The Bioeffects of a High Frequency Electromagnetic Field in the Microwave Range

The bioeffects of a high frequency electromagnetic field in the microwave range

Submitted in total fulfilment of the requirements for the degree of

Doctor of Philosophy By The Hong Phong Nguyen

Faculty of Science, Engineering and Technology Swinburne University of Technology 2018

ABSTRACT

The World Health Organization (WHO) has stated that there is no evidence available to confirm the existence of any health consequences arising from human exposure to low level EMFs. Some gaps in knowledge exist regarding the biological effects that may be triggered by exposure to high levels of EMFs and hence these possible effects require further research. Notwithstanding the sterilization/inactivation effects that come about from EMF exposure, a number of specific effects have been observed that cannot be explained by the increase in bulk temperature that occurs through EMF exposure alone. The exact mechanism/s of those effects is/are not fully understood and therefore have been the subject of debate.

In this thesis, the effects of high frequency (18 GHz) EMF exposure have been studied using typical representatives of prokaryotic and eukaryotic taxa, including two Gram-negative bacteria (Branhamella catarrhalis ATCC 23246 and Escherichia coli ATCC 15034), six Gram-positive bacteria (Kocuria rosea CIP 71.15T, Planococcus maritimus KMM 3738, Staphylococcus aureus CIP 65.8T, Staphylococcus aureus ATCC 25923, Staphylococcus epidermidis ATCC 14990T, and Streptomyces griseus ATCC 23915), a eukaryotic unicellular organism (yeast Saccharomyces cerevisiae ATCC 287) and red blood cells obtained from a New Zealand rabbit. Three parameters of the EMF exposure (power, duration, and exposure number) were varied to determine their effect on the biological system being studied. Advanced microscopy techniques were employed, comprising Scanning Electron Microscopy (SEM), Transmission Electron Microscopy (TEM), and Confocal Scanning Laser Microscopy (CSLM) together with fluorescent probes, in order to allow a thorough examination of the cell membrane morphology and permeability following EMF exposure/s to be studied.

It was determined, for the first time, that regardless of the differences in cell wall/ membrane structure, exposure to 18 GHz EMF induced cell permeabilization, as confirmed via the ability of the cells to uptake silica nanospheres (23 nm and 46 nm in diameter), in all the cell types studied, as confirmed directly by transmission electron microscopy (TEM) and indirectly by confocal laser scanning microscopy (