Carboxymethyl chitosan functionalized magnetic nanoparticles for disruption of biofilms of straphylococcus aureus and escherichia coli

CARBOXYMETHYL CHITOSAN-FUNCTIONALIZED MAGNETIC NANOPARTICLES FOR DISRUPTION OF BIOFILMS OF STAPHYLOCOCCUS AUREUS AND ESCHERICHIA COLI CHEN TONG (B. ENG DUT) A THESIS SUBMITTED FOR THE DEGREE OF MASTER OF ENGINEERING DEPARTMENT OF CHEMICAL AND BIOMOLECULAR ENGINEERING NATIONAL UNIVERSITY OF SINGAPORE 2012 DECLARATION I hereby declare that this thesis is my original work and it has been written by me in its entirety. I have duly acknowledged all the sources of information which have been used in the thesis. This thesis has also not been submitted for any degree in any university previously. Chen Tong 25 Jan 2013 ACKNOWLEDGEMENT It is a great pleasure to thank many people whose help and suggestions were so valuable in my one year research work. First and foremost, I would like to express my sincerest and deepest appreciation to my supervisors, Professor Neoh Koon-Gee and Professor Kang En-Tang, at National University of Singapore, for their invaluable guidance, instructions, and discussion throughout this work. Professor Neoh’s abundant knowledge in biology related areas is always a source of inspiration to me in carrying out this project. Their enthusiasm, diligence, patience, and preciseness enlighten me on the road of scientific research, and even my future road of life. I am also indebted to Dr. Shi Zhilong, Dr. Liu Gang, Dr. Li Min, Cai Tao, Yang Wenjing, Xu Liqun, Wang Rong, for their fruitful discussion and comments during this work. I would like to express my particular gratitude to Xu Liqun, from whose generous consultation and invaluable experience I learnt heavily for my own work. In addition, my parents, Mr. Chen Dongsheng and Ms. Sun Yulan also gave me great support during this one year study in Singapore. Their unconditional love and sacrifice made me fully concentrate on my research work without concerning too much about the daily issues. Their consistent care and support enable me healthy enough, both mentally and physically, to finish this work. Last but not least, I would like to appreciate the financial support provided by the National University of Singapore. i TABLE OF CONTENTS ACKNOWLEDGEMENT ............................................................................................. i TABLE OF CONTENTS .............................................................................................. ii SUMMARY ................................................................................................................. iv NOMENCLATURE .......................................................................................................v LIST OF FIGURES .................................................................................................... vii CHAPTER 1 INTRODUCTION ...................................................................................1 CHAPTER 2 LITERATURE REVIEW.........................................................................5 2.1 Biofilm ................................................................................................................................ 6 2.1.1 Formation and development of biofilm ................................................................... 6 2.1.2 The mechanisms of resistance to antibiotics ........................................................... 7 2.1.3 Infectious diseases................................................................................................... 9 2.1.4 Disruption of biofilm............................................................................................. 10 2.2 Chitosan ............................................................................................................................ 11 2.2.1 Physical and chemical characterization ................................................................. 13 2.2.2 Antimicrobial action .............................................................................................. 14 2.2.3 Applications of chitosan ........................................................................................ 18 CHAPTER 3 EXPERIMENTIAL ...............................................................................21 3.1 Materials ........................................................................................................................... 22 3.2 Synthesis of carboxymethyl chitosan ................................................................................ 22 3.3 Synthesis of magnetic iron oxide nanoparticles (MNPs) .................................................. 23 3.4 Synthesis of magnetic carboxymethyl chitosan nanoparticles (CMCS-MNPs) ................ 23 3.5 Determination of antibacterial effcacy against planktonic cells ........................................ 25 3.6 Determination of biofilm disruption efficacy .................................................................... 26 3.7 Bacterial quantification ..................................................................................................... 29 3.8 Cytotoxicity of nanoparticles ............................................................................................ 29 3.9 Characterization ................................................................................................................ 30 CHAPTER 4 RESULTS AND DISCUSSIONS ..........................................................31 ii 4.1 Characterization of CMCS-MNPs .................................................................................... 32 4.2 Antibacterial efficacy against planktonic cells .................................................................. 35 4.3 Biofilms disruption ........................................................................................................... 38 4.4 Cytotoxicity of nanoparticles ............................................................................................ 47 CHAPTER 5 CONCLUSION AND RECOMMENDATIONS ...................................49 5.1 Conclusion ........................................................................................................................ 50 5.2 Recommendations ............................................................................................................. 51 REFERENCES ............................................................................................................53 iii SUMMARY Bacteria in biofilms are much more resistant to antibiotics and microbicides compared to their planktonic stage. Thus, to achieve the same antibacterial efficacy, a much higher dose of antibiotics is required for biofilm bacteria. However, the widespread use of antibiotics has been recognized as the main cause for the emergence of antibiotic-resistant microbial species, which has now become a major public health crisis globally. In this work, we present an efficient non-antibiotic-based strategy for disrupting biofilms using carboxymethyl chitosan (CMCS) coated on magnetic iron oxide nanoparticles (CMCS-MNPs). CMCS-MNPs demonstrate strong bactericidal activities against both Gram-positive Staphylococcus aureus (S. aureus) and Gram-negative Escherichia coli (E. coli) planktonic cells. More than 99% S. aureus and E. coli planktonic cells were killed after incubation with CMCS-MNPs for 10 h and 5 h, respectively. In the presence of a magnetic field (MF), CMCS-MNPs can effectively penetrate into both S. aureus and E. coli biofilms, resulting in a reduction of viable cells counts by 84% and 95%, respectively, after 48 h incubation, compared to the control experiment without CMCS-MNPs or CMCS. CMCS-MNPs are non-cytotoxic towards mammalian cells and can potentially be a useful antimicrobial agent to eliminate both planktonic and biofilm bacteria. iv NOMENCLATURE ATCC American type culture collection CH Chitosan CLSM Confocal laser scanning microscopy CMCS Carboxymethyl chitosan E. coli Escherichia coli DMEM Dulbecco’s modified eagle’s medium FAC Ferric ammonium citrate FTIR Fourier transform infrared spectroscopy GV Gentian violet LPS Lipopolysaccharide LTA Lipoteichoic acid MTT 3-(4, 5-dimethylthiazol-2-yl)-2, 5-diphenyltetrazolium bromide MNP Magnetic nanoparticle NIR NIR diode otopathogenic OM Outer membrane OPPA8 Otopathogenic pseudomonas aeruginosa PA Pseudomonas aeruginosa PDA Polydopamine PG Peptidoglycan PNIPAAm Poly(N-isopropylacrylamide) v P. mirabilis Proteus mirabilis PP Polypropylene S. aureus Staphylococcus aureus SEM Scanning electron microscopy SW Q-switched Nd-YAGSW TA Teichoic acid TGA Thermogravimetric analysis XPS X-ray photoelectron spectroscopy vi LIST OF FIGURES Figure 2-1 Biofilm maturation is a complex developmental process involving five stages Figure 2-2 Three hypotheses for mechanisms of antibiotic resistance in biofilms Figure 2-3 Structures of chitin and chitosan Figure 2-4 Schematic diagram illustrating synthesis of carboxymethyl chitosan Figure 2-5 Schematic view of the Gram-negative cell envelope Figure 2-6 Gram-positive cell walls Figure 3-1 Schematic illustration for the preparation of CMCS-MNPs