catheter samples after 1 day, post lyophilization (1 day) and
post 1 week at 60C (equivalent 4 months at RT). Red bar – NPD-500 represents
NP coated catheter segment with initial DispersinB concentration 500 µg/ml;
Green bar – D-500 represents plain catheter segment with initial DispersinB
concentration of 500 µg/ml; (Mean ± SD; n= 2 for 1st two sets and n = 3 for
accelerated aging study )
In case of samples coated with DispersinB, we found that the rate of hydrolysis for
uncoated catheter segment was not significantly different (p>0.05; n =3; One way
ANOVA) for both plain and NP coated catheter segments indicating that DispersinB
possibly lost its activity during the accelerated testing study (Figure 7.11). But, we did
observe ~ 90% percentage drop in DispersinB activity on catheter segments (compared to
162
its activity before freeze-drying) via the colorimetric assay after a 24 h lyophilization
period suggesting that DispersinB
(unlike lysostaphin) did not retain its hydrolytic
activity after the freeze-drying step. During lyophilization process, stresses associated
with freezing and drying cycle can sometimes lead to aggregation of the protein molecules
with a consequent loss in enzyme activity. This happens because solutes (including salts in
solution) may reach concentrations as high as 50 times their initial concentration due to
separation of ice, leading to increased molecular interactions in the freeze-dried state [359,
360]. In order to prevent loss of enzyme activity during lyophilization, enzymes may be
modified chemically or stabilizers may be added to the buffer solution [342, 344, 361,
362]. For, e.g. addition of cryoprotectants or stabilizers like Sucrose might help prevent
denaturation of the functional enzymes such as DispersinB during the lyophilization
process [363].
7.4. Conclusion
In conclusion, adsorbed enzyme activity for both lysostaphin and DispersinB,
enzyme leaching, cytotoxicity and antibiofilm activity were not significantly different for
plain and NP coated catheter segments. Our study showed that coating of nanoparticles on
catheters overall did not significantly improve performance of enzymes on the catheters
when adsorbed alone or through coadsorption. This is in part, possibly due to lack of a
significantly higher surface area available for enzyme binding due to presence of
nanoparticles, as it appears that enzymes were bound to the bare part of the catheter
segment. We were unable to achieve good coating of PLA NPs on catheter surface,
despite using excess of the NPs (based on theoretical surface area of catheter) in the initial
163
NP suspension. Furthermore, expected improvement in stability has not been achieved,
again likely because most of the enzyme was present as adsorbed not on nanoparticles but
on the bare part of the catheter segment.
Our accelerated shelf life testing showed lysostaphin retaining all of its activity
after freeze-drying cycle and shelf life study, while DispersinB lost most of its activity
during the lyophilization process and all of its activity after the shelf life study. Further
work involving addition of stabilizers or cryoprotectant to preserve enzyme activity,
especially for a temperature sensitive enzyme such as DispersinB may be performed.
However, based on the overall similar antibiofilm activity of the segments coated by
lysostaphin, we conclude that use of lysostaphin may be preferable instead of DispersinB
because of its higher stability.
164
CHAPTER 8
CONCLUSIONS AND FUTURE RECOMMENDATIONS
8.1 Conclusions
One of the most serious complications in the use of prosthetic devices such as Central
venous catheters is device-associated infections and S.aureus is most frequently
encountered nosocomial pathogen in this setting. However, the emergence of antibiotic
resistant strains such as MRSA, has spurred the need for alternative sources of treatment.
Coatings of devices and implants using antibacterial enzymes and peptides have numerous
advantages including low rates of resistance, broad spectrum of action, short treatment
time and synergism (between peptides). Our study proposed to use nanoparticles as
coatings on biomedical devices for higher enzyme loading, low leaching and long term
durability. Based on the results from this study, the following conclusions can be drawn
regarding the application of nanotechnology for targeted delivery of antibacterial enzymes
to planktonic cell suspensions and as potential „surrogate surfaces‟ for enzyme adsorption
on devices to prevent bacterial colonization and subsequent biofilm formation:

Intrinsic properties of nanoparticles such as charge and surface charge density can
be used to improve targeting of enzymes to bacterial cell walls. Lysozyme
covalently conjugated to positively charged nanoparticles (modified with amine
groups) shows better lytic activity to Micrococcus lysodeikticus than free enzyme
and lysozyme conjugated to negatively charged nanoparticles (modified with
chloromethyl and sulfate groups) through charge directed targeting to negatively
charged bacterial cell walls.
165

High surface to volume ratio enables attachment of multiple ligands on the surface
of nanoparticles including a specific antibody along with the corresponding
antibacterial enzyme to improve targeting to bacterial pathogens. Lysostaphin
adsorbed to Poly (lactic) acid nanoparticles in the presence and absence of a S.
aureus antibody showed significantly better activity due to multiple-ligand
targeting and antibody-directed targeting compared to free enzyme against
S.aureus.

Coating of antibacterial enzymes (lysostaphin) to synthetic hernia-repair meshes
through physical adsorption can prevent bacterial colonization and implant
associated infection. Our data specifically showed that enzyme leaching surfaces
could be effective in inhibiting bacterial growth and is significantly dependent
upon coating concentration in vivo.

We did not observe a significant difference in enzyme loading, leaching,
cytotoxicity, antibiofilm activity and long term durability for enzymes (lysostaphin
and DispersinB) coated on plain and PLA nanoparticle coated Polyurethane
catheter segments. This was in part due to the absence of a significant monolayer
of nanoparticle coating on the surface of the catheter segment.
8.2 Future recommendations
Our studies aimed at using nanotechnology for targeted delivery of antibacterial enzymes
and as surface coatings for enzyme-based coatings on catheter segments. However, while
we showed that nanoparticles can be used to improve targeted delivery of antibacterial
enzymes to bacterial cells, we were not able to significantly improve characteristics of
enzyme coated catheters after precoating catheters with nanoparticles. PLA nanoparticles
166
did not strongly bind to the surface of Polyurethane catheter segment despite using several
fold excess of nanoparticles in the initial NP suspension, suggesting that nonspecific
adsorption might not be an appropriate method and PLA not the most appropriate material
for surface coating on Polyurethane. Stability of DispersinB could be improved during the
lyophilization process by addition of cryoprotectants although the theory behind coating
multiple enzymes including lysostaphin and DispersinB to achieve a synergistic
interaction seems a little unlikely. Lysostaphin, by itself seems to have antibiofilm activity
against S. aureus specifically and is more stable as an enzyme; hence coating of catheters
with lysostaphin alone could probably be just as effective. However, DispersinB could be
used in combination with other antimicrobials in inhibition of growth and biofilm
formation for other slime producing bacterial strains other than S. aureus. Addition of
cryoprotectants or stabilizers could prevent loss of activity during the lyophilization
process [363]. Before choosing an appropriate stabilizer, it is important that it does not
react with the functional enzyme and salt concentration in the buffer be low during the
adsorption process.
167
APPENDICES
168
CHAPTER 9
Appendix A
Structure of tween 20
169
Appendix B
Structure of p-nitrophenyl penta-N-acetyl-ß-chitopentaoside
170
Appendix C
Structure of Alexafluor succidimyl ester
171
Appendix D
Structure of Pluronic F68
172
Appendix E
Digital camera images showing reduction of turbidity of S. aureus cell suspension
(clearing) through bacterial lysis by ultrapro samples coated with lysostaphin at
different coating concentrations compared to uncoated samples.
173
Appendix F
Tapping mode AFM images showing topography and height of a polyurethane
catheter segment.
153.89 nm
20
Z[nm]
15
10
5
0
200nm
0
0.00 nm
174
200
400
X[nm]
600
800
1000
Appendix G
Structure of 4-nitrophenyl-N-acetyl- D-glucosaminide
175
Appendix H
Schematic of MTS assay
176
CHAPTER 10
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