and RITC-CMCS-MNPs Figure 3-2 Schematic representation of antibacterial assay using CMCS-MNPs against planktonic cells Figure 3-3 Schematic representation of antibacterial assay using CMCS-MNPs against biofilm Figure 4-1 FT-IR spectra of (a) MNPs, (b) PDA-MNPs and (c) CMCS-MNPs Figure 4-2 TGA curves of (a) MNPs, (b) PDA-MNPs and (c) CMCS-MNPs Figure 4-3 Hydrodynamic size of CMCS-MNPs after incubation in PBS for different periods Figure 4-4 Antibacterial effect of CMCS-MNPs (2.0 mg/mL) and CMCS (0.34 mg/mL) on (a) S. aureus and (b) E. coli suspensions (106 cells/mL). The controls refer to the bacterial suspensions without CMCS or vii CMCS-MNPs Figure 4-5 Antibacterial effect of MNPs (2.0 mg/ml) on (a) S. aureus and (b) E. coli suspensions (106 cells/mL). The controls refer to the bacterial suspensions without CMCS or CMCS-MNPs Figure 4-6 Effect of CMCS-MNPs (with or without MF) and CMCS on pre-grown (a) S. aureus biofilms and (b) E. coli biofilms after 12, 24, and 48 h. The controls refer to the respective pre-grown biofilms in sterile PBS without addition of CMCS or CMCS-MNPs. The prefix 1.0 and 2.0 represent 1.0 mg/mL and 2.0 mg/mL CMCS-MNPs suspension respectively; and the suffix (MF) indicates the application of magnetic field in the 5 min period when the biofilms were exposed to the CMCS-MNPs suspension. * denotes significant differences (p < 0.05) compared to the control experiment at the same incubation time Figure 4-7 CLSM (a,c) volume view and (b,d) cross-sectional view images of S. aureus biofilms exposed to RITC-CMCS-MNPs (2.0 mg/mL) (a,b) without a MF and (c,d) with a MF. Scale bar = 100 µm Figure 4-8 CLSM volume view images of (a-c) E. coli biofilms and (d-f) S. aureus biofilms: (a) and (d) pre-grown biofilms after incubation in PBS for 24 h, (b) and (e) with addition of CMCS-MNPs (2.0 mg/mL) without MF for 5 min and after incubation in PBS for 24 h, (c) and (f) with addition of CMCS-MNPs (2.0 mg/mL) with MF for 5 min and after incubation in PBS for 24 h. Scale bar = 100 µm viii Figure 4-9 SEM images of (a-c) E. coli biofilms and (d-f) S. aureus biofilms: (a) and (d) pre-grown biofilms after incubation in PBS for 24 h, (b) and (e) with addition of CMCS-MNPs (2.0 mg/mL) without a MF for 5 min and after incubation in PBS for 24 h, (c) and (f) with addition of CMCS-MNPs (2.0 mg/mL) with MF for 5 min and after incubation in PBS for 24 h Figure 4-10 Viability of 3T3 fibroblast cells incubated for 24 h in growth medium containing CMCS (0.34 mg/ml) and CMCS-MNPs (2.0 mg/ml) relative to the control (no CMCS or CMCS-MNPs added). The suffix (MF) indicates the application of magnetic field throughout the incubation period. Results are represented as mean ±standard deviation ix CHAPTER 1 INTRODUCTION 1 Chapter 1 Introduction Bacteria growing in biofilms are embedded within a self-produced matrix of extracelluar polymeric substance (EPS), and thus can be insensitive to antibiotics and microbicides that could eliminate them in the plankonic state (Branda et al., 2005, Ramage et al., 2010). In general, biofilm cells are 100- to 1000-fold more resistant to antibiotic treatment. The resistance mechanisms are associated with the morphology of the biofilms, whereby the EPS matrix of biofilms can present a generic barrier to the diffusion of antibiotics. Measurements of antibiotics penetration into biofilms have shown that some antibiotics cannot readily permeate biofilms (Stewart et al., 1996). Furthermore, the exchange of genetic materials and the mutation of bacteria in biofilms occur more frequently than in planktonic populations. Therefore, development of resistance mechanisms can quickly be selected for and propagated throughout the community. In addition, the cells in the deep layers of biofilms grow at a slower rate because of insufficiency of oxygen and nutrients compared to those located on the surface, and they become insensitive to antibiotics due to their reduced metabolic activities (Richards et al., 2009, Stewart et al., 2001, He et al., 2011). As a result of these resistance mechanisms, a much higher dosage of antibiotics is required to achieve the same antimicrobial efficacy on biofilm microbes than on planktonic ones (Anwar et al., 1990, Costerton et al., 1987, Khoury et al., 1992). Biofilm-associated infections have become one of the most devastating medical complications. For instance, the US Centers for Disease Control and Prevention estimated that healthcare-associated infections were among the top ten leading causes 2 Chapter 1 Introduction of death in the United States, accounting for 1.7 million infections and 99,000 associated deaths (Klevens et al., 2007). Many antibiotics including penicillin, methicillin and sulfonamides have been used in the treatment of bacterial infections. However, the widespread use of antibiotics in the agricultural and biomedical fields has been identified as the main cause for the emergence of multidrug-resistant microbes. Clearly, an antimicrobial strategy which is not antibiotic-based would be desirable for combating biofilm-associated infections. Lasers have been used for disrupting biofilms in recent years (Krespi et al., 2008). For instance, the combination of Q-switched Nd-YAGSW (SW) and NIR diode (NIR) lasers can result in a decrease of more than 43% of methicillin-resistant Staphylococcus aureus (S. aureus) biofilm cells (Krespi et al. 2011). However, the need for specialized equipment such as SW and NIR could be a limitation for the widespread use of these radiation-based treatment methods. Recently, it was reported that gentian violet (GV) and ferric ammonium citrate (FAC) possess biofilm disruption properties. After 24 h of continuous exposure to GV (1225 µmol/L), few live Pseudomonas aeruginosa (PA) biofilm cells were detected, and FAC at 250 µmol/L significantly decreased the fluorescence of otopathogenic Pseudomonas aeruginosa (OPPA8) biofilms after 24 h of exposure (p < 0.03) (Eric et al., 2008). In other investigations, MgF2 nanoparticles were shown to be capable of penetrating both Escherichia coli (E. coli) and S. aureus cells, and could restrict the formation of biofilms (Lellouche et al., 2009). Ag-loaded 3 Chapter 1 Introduction chitosan nanoparticles also show synergistic antimicrobial effect against S. aureus bacteria (Ali et al., 2011). Nevertheless, the use of MgF2 and Ag may not be appropriate as they pose possible environmental problems and toxicity to certain mammalian cells (Mukherjee et al., 2012, Kim et al., 2011). In this present study, an antimicrobial and anti-biofilm strategy involving the use of carboxymethyl chitosan (CMCS) coated on polydopamine (PDA) pre-treated magnetic iron oxide nanoparticles (MNPs) is presented. Chitosan is a cationic polysaccharide derived from chitin which is commonly extracted from crustacean shells. Its antibacterial properties (Li et al., 2008, Raafat et al., 2008, Lou et al., 2011) and biocompatible nature (Ahmadi et al., 2008, Mattanvee et al., 2009) have attracted considerable interest in recent years. The carboxymethylation of chitosan increases its solubility in water, and promotes the dispersion of CMCS-coated MNPs in aqueous media. The increase in –NH3+ groups, resulting from the intra- and intermolecular interaction between –COOH and –NH2 groups may also enhance the antibacterial properties of CMCS-coated MNPs (Liu et al., 2001). Our results showed that this antimicrobial system is highly effective in eliminating planktonic cells of both Gram-positive S. aureus and Gram-negative E. coli. The use of a magnetic field in combination with the CMCS-MNPs can also effectively disrupt the biofilms of these bacteria. 4 CHAPTER 2 LITERATURE REVIEW 5 Chapter 2 Literature Review 2.1 Biofilm A biofilm is a gathering of bacterial cells enclosed in a self-produced polymeric matrix composed of extracellular polymeric substances, mainly exopolysaccharides, proteins and nucleic acids. Biofilms may form on living or non-living surfaces and can be prevalent in natural, industrial and hospital settings (Hall-Stoodley et al., 2004, Lear et al., 2012). Biofilm cells often display enhanced tolerance towards antibiotics and immune responses and they also exhibit an altered phenotype with respect to growth rate and gene transcription, which are very different from the single-cells in a liquid medium (Madsen et al., 2012). 2.1.1 Formation and development of biofilm Biofilms are present on nearly all types of surfaces, ranging from industrial equipment to surgical implants, medical devices as well as living tissues. The formation of a biofilm begins with the initial attachment of free-floating microorganisms to surface. The first colonists adhere to surface initially through weak, reversible adhesion via van der Waals forces. Those cells can anchor themselves more permanently using cell adhesion structures such as pili (Karatan et al., 2009), when they are not immediately separated from the surface. Once the colonization has begun, the cells in biofilms grow through a combination of cell division and recruitment. The formation of a biofilm ended with the last step known as development, which may result in an aggregate cell colony becoming increasingly antibiotic resistant. Figure 2-1 shows a complex developmental process of biofilm maturation involving five stages: stage 1, 6 Chapter 2 Literature Review initial attachment; stage 2, irreversible attachment; stage 3, maturation Ⅰ; stage 4, maturation Ⅱ; stage 5, dispersion. Each stage of development in the diagram is paired with a photomicrograph of a developing Pseudomonas aeruginosa biofilms. All photomicrographs are shown to same scale. Figure 2-1 Biofilm maturation is a complex developmental process involving five stages (Monroe, 2007) 2.1.2 The mechanisms of resistance to antibiotics Bacteria growing in biofilms are embedded within a self-produced matrix of extracelluar polymeric substance (EPS), and thus can be insensitive to antibiotics and microbicides that could eliminate them in the plankonic state. In addition, this matrix protects the cells within it and facilitates communication among them through biochemical signals. Figure 2-2 shows the three main hypotheses for antibiotic resistance mechanisms in biofilms. 7 Chapter 2 Literature Review The first hypothesis is the possibility of slow or incomplete penetration of the antibiotics into the biofilms. Measurements of antibiotics penetration into biofilms in vitro have shown that some antibiotics readily permeate bacterial biofilms (Stewart et al., 1996). However, some antibiotics are adsorbed on the biofilm matrix which can reduce its penetration into the biofilms. This may account for the slow penetration of aminoglycoside antibiotics (Kumon et al., 1994) since these positively charged agents bind to the negatively charged polymers in the biofilm matrix. Secondly, the exchange of genetic materials and the mutation of bacteria in biofilms occur more frequently than in planktonic populations. Therefore, development of resistance mechanisms can quickly be selected for and propagated throughout the community. Some of the bacteria may differentiate into a protected phenotypic state and become more resistance to antiobics (Tamilvanan, 2010). The third mechanism of antibiotic resistance is the altered chemical microenvironment within the biofilms. The depletion of a substrate or accumulation of an inhibitive waste product may cause some bacteria to enter into a non-growing state, in which they become insensitive to antibiotics. De Beer et al. (1994) reported that oxygen can be completely consumed in the surface layers of a biofilm, leading to anaerobic niches in the deep layers of the biofilms. Aminoglycoside antibiotics, for instance, are less effective against the same microorganism in anaerobic than in 8 Chapter 2 Literature Review aerobic conditions (Tack et al., 1985). Local accumulation of acidic waste products may lead to pH differences between the biofilm surface and the biofilm interior (Vroom et al., 1999), which could directly antagonise the action of an antibiotic. For instance, Baudoux et al. (2007) reported that antibacterial activities against methicillin-susceptible S. aureus decreased 8-fold of oxacillin between pH 7.4 and 5.0. Figure 2-2 Three hypotheses for mechanisms of antibiotic resistance in biofilms (Stewart et al., 2001) 2.1.3 Infectious diseases 9 Chapter 2 Literature Review Biofilms have been found to be involved in a wide variety of microbial infections in the body, and they account for nearly 80% of all infections. The US Centers for Desease Control and Prevention reported that biofilm-associated infections were among the top ten leading causes of death in the United State, accounting for 1.7 million infections and 99,000 associated deaths (Klevens et al., 2007). There are two main aspects of biofilm-associated infections, common problems such as urinary tract infections, catheter infections, coating contact lenses, gingivitis, and the less common but more lethal processes such as endocarditis, infections in cystic fibrosis and infections of permanent indwelling devices such as joint prostheses and heart valves. It is apparent that biofilm-associated infections can potentially become one of the most devastating medical complications, if new and better approaches for combating them are not implemented. 2.1.4 Disruption of biofilm Much of work has been done with the purpose of disrupting the biofilms: (1) Laser and photodynamic treatment have been used to disrupt bacterial biofilms. Krespi et al (2011) reported that the combination of Q-switched Nd-YAGSW (SW) and NIR diode (NIR) lasers can result in a decrease of more than 43% of methicillin-resistant S. aureus biofilm cells. However, the need for specialized equipment such as SW and NIR could be a limitation for the widespread use of these radiation-based treatment methods. (2) Gentian violet (GV) and ferric ammonium citrate (FAC) have also been reported to possess biofilm disruptive activity. After 24 h of continuous exposure to 10 Chapter 2 Literature Review GV (1225 µmol/L), few live Pseudomonas aeruginosa (PA) biofilm cells were detected (Eric et al., 2008). FAC at 200 µM caused disruption of PA biofilms after a 5-day incubation period (Musk et al., 2005). (3) Lellouche et al. (2009) demonstrated that nanosized magnesium fluoride (MgF2) was capable of penetrating E. coli and S. aureus cells and inhibiting biofilm formation. (4) Magnetic microspheres coated with Ag nanoparticles-loaded multilayers were also shown to possess significant bactericidal properties against both Gram-positive Staphylococcus epidermidis and Gram-negative E. coli bacteria (Lee et al., 2005). Nevertheless, the use of MgF2 and Ag may not be appropriate as they pose possible environmental problems and toxicity to certain mammalian cells (Mukherjee et al., 2012, Kim et al., 2011). In the present work, magnetic iron oxide nanoparticles (MNPs) functionalized with bactericidal moieties are used for disruption of biofilms. MNPs are iron oxide particles with diameters between about 1 and 100 nm, and they have attracted extensive interest in biomedical field due to their superparamagnetic properties, biocompatibility and lack of toxicity to humans (Hanini et al., 2011, Markides et al., 2012). With the use of a magnetic field, the functionalized nanoparticles can then be delivered to specific locations where bacteria were present. 2.2 Chitosan Chitosan is a cationic polysaccharide derived from chitin, which is commonly extracted from crustacean shells such as crabs and shrimp, the cuticles of insects, and 11 Chapter 2 Literature Review the cell walls of fungi. Figure 2-3 shows the structures of chitin and chitosan. Chitin is the most abundant natural amino polysaccharide (Majeti N. V. R. K., 2000) and represents the major source of nitrogen accessible to countless living terrestrial and marine organisms. The antibacterial properties (Li et al., 2008, Raafat et al., 2008, Lou et al., 2011) and biocompatible nature (Ahmadi et al., 2008, Mattanavee et al., 2009) of chitosan have attracted considerable interest in recent years. The carboxymethylation of chitosan increases its solubility in water, and the increase in −NH3+ groups, resulting from the intra- and intermolecular interaction between −COOH and −NH2 groups, may also enhance the antibacterial properties of CMCS (Liu et al., 2001). The aim of the present study is to formulate an antimicrobial and antibiofilm strategy, and chitosan is considered one of the most promising materials for this purpose. OH Chitin H3COC O NH HO O O * HO H3COC OH O NH HO H3COC OH * O NH Deacetylation OH Chitosan OH O NH2 HO O O * HO O NH2 * O HO NH2 OH Figure 2-3 Structures of chitin and chitosan (Jayakumar et al., 2010) 12 Chapter 2 Literature Review 2.2.1 Physical and chemical characteristics Chitosan is a polysaccharide composed of N-glucosamine and N-acetyl-glucosamine units, in which the number of N-glucosamine units exceeds 50% (Sodhi Rana et al., 2001). Chitosan has found several applications due to its excellent chemical, physical, and biological properties, such as biocompatibility, biodegradability, nontoxicity, adsorptive properties, and most importantly, antimicrobial activity. Some properties of chitosan such as the degree of N-deacetylation, molecular weight and solubility can, and to a great extent, influence the antibacterial efficacy. One of the most important parameter to examine closely is the degree of deacetylation of chitin. Takahashi et al. (2008) reported that the higher degree of deacetylation, the higher antibacterial efficacy of chitosan against S. aureus and E. coli bacteria. In addition, the molecular weight of chitosan can also affect the antimicrobial ability (Tsai et al., 2006). Viscometry is the simplest and most rapid method for determining the molecular weight. The constants а and κ in the Mark-Houwink equation have been determined in 0.1 M acetic acid and 0.2 M sodium chloride solution. The intrinsic viscosity is expressed as [η] = κMа = 1.81 * 10-3 M0.93, η is the intrinsic viscosity of chitosan solution and M is the average molecular weight (Kumar, 2000). Chitosan is a polyelectrolyte in acidic media because of the protonation of the amine (-NH2) groups. For instance, when chitosan is dispersed in acetic acid solution at different concentrations the following equilibria have to be considered: 13 Chapter 2 Literature Review CH3COOH + H2O CH3COO- + H3O+ Chit-NH2 + H3O+ Chit-NH3+ + H2O Rinaude et al. (1999) reported that complete solubilization was obtained when the degree of protonation exceeded 50% and the ([CH3COOH] / [Chit-NH2]) ratio was 0.6. Despite chitosan’s desirable solubility in acid media, its actual use is limited by the poor solubility in water. A lot of modification techniques and derivatives such as O-carboxymethyl chitosan, N-carboxymethyl chitosan and O-succinyl chitosan have been developed to improve its solubility. Among the water-soluble chitosan derivatives, O-carboxymethyl chitosan (Figure 2-4) is an amphiprotic ether derivative, containing –COOH groups and –NH2 groups in the molecule. There are many outstanding properties of O-carboxymethyl chitosan such as non-toxicity, biocompatibility, antibacterial, and antifungal bioactivity (Jayakumar et al., 2010). COOH O OH * O HO O NH2 * ClCH3COOH, 60OC * O HO O * NH2 Figure 2-4 Schematic diagram illustrating the synthesis of O-carboxymethyl chitosan 2.2.2 Antimicrobial action The exact mechanisms of antibacterial activities of chitosan and its derivatives are still unknown. It is known that the antimicrobial activity is influenced by a number of factors. 14 Chapter 2 Literature Review (a) Chitosan structure The polycationic structure of chitosan is a prerequisite for antibacterial activity. When the environmental pH is below 6.5 (the pKa value of chitosan), electrostatic interaction between the polycationic chitosan and the predominantly anionic components of the microbial cell membrane plays a primary role in the antibacterial activity. Through this process, chitosan can disrupt the normal functions of the cell membrane by promoting cell lysis and by inhibiting nutrients transport (Eldin et al., 2008, Gu et al., 2007). When the positive charge density of chitosan increases, the antibacterial property will increase correspondingly, as is the case with quaternized chitosan (Xie et al., 2007). In addition, the number of amino groups linking to C-2 on the chitosan backbone also plays an important role in the electrostatic interaction. Large numbers of amino groups are able to enhance the antibacterial activity. Another but still controversial mechanism is that the positively charged chitosan interacts with cellular DNA of some fungi and bacteria, which consequently inhibits the RNA and protein synthesis (Meng et al., 2012). In this mechanism, chitosan must be hydrolyzed to low molecular weight to penetrate into the cell of microorganisms (Tokura et al., 2007). (b) Microorganism structure Gram-negative bacteria possess an outer membrane (OM) that contains lipopolysaccharide (LPS), which provide the bacteria with a hydrophilic surface 15 Chapter 2 Literature Review (Figure 2-5). The lipid components and the inner core of the LPS molecules contain anionic groups (phosphate, carboxyl), which contribute to the stability of the LPS layer through electrostatic interactions with divalent cations (Helander et al., 1997). Removal of these cations by chelating agents results in destabilization of the OM through the release of LPS molecules. The OM serves as a penetration barrier against macromolecules and hydrophobic compounds, and thus Gram-negative bacteria are relatively resistant to hydrophobic antibiotics and toxic drugs. Therefore, overcoming the outer membrane is a prerequisite for any material to exert bactericidal activity towards Gram-negative bacteria (Kong et al., 2008a). Figure 2-5 Schematic view of the Gram-negative cell envelope (Helander et al., 1997) The cell wall of Gram-positive bacteria comprises peptidoglycan (PG) and teichoic acid (TA) (Figure 2-6). TA is an essential polyanionic polymer of the cell wall of 16 Chapter 2 Literature Review Gram-positive bacteria, which traverses the wall to contact with the PG layer. They can be either anchored into the outer leaflet of the cytoplasmic membrane via a glycolipid (lipoteichoic acids, LTA) or covalently linked to N-acetylmuramic acid of the PG layer (Raafat et al., 2008). Poly (glycerol phosphate) anion groups make TA responsible for structural stability of the cell wall. Besides, it is crucial for the function of various membrane-bound enzymes. Comparatively, TA's counterpart, LPS in the cell wall of Gram-negative bacteria, acts in a similar fashion. Figure 2-6 Gram-positive cell walls (Cabeen et al., 2005) Despite the distinction between Gram-negative and Gram-positive bacterial cell walls, antibacterial modes both begin with the interactions at the cell surface which compromise the OM or cell wall. The LPS and proteins in the Gram-negative bacteria OM are held together by electrostatic interactions with divalent cations that are required to stabilize the OM. Polycations may compete with divalent metals for binding with polyanions when the pH is below pKa of chitosan and its derivatives. However, chelation occurs when pH is above the pKa. Replacement of divalent 17 Chapter 2 Literature Review metals present in the cell wall will likely disrupt the integrity of the cell wall or influence the activity of degradative enzymes. For Gram-positive bacteria, LTA could provide a molecular linkage for chitosan at the cell surface, allowing it to disturb membrane functions (Raafat et al., 2008). Once the cells lose the protection of the cell wall, the cell membrane is exposed to the external influence. The functions of cell membrane can be changed consequently, with alteration in the membrane permeability (Kong et al., 2008a). 2.2.3 Applications of chitosan (a) Food preservation Chitosan has been approved as a food additive in Korea and Japan since 1995 and 1983, respectively (KFDA, 1995, Weiner, 1992). Due to its ability of forming semi-permeable film, chitosan coating can be expected to modify the environment of packaged food, to decrease the transpiration loss (Elghaouth et al., 1991) and to delay the ripening of fruits (Elghaouth et al., 1992). As a component of packaging material, chitosan not only retards microorganism growth in food, it also improves the quality and shelf life of food. Various kinds of chitosan-based packaging films modified with new polymeric material such as chitosan/polyethylene oxide film (Maher et al., 2008) and chitosan-nylon-6/Ag blended membranes (Ma et al., 2008) have been developed. Instead of polyethylene or polypropylene petrochemical materials which are inedible or not made from renewable natural resources, these new materials are environmentally-friendly and biodegradable. 18 Chapter 2 Literature Review (b) Medical industry Chitosan has been used in the area of health care and hygienic applications because it is a natural, biocompatible, anti-infective mucoadhesive, and hemostatic polymer, which may be incorporated into fibers, membrane, or hydrogel, and used for wound dressing, drug delivery carrier and orthopaedic tissue engineering. An ideal wound dressing material must be capable of absorbing the exuded liquid from the wounded area and should permit water evaporation at a certain rate and allow no microbial transport (Yang et al., 2004). Chitosan immobilized on poly(N-isopropylacrylamide) (PNIPAAm) gel/polypropylene (PP) nowoven composites surface have hydrogel-forming properties and are considered to be advantageous in their application as a wound dressing material (Chen et al., 2005). Surgical and pharmaceutical materials introduced into human body for tissue engineering or as drug release systems, for instance, suffer from potential complications arising from microorganism infections. It is apparent that once the introduced materials are infected, high morbidity and mortality rate can be expected. Therefore, efforts have focused on the development of bacterial-resistant prosthetic parts through binding of antimicrobial polymers to the materials. For instance, chitosan hydrogel coated grafts, crosslinked upon ultraviolet light irradiation, exhibited a resistance against E. coli in vitro and in vivo (Fujita et al., 2004). Silicone is widely used for implantable biomedical devices such as catheters (Stevens et al., 2009) and stents (Venkatesan et al., 2010). Wang et al. (2012) reported that O-carboxymethyl chitosan coated silicone 19 Chapter 2 Literature Review surface can inhibit the formation of E. coli and Proteus mirabilis (P. mirabilis) biofilms under both static and flow conditions. 20 CHAPTER 3 EXPERIMENTAL 21 Chapter 3 Experimental 3.1 Materials Polystyrene (PS) sheets of 1.2 mm thickness were purchased from Goodfellow. Ferric chloride hexahydrate (FeCl3·6H2O, > 99%), ferrous chloride tetrahydrate (FeCl2·4H2O, > 99%), dopamine hydrochloride (> 99%), monochloroacetic acid (> 99%), rhodamine isothiocyanate (RITC), dimethyl sulfoxide (DMSO), 3-[4,5-dimethyl-thiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT) and folate-free Dulbecco’s modified Eagle’s medium (DMEM) were obtained from Sigma-Aldrich (St. Louis, MO). Chitosan was purchased from CarboMer Inc. and used as received. Ultra-pure water (> 18.2 MΩ cm, Millipore Milli-Q system) was used in the experiments. S. aureus 25923, E. coli DH5α and 3T3 fibroblasts were obtained from American Type Culture Collection (ATCC). Sodium hydroxide (NaOH), potassium bromide (KBr), isopropanol, ethanol and acetone were all analytical reagent (AR) grade and obtained from Sigma-Aldrich or Merck Chem. Co.. 3.2 Synthesis of Carboxymethyl Chitosan (CMCS) Carboxymethyl chitosan (CMCS) was prepared according to a method described by Chen et al. (2003). 3.00 g of purified chitosan was added to 40% (w/w) aqueous NaOH and kept at 0°C overnight for alkalization. The cold alkali solution was put into a 250 mL reactor containing 60 mL isopropanol, and then 9.00 g of monochloroacetic acid in isopropanol (3 mL) was slowly added to the mixture over a 30 min period. After reaction for 12 h at room temperature, 200 mL of 70% (v/v) ethanol was added to stop the reaction. Finally, the solid was filtered, washed with ethanol and dried in a 22 Chapter 3 Experimental vacuum oven at 60°C for 24 h. The products were dissolved in dilute ammonia (0.1 g/mL) and centrifuged to remove the unreacted chitosan. The CMCS was precipitated by ethanol from the water-soluble portion, filtered and dried under reduced pressure at 60°C for 24 h. 3.3 Synthesis of Magnetic Iron Oxide Nanoparticles (MNPs) The MNPs were prepared using a controlled coprecipitation method following the reported procedure (Mikhaylova et al., 2004). In brief, FeCl3·6H2O (6.75g, 25 mmol), FeCl2·4H2O (2.48g, 12.5 mmol) and 1 mL 37% (v/v) HCl were dissolved in 24 mL ultra-pure water under vigorous stirring. The coprecipitation of MNPs was achieved by adding the iron solution to 250 mL of 0.5 M NaOH (under stirring at 1000 rpm), which was preheated to 80°C. The reaction was carried out for 1 h under the protection of nitrogen. The particles were then collected by sedimentation with a help of an external magnet and washed several times with ultra-pure water until a stable ferrofluid was obtained. The solid MNPs were freeze-dried and stored under nitrogen prior to further modification and characterization. 3.4 Synthesis of Magnetic Carboxymethyl Chitosan Nanoparticles (CMCS-MNPs) The CMCS-MNPs were synthesized as reported by Lee et al. (2007) with some minor modifications. 30 mg of MNPs and 45 mg of dopamine hydrochloride were added into 30 mL of 10 mM Tris-Cl solution (pH = 8.5) and dispersed by sonication for 1 h 23 Chapter 3 Experimental in an ice bath (Young et al., 2009). The reaction mixture was stirred at room temperature for 3 h to obtain the polydopamine coated magnetic nanoparticles (PDA-MNPs). The PDA-MNPs were collected under a magnetic field, washed three times with ultra-pure water to remove any loosely adsorbed PDA , and then dispersed in 20 mL phosphate buffered saline (PBS (10 mM, pH = 7.4)). After that, 20 mL of CMCS solution (10 mg/mL in PBS) was added, and the reaction mixture was incubated overnight in an orbital shaker at 180 rpm. The CMCS-MNPs were collected by centrifugation, and washed three times with ethanol and water to remove the excess CMCS. For the preparation of fluorescent RITC-CMCS-MNPs (Bhattacharya et al., 2011), 10 mg CMCS-MNPs was dispersed in 30 mL PBS, and 1 mL of RITC solution (1 mg/mL in DMSO/H2O (1/1, v/v)) was then added dropwise to the mixture. The reaction mixture was ultrasonicated in the dark for 1 h. The nanoparticles were collected under a magnetic field and washed with ultra-pure water. 24 Chapter 3 Figure 3-1 Experimental Schematic illustration for the preparation of CMCS-MNPs and RITC-CMCS-MNPs 3.5 Determination of Antibacterial Efficacy against Planktonic Cells S. aureus and E. coli were cultured in tryptic soy broth and nutrient broth, respectively, overnight at 37°C. The bacterial suspensions were centrifuged at 2700 rpm for 10 min. After removal of the supernatant, the cells were washed twice with sterile PBS and then resuspended in PBS to reach a concentration of 106 cells/mL. All lab wares were sterilized under UV irradiation for 1 h before the experiments. Five mL of the bacterial-containing PBS suspension was mixed with 5 mL CMCS-MNPs (4.0 mg/mL) or 5 mL CMCS solution (0.68 mg/mL, to maintain the same concentration of CMCS as that in CMCS-MNPs which contained ~ 17% CMCS 25 Chapter 3 Experimental as determined by thermogravimetric analysis (TGA)). The final concentration of CMCS-MNPs and CMCS in the bacteria-containing PBS suspension was 2.0 mg/mL and 0.34 mg/mL, respectively. Control experiments were carried out with PBS solution without CMCS-MNPs or CMCS. The suspensions were then placed in sterile tubes in an orbital shaker maintained at 37°C and 200 rpm (Figure 3-1). The number of viable bacteria at 2, 4, 6, 8 and 10 h for S. aureus and at 1, 2, 3, 4 and 5 h for E. coli was determined using the method described in the section on "Bacterial Quantification". CMCS-MNPs added 37 °C, 200 rpm Bacterial suspension Figure 3-2 Schematic representation of antibacterial assay using CMCS-MNPs against planktonic cells 3.6 Determination of Biofilm Disruption Efficacy PS sheets were cut into 1 × 1 cm2 pieces, washed ultrasonically in acetone and ethanol, for 10 min in each step, and then rinsed with copious ultra-pure water after each wash. After that, the substrates were immersed in ultra-pure water for 10 min, and then blown dry under a flow of purified N2. The PS substrates were sterilized with UV 26 Chapter 3 Experimental irradiation for 1 h before use. Bacterial broth suspension (1 mL) at a concentration of 106 cells/mL was added to each 24-well plate with PS substrates (for scanning electron microscopy (SEM) and confocal laser scanning microscopy (CLSM) observation) or without PS substrates (for viable bacterial cell count). The biofilms were allowed to grow at 37°C for 48 h, with the culture broth replenished after 24 h. For the viable bacterial cell count experiment, 1 mL of CMCS-MNPs (1.0 or 2.0 mg/mL) or CMCS (0.34 mg/mL) in PBS solution was added to each well with pre-grown biofilms. A magnet (39.5 mm × 24.5 mm × 5.0 mm, field strength 355 ± 30 G) was placed under the 24-well plate and the magnetic field was maintained for 5 min before the suspension was removed (Figure 3-2). The 5 min exposure time to CMCS-MNPs was chosen because it was found that ~ 95% of these nanoparticles would settle to the bottom of the well within 5 min under the magnetic field (as determined by UV-visible absorption). The wells were then refilled with 1 mL sterile PBS (10 mM, pH = 7.4), and the bacteria were allowed to incubate at 37°C for 12, 24 and 48 h. Control experiments were carried out with the pre-grown biofilms in sterile 1 mL PBS (10 mM, pH = 7.4) without any CMCS-MNPs or CMCS. For the SEM observation, the PS substrates were removed from the wells with sterile forceps, washed three times with sterile PBS (10 mM, pH= 7.4), fixed in 3% (v/v) glutaraldehyde in PBS solution for 30 min at room temperature, and immersed in 25%, 75%, 100% ethanol stepwise for dehydration. The PS substrates were dried, coated with platinum, and observed under SEM (JEOL, model 27 Chapter 3 Experimental 5600LV, Tokyo, Japan). For the CLSM observation, the CMCS-MNPs-treated biofilms were stained with LIVE/DEAD Baclight viability kit (Molecular Probes Inc, L1352). In addition, the biofilms treated with fluorescent RITC-CMCS-MNPs were only stained with SYTO9 dye from the kit to differentiate between the live bacteria (green fluorescence) and the nanoparticles (red fluorescence). After 20 min incubation in the dark, images were taken using a Nikon Ti-E microscope with A1 confocal system (Nikon, Tokyo, Japan). A Multi-Argon 488 nm laser was used as the source of illumination, with 488 nm excitation, long-pass 500-530 nm emission filter settings for the green signal and 570-620 nm emission filter settings for the red signal. NIS-Elements C software was used to generate the volume view images of the biofilms. CMCS-MNPs added MF applied 5 min Pre-grown biofilms CMCS-MNPs replaced by PBS MF removed Figure 3-3 Schematic representation of antibacterial assay using CMCS-MNPs against biofilm 28 Chapter 3 Experimental 3.7 Bacterial Quantification For quantification of the viable bacterial cells in the biofilms, the 24-well plate after the pre-determined incubation period (12, 24, and 48 h) was subjected to ultrasonication for 10 min in a 100 W ultrasonic bath operating at a nominal frequency of 50 Hz. The bacteria-containing suspensions were transferred into 15 mL centrifuge tubes and subjected to rapid vortex mixing for 20 s. Serial ten-fold dilutions were performed and the number of bacteria cells was counted using the spread plate method. For the planktonic cells experiments, 0.1 mL of the bacterial suspension after the specified incubation period was pipetted out and added into 0.9 mL sterile PBS (10 mM, pH = 7.4). Serial ten-fold dilutions were performed and the number of bacteria cells was counted using the spread plate method. All experiments were performed in quintuplicate with five substrates and the mean values were calculated. 3.8 Cytotoxicity of Nanoparticles 3T3 fibroblast cells were cultured in DMEM supplemented with 10% fetal bovine serum, 100 IU/mL penicillin, and 1 mM L-glutamine. The cells were seeded at a density of 104 cells per well and incubated for 24 h at 37°C before the medium was replaced with fresh one containing CMCS and CMCS-MNPs at a concentration of 0.34 mg/mL and 2.0 mg/mL, respectively. The runs with CMCS-MNPs were carried out with and without a magnet placed at the bottom of the well. Control experiments were carried out using the complete growth culture medium without CMCS or 29 Chapter 3 Experimental CMCS-MNPs. The cells were incubated at 37°C for another 24 h in the medium. The culture medium from each well was then removed and 0.9 mL of medium and 0.1 mL of MTT solution (5 mg/mL in PBS) were added to each well. After 4 h of incubation at 37°C, the medium was removed and the formazan crystals were solubilized with 1 mL DMSO for 15 min. The optical absorbance was then measured at 570 nm on a microplate reader (Tecan GENios). The results were expressed as percentages relative to the results obtained from the control experiments. 3.9 Characterization The hydrodynamic size of the nanoparticles was determined using dynamic light scattering (DLS) with a Brookhaven LLS 90 Plus Particle Size Analyzer. To test the stability of the CMCS-MNPs in aqueous medium, 10 mg CMCS-MNPs were dispersed in 30 mL PBS (10 mM, pH= 7.4) and stored at 4°C in a refrigerator. After pre-determined periods of time, 1 ml of the suspension was pipetted out and ultrasonicated for 5 min followed by the determination of the hydrodynamic size. The zeta potential of the nanoparticles in PBS and bacterial suspension in PBS (106 cells/mL) were measured using a Zetasizer Nano-ZS analyzer (Malvern Instruments). Thermogravimetric analysis was carried out with a TGA 2050 analyzer (TA Instruments). The samples were heated from room temperature to 800°C at a heating rate of 10 °C/min in air. Fourier transform infrared (FT-IR) spectra of samples dispersed in KBr pellets were obtained in the transmission mode on a Bio-Rad FT-IR spectrophotometer (Model FTS135). 30 CHAPTER 4 RESULT AND DISCUSSIONS 31 Chapter 4 Results and Discussions 4.1 Characterization of CMCS-MNPs Figure 4-1 shows the FT-IR spectra of MNPs, PDA-MNPs, and CMCS-MNPs, respectively. Comparing the FT-IR spectrum of MNPs (Figure 4-1a) and PDA-MNPs (Figure 4-1b), an absorption band at 1503 cm-1 can be observed in the latter. This band is the characteristic absorption band of aromatic rings of PDA (Zhu et al., 2011), indicating the successful coating of PDA on the surface of MNPs. After the conjugation with CMCS, the spectrum of CMCS-MNPs (Figure 4-1c) showed not only the main absorption peak of naked Fe3O4 nanoparticles at 590 cm-1 (Figure 4-1a) but also the characteristic peaks of CMCS at 1596 cm-1 and 1422 cm-1, which are assigned to the respective asymmetry and symmetry stretch vibration of the COOgroups (Liang et al., 2007). The large difference between the zeta potential of PDA-MNPs (41.6 ± 9.1 mV) and CMCS-MNPs (-40.4 ± 5.8 mV) also indicates that CMCS has been successfully coated on the PDA-MNPs (Table 4-1). The positive zeta potential of PDA-MNPs is due to the amine groups in PDA (Gomes et al., 2009, Aviles et al., 2010), while the negative zeta potential of CMCS-MNPs can be attributed to –COOH groups in CMCS (Shi et al., 2009). Despite the fact that CMCS-MNPs are highly negatively-charged, by using these nanoparticles with a magnetic field, the nanoparticles can effectively penetrate into and disrupt bacterial biofilms (see Section 4.3). The hydrodynamic size of PDA-MNPs and CMCS-MNPs are 105.2 ± 1.1 nm and 268.9 ± 5.3 nm, respectively. The increase in the hydrodynamic diameter of CMCS-MNPs further confirms the presence of the CMCS layer on the surface of PDA-MNPs. 32 Chapter 4 Results and Discussions Table 4-1 Properities of PDA-MNPs and CMCS-MNPs PDA- MNPs CMCS-MNPs  Potential/ mV 41.6 ±9.1 -40.4 ±5.8 Hydrodynamic Size/ nm 105.2 ±1.1 268.9 ±5.3 590 Transmittance (%) (a) 1503 (b) (c) 1596 4000 3000 2000 1422 1000 Wavenumber (cm-1) Figure 4-1 FT-IR spectra of (a) MNPs, (b) PDA-MNPs and (c) CMCS-MNPs Figure 4-2 shows the respective TGA plot of MNPs, PDA-MNPs and CMCS-MNPs in an air atmosphere. The PDA and CMCS layers accounted for 28% and 17% of the weight of the CMCS-MNPs, respectively. The PDA and CMCS confer a high level of dispersibility to the nanoparticles in aqueous media, and Figure 4-3 shows that the hydrodynamic size of CMCS-MNPs did not change significantly during one month in PBS. 33 Chapter 4 Results and Discussions 100 Weight (%) (a) 80 (b) 60 (c) 40 0 200 400 600 800 Temperature (OC) Figure 4-2 TGA curves of (a) MNPs, (b) PDA-MNPs and (c) CMCS-MNPs Hydrodynamic Size (nm) 400 300 200 100 0 1 2 3 4 Weeks Figure 4-3 Hydrodynamic size of CMCS-MNPs after incubation in PBS for different periods. 34 Chapter 4 Results and Discussions 4.2 Antibacterial Efficacy against Planktonic Cells The number of viable S. aureus and E. coli cells as a function of time after the addition of CMCS and CMCS-MNPs to the bacterial suspension (106 cells/mL in PBS) was investigated by the spread plate method. The antibacterial activity of CMCS and CMCS-MNPs are compared in Figure 4-4. As shown in Figure 4-4a, the number of S. aureus viable cells in the suspension decreased by 26% and 30% after 2 h in contact with CMCS and CMCS-MNPs, respectively, whereas the viable cell number did not change significantly in the absence of CMCS and CMCS-MNPs (control). The number of viable cells progressively decreased in the presence of CMCS and CMCS-MNPs, and more than 99% cells were killed after 10 h in these suspensions. For E. coli (Figure 4-4b), the reduction in viable cells progressed more rapidly, and the number of viable cells decreased more than 50% and 54% after 2 h in contact with CMCS and CMCS-MNPs, respectively, and in 5 h, almost all bacterial cells have been killed. Eric et al. (2008) had suggested that iron in FAC is capable of killing bacteria, but we found that when the antibacterial assays were carried out with MNPs and PDA-MNPs, no significant antibacterial properties against both S. aureus and E. coli bacteria was observed (p > 0.05 compared to control, Figure 4-5). The difference in antibacterial action of FAC and MNPs may be attributed to the presence of free iron ions in FAC, but not in MNPs. 35 Chapter 4 Results and Discussions 5 No. of Bacterial Cells (10 /mL) a 12.5 Control CMCS CMCS-MNPs 10.0 7.5 5.0 2.5 0.0 0 2 4 6 8 10 12 Time (h) 5 No. of Bacterial Cells (10 /mL) b 12.5 Control CMCS CMCS-MNPs 10.0 7.5 5.0 2.5 0.0 0 1 2 3 4 5 6 Time (h) Figure 4-4 Antibacterial effect of CMCS-MNPs (2.0 mg/mL) and CMCS (0.34 mg/mL) on (a) S. aureus and (b) E. coli suspensions (106 cells/mL). The controls refer to the bacterial suspensions without CMCS or CMCS-MNPs. 36 Chapter 4 Results and Discussions Control 12.5 PDA-MNPs MNPs 5 No. of Bacterial Cells (10 /mL) a 10.0 7.5 5.0 2.5 0.0 0 2 4 6 8 10 12 Time (h) Control 12.5 PDA-MNPs MNPs 5 No. of Bacterial Cells (10 /mL) b 10.0 7.5 5.0 2.5 0.0 0 1 2 3 4 5 6 Time (h) Figure 4-5 Antibacterial effect of MNPs (2.0 mg/ml) and PDA-MNPs (2.0 mg/ml) on (a) S. aureus and (b) E. coli suspensions (106 cells/mL). The controls refer to the bacterial suspensions without MNPs or PDA-MNPs. 37 Chapter 4 Results and Discussions The observed bactericidal action in Figure 4-4 is attributed to the effect of CMCS, which can bind to bacterial cell surface via electrostatic interaction of its positively charged amino groups with the negatively charged cell membrane (Raafat et al., 2008). Through this process, CMCS can disrupt the normal functions of the membrane by promoting cell lysis and by inhibiting nutrients transport (Eldin et al., 2008, Gu et al., 2007, Du et al., 2009). Figure 4-4 also shows that S. aureus have a comparatively greater resistance to CMCS than E. coli. This may be explained by the differences in cell wall structure and composition between those two types of bacteria. In addition, zeta potential measurements of bacterial suspensions (106 cells/mL in PBS (10 mM, pH = 7.4)) indicate that the negative charge on the cell surface of E. coli (zeta potential of -32.3 ± 1.1) is higher than on S. aureus (zeta potential of -22.5 ± 3.8), and thus the interaction of CMCS with E. coli is probably stronger than with S. aureus. 4.3 Biofilms Disruption The results in the preceding section showed that CMCS and CMCS-MNPs can effectively eliminate both S. aureus and E. coli bacteria in suspension. However, antibacterial assays against planktonic bacteria may not give a representative indication of the efficacy against bacteria in biofilms since the dense and protected environment of the biofilms may shield the bacteria from antimicrobial agents. Thus, the antibacterial properties of CMCS-MNPs on S. aureus and E. coli biofilms were also investigated. Figure 4-6 shows the viable cell count in S. aureus and E. coli biofilms after treatment with CMCS and CMCS-MNPs for only 5 min. As shown in 38 Chapter 4 Results and Discussions Figure 4-6a, in the control experiment (pre-grown biofilms in sterile PBS without addition of CMCS or CMCS-MNPs), there is a decrease in the number of viable cells in the S. aureus biofilm throughout the 48 h incubation period. This finding was consistent with the previous study by Fujimoto et al. (2006) that S. aureus required many macronutrients and micronutrients to grow. Due to a lack of such nutrients in PBS used in the experiments, the S. aureus cells in the biofilm gradually died during the incubation period. The S. aureus biofilms treated with CMCS (0.34 mg/mL) and CMCS-MNPs (1.0 mg/mL and 2.0 mg/mL) in the absence of MF did not exhibit significant differences in viable bacterial cell count as compared to the control (p > 0.05), throughout the 48 h incubation period. This result illustrates the difficulty for antibacterial agents to penetrate the biofilm. However, there was a clear decrease in viable bacterial cells within the biofilms when a MF was applied below the biofilm-containing wells in the presence of CMCS-MNPs. The difference in the results obtained in the absence and presence of MF indicates that penetration of the CMCS-MNPs into the biofilms is essential for biofilm disruption. This can be confirmed by the use of red fluorescent RITC-CMCS-MNPs to study the penetration of nanoparticles into the biofilms in the presence of MF. Figure 4-7 shows the CLSM images of S. aureus biofilms (with viable bacterial cells stained green, RITC-CMCS-MNPs stained red, and the yellow signal is a combination of the green and red signals) treated with RITC-CMCS-MNPs for 5 min with or without a MF. In the absence of MF, a few nanoparticles have settled on the biofilms (Figure 4-7a and 4-7b) within the 5 min exposure period. However, in the presence of MF, the strong 39 Chapter 4 Results and Discussions yellow signal throughout the biofilms confirms that the RITC-CMCS-MNPs rapidly deposited on and penetrated into the biofilms (Figure 4-7c and 4-7d) within 5 min. It can also be observed from Figure 4-6a that a higher concentration of CMCS-MNPs generally resulted in greater disruption of the biofilms. For instance, after 24 h incubation in PBS, the number of viable cells in the biofilm treated with 1.0 mg/mL CMCS-MNPs under MF decreased by 54% compared to the control experiment, while the corresponding value when 2.0 mg/mL CMCS-MNPs were used in the presence of MF was 79%. 40 Chapter 4 Results and Discussions a No. of Bacterial Cells (106/cm2) 700 600 Control 1.0 CMCS-MNPs 1.0 CMCS-MNPs (MF) CMCS 2.0 CMCS-MNPs 2.0 CMCS-MNPs (MF) 500 400 * 300 * * 200 * 100 0 ** 12 h 24 h 48 h No. of Bacterial Cells (106/cm2) b 140 120 Control 1.0 CMCS-MNPs 1.0 CMCS-MNPs (MF) CMCS 2.0 CMCS-MNPs 2.0 CMCS-MNPs (MF) 100 80 60 * 40 0 * * 20 12 h * * 24 h * 48 h Figure 4-6 Effect of CMCS-MNPs (with or without MF) and CMCS on pre-grown (a) S. aureus biofilms and (b) E. coli biofilms after 12, 24, and 48 h. The controls refer to the respective pre-grown biofilms in sterile PBS without addition of CMCS or CMCS-MNPs. 41 Chapter 4 Results and Discussions The prefix 1.0 and 2.0 represent 1.0 mg/mL and 2.0 mg/mL CMCS-MNPs suspension respectively; and the suffix (MF) indicates the application of magnetic field in the 5 min period when the biofilms were exposed to the CMCS-MNPs suspension. * denotes significant differences (p < 0.05) compared to the control experiment at the same incubation time. a c b d Figure 4-7 CLSM (a,c) volume view and (b,d) cross-sectional view images of S. aureus biofilms exposed to RITC-CMCS-MNPs (2.0 mg/mL) (a,b) without a MF and (c,d) with a MF. Scale bar = 100 µm. Viable bacterial cells are stained green, RITC-CMCS-MNPs are stained red, and the yellow signal arises from a combination of the green and red signals. The effect of CMCS and CMCS-MNPs on E. coli biofilms in the absence or presence of MF is presented in Figure 4-6b. Unlike the S. aureus biofilms (Figure 4-6a), the E. 42 Chapter 4 Results and Discussions coli biofilms in the control experiment did not exhibit a reduction in viable cell number. Young et al. (1997) had reported that growth of E. coli cells was observed even after 6 months in intraluminal saline. Similar to the results shown in Figure 6a, CMCS and CMCS-MNPs in the absence of MF are not effective in disrupting the biofilm over 48 h. However, when the E. coli biofilms were treated with 1.0 mg/mL and 2.0 mg/mL CMCS-MNPs in the presence of MF, the viable cell count decreased by nearly 60% and 83% after 12 h compared to the control experiment. Furthermore, the number of viable cells in the biofilms decreased significantly with time and more than 85% and 95% of the cells in biofilms treated with 1.0 mg/mL and 2.0 mg/mL CMCS-MNPs under MF, respectively, were killed after 48 h. CLSM and SEM were used to provide a more illustrative description of the biofilm disruption capabilities of CMCS-MNPs under MF. Figure 4-8 shows the CLSM images (volume view) of S. aureus and E. coli biofilms (with viable cells stained green) after exposure to CMCS-MNPs with or without a MF, following by incubation for 24 h in PBS. As can be seen from Figure 4-8a and 4-8d, S. aureus produced thicker and denser biofilms than E. coli. This finding is consistent with the SEM images in Figure 4-9a and 4-9d. After 24 h, a large portion of the original S. aureus and E. coli biofilms treated with 2.0 mg/mL CMCS-MNPs with MF have been disrupted (Figure 4-8c and 4-8f, Figure 4-9c and 4-9f). It can be observed from Figure 4-9c and 4-9f that there is a substantial amount of debris on the PS substrate. This debris is attributed to a mixture of the CMCS-MNPs, cell and biofilm fragments, and 43 Chapter 4 Results and Discussions it can be removed by using a magnet during the rinsing process (data not shown). The reduction in the biofilm mass after treatment with CMCS-MNPs in the absence of MF (Figure 4-8b and 4-8e, Figure 4-9b and 4-9e) is very much less than that with MF, consistent with the quantitative viable bacterial cell count results in Figure 4-6. The results in Figure 4-8 and 4-9 clearly indicate that application of CMCS-MNPs with a MF is effective in disrupting the biofilms although the percentage decrease in bacterial cell count is not as high as that observed in the planktonic cells experiments (Figure 4-4). In the experiments with planktonic cells, the direct contact between the nanoparticles and the bacterial cells was enhanced by the continuous agitation provided by the orbital shaker, and enhanced contact would facilitate the membrane-disruptive effect of the CMCS-MNPs. On the other hand, in the biofilms experiments, the nanoparticles were basically effective against the bacterial cells in their immediate vicinity. A number of bacterial cells may not be in contact with the nanoparticles despite the application of MF, and a higher nanoparticle concentration will increase the probability of contact. In addition, bacteria in biofilms are more resistant to antimicrobial agents than their planktonic counterparts as mentioned above, possibly due to different growth characteristics (Dusane et al., 2008) and alterations in the membrane protein composition of the biofilm cells (Otto et al., 2001). 44 Chapter 4 Results and Discussions a d b e c f Figure 4-8 CLSM volume view images of (a-c) E. coli biofilms and (d-f) S. aureus biofilms: (a) and (d) pre-grown biofilms after incubation in PBS for 24 h, (b) and (e) with addition of CMCS-MNPs (2.0 mg/mL) without MF for 5 min and after incubation in PBS for 24 h, (c) and (f) with addition of CMCS-MNPs (2.0 mg/mL) with MF for 5 min and after incubation in PBS for 24 h. Scale bar = 100 µm. 45 Chapter 4 Results and Discussions a d b e c f 1 Figure 4-9 SEM images of (a-c) E. coli biofilms and (d-f) S. aureus biofilms: (a) and (d) pre-grown biofilms after incubation in PBS for 24 h, (b) and (e) with addition of CMCS-MNPs (2.0 mg/mL) without a MF for 5 min and after incubation in PBS for 24 h, (c) and (f) with addition of CMCS-MNPs (2.0 mg/mL) with MF for 5 min and after incubation in PBS for 24 h 46 Chapter 4 Results and Discussions 4.4 Cytotoxicity of Nanoparticles While the above results have clearly illustrated the antibacterial efficacy of the CMCS-MNPs, its potential cytotoxic effects on mammalian cells must be considered. The MTT assay results after incubation of 3T3 fibroblast cells with MNPs (2.0 mg/ml), CMCS (0.34 mg/mL) and CMCS-MNPs (2.0 mg/mL, with and without MF) for 24 h are shown in Figure 4-9. The viability of the fibroblasts in the presence of MNPs, CMCS and CMCS-MNPs remained high (96 – 98% as compared to the control) regardless of the presence of MF. The lack of cytotoxicity of the CMCS and CMCS-MNPs are as expected since Jaiswal et al. (2012) had reported that more than 95% of both U-87 MG (human glioblastoma astrocytoma) and HT29 (human colon adenocarcinoma) cells were viable after 24 h treatment with folic acid conjugated chitosan nanocarriers over a wide range of concentration from 0 to 30 µg/mL. Another investigation showed that polyethylene glycol (PEG) coated MNPs were nontoxic to infinity telomerase-immortalized primary human fibroblasts (Gupta et al., 2004). In addition, Milovic et al. (2005) had reported that polycationic polyethylenimine immobilized on a glass slide can effectively kill E. coli cells by a similar membrane-rupturing mechanism as exhibited by chitosan without adverse effects on mammalian cells. Though mammalian cell membranes are also negatively charged (Mishra et al., 2009), contact with CMCS did not result in the membrane-disruptive action observed with bacteria. This phenomenon may be attributed to the differences in size and membrane composition between mammalian cells and bacterial cells. It was found that the presence of cholesterols (not found in 47 Chapter 4 bacterial Results and Discussions cell membranes) in mammalian cell membranes inhibited the membrane-rupturing ability of the cationic antimicrobial pardaxin, though the inhibition mechanism was not well understood (Hallock et al., 2002). Therefore, it was possible that the same mechanism also applied when mammalian cells were exposed to CMCS. 100 80 60 40 20 N (M Ps S C C M C C S- M M C N S- Ps M M C N M F) 0 Ps Cell Viability (% of Control) 120 Figure 4-10 Viability of 3T3 fibroblast cells incubated for 24 h in growth medium containing MNPs (2.0 mg/ml), CMCS (0.34 mg/ml) and CMCS-MNPs (2.0 mg/ml) relative to the control (i. e. no CMCS or CMCS-MNPs added). The suffix (MF) indicates the application of magnetic field throughout the incubation period. Results are represented as mean ± standard deviation 48 CHAPTER 5 CONCLUSION AND RECOMMENDATIONS 49 Chapter 5 Conclusion and recommendations 5.1 Conclusion Bacteria in biofilms develop resistance to antibiotics via a combination of mechanisms. In the present work, a non-antibiotic-based strategy of using CMCS-MNPs under an external magnetic field was shown to exhibit strong bactericidal activities against biofilms. These nanoparticles are produced from readily available chemicals and the process is easily scalable. The bactericidal effects arise from the CMCS component on the surface of CMCS-MNPs, while the MNPs acting in conjunction with the magnetic field facilitate the penetration of the bactericidal agent deep into the biofilms. The antibacterial efficiency is dependent on the concentration of CMCS-MNPs and the incubation time. The number of viable cells in S. aureus and E. coli biofilms after exposure to 2.0 mg/mL CMCS-MNPs under a magnetic field decreased by 84% and 95%, respectively, after 48 h. In addition, CMCS-MNPs are also highly effective against planktonic S. aureus and E. coli cells, and more than 99% of the cells in contact with these nanoparticles are killed after 10 h and 5 h, respectively. CMCS-MNPs are not cytotoxic to mammalian cells, and can potentially be used as an antimicrobial agent in a wide range of applications including targeting biofilms associated with industrial equipment, biomedical devices or in food processing. 50 Chapter 5 Conclusion and recommendations 5.2 Recommendations A number of possible methods which may enhance the dispersion and the antibacterial efficacy of the functional magnetic nanoparticles can be foreseen. Two examples which need further study are given blow: Enhance the dispersion of the functional MNPs In this thesis, the carboxymethylation of chitosan has increased its solubility in water, and promoted the dispersion of CMCS-MNPs in aqueous media. Recent studies have showed that the chitosan functionalized by other groups such as succinyl and dicarboxymethyl possessed a better solubility in water (Jayakumar et al., 2010). It can be expected that the dispersion of functional MNPs may be further enhanced by using these functional groups in chitosan. Enhance the antibacterial efficacy of the functional MNPs Through the electrostatic interaction of the positively charged amino groups in chitosan with the negatively charged cell membrane, chitosan can disrupt the normal functions of bacterial membrane by promoting cell lysis and by inhibiting nutrients transport. Electrostatic interaction plays an important role in the antibacterial process. The results in Section 4.3 showed that despite exposure to CMCS-MNPs under MF, a number of bacterial cells in biofilms remained unaffected by the magnetic nanoparticles. This may due to the poor attachment between cell membrane and magnetic nanoparticles. Some carbohydrates such as glucose, mannose has been 51 Chapter 5 Conclusion and recommendations reported to attach well to the bacterial membrane (Ip et al., 2009, Eboigbodin et al., 2007). 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Polymer 2011, 52, 2141. 65 [...]... images of (a-c) E coli biofilms and (d-f) S aureus biofilms: (a) and (d) pre-grown biofilms after incubation in PBS for 24 h, (b) and (e) with addition of CMCS-MNPs (2.0 mg/mL) without a MF for 5 min and after incubation in PBS for 24 h, (c) and (f) with addition of CMCS-MNPs (2.0 mg/mL) with MF for 5 min and after incubation in PBS for 24 h Figure 4-10 Viability of 3T3 fibroblast cells incubated for. .. decreased the fluorescence of otopathogenic Pseudomonas aeruginosa (OPPA8) biofilms after 24 h of exposure (p < 0.03) (Eric et al., 2008) In other investigations, MgF2 nanoparticles were shown to be capable of penetrating both Escherichia coli (E coli) and S aureus cells, and could restrict the formation of biofilms (Lellouche et al., 2009) Ag-loaded 3 Chapter 1 Introduction chitosan nanoparticles also show... biofilms have shown that some antibiotics cannot readily permeate biofilms (Stewart et al., 1996) Furthermore, the exchange of genetic materials and the mutation of bacteria in biofilms occur more frequently than in planktonic populations Therefore, development of resistance mechanisms can quickly be selected for and propagated throughout the community In addition, the cells in the deep layers of biofilms. .. Disruption of biofilm Much of work has been done with the purpose of disrupting the biofilms: (1) Laser and photodynamic treatment have been used to disrupt bacterial biofilms Krespi et al (2011) reported that the combination of Q-switched Nd-YAGSW (SW) and NIR diode (NIR) lasers can result in a decrease of more than 43% of methicillin-resistant S aureus biofilm cells However, the need for specialized... would be desirable for combating biofilm-associated infections Lasers have been used for disrupting biofilms in recent years (Krespi et al., 2008) For instance, the combination of Q-switched Nd-YAGSW (SW) and NIR diode (NIR) lasers can result in a decrease of more than 43% of methicillin-resistant Staphylococcus aureus (S aureus) biofilm cells (Krespi et al 2011) However, the need for specialized equipment... properties of chitosan such as the degree of N-deacetylation, molecular weight and solubility can, and to a great extent, influence the antibacterial efficacy One of the most important parameter to examine closely is the degree of deacetylation of chitin Takahashi et al (2008) reported that the higher degree of deacetylation, the higher antibacterial efficacy of chitosan against S aureus and E coli bacteria... Gram-negative E coli The use of a magnetic field in combination with the CMCS-MNPs can also effectively disrupt the biofilms of these bacteria 4 CHAPTER 2 LITERATURE REVIEW 5 Chapter 2 Literature Review 2.1 Biofilm A biofilm is a gathering of bacterial cells enclosed in a self-produced polymeric matrix composed of extracellular polymeric substances, mainly exopolysaccharides, proteins and nucleic acids Biofilms. .. cells were detected (Eric et al., 2008) FAC at 200 µM caused disruption of PA biofilms after a 5-day incubation period (Musk et al., 2005) (3) Lellouche et al (2009) demonstrated that nanosized magnesium fluoride (MgF2) was capable of penetrating E coli and S aureus cells and inhibiting biofilm formation (4) Magnetic microspheres coated with Ag nanoparticles- loaded multilayers were also shown to possess... Staphylococcus epidermidis and Gram-negative E coli bacteria (Lee et al., 2005) Nevertheless, the use of MgF2 and Ag may not be appropriate as they pose possible environmental problems and toxicity to certain mammalian cells (Mukherjee et al., 2012, Kim et al., 2011) In the present work, magnetic iron oxide nanoparticles (MNPs) functionalized with bactericidal moieties are used for disruption of biofilms MNPs are... slow or incomplete penetration of the antibiotics into the biofilms Measurements of antibiotics penetration into biofilms in vitro have shown that some antibiotics readily permeate bacterial biofilms (Stewart et al., 1996) However, some antibiotics are adsorbed on the biofilm matrix which can reduce its penetration into the biofilms This may account for the slow penetration of aminoglycoside antibiotics ... (a-c) E coli biofilms and (d-f) S aureus biofilms: (a) and (d) pre-grown biofilms after incubation in PBS for 24 h, (b) and (e) with addition of CMCS-MNPs (2.0 mg/mL) without MF for and after... without MF) and CMCS on pre-grown (a) S aureus biofilms and (b) E coli biofilms after 12, 24, and 48 h The controls refer to the respective pre-grown biofilms in sterile PBS without addition of CMCS... (d-f) S aureus biofilms: (a) and (d) pre-grown biofilms after incubation in PBS for 24 h, (b) and (e) with addition of CMCS-MNPs (2.0 mg/mL) without a MF for and after incubation in PBS for 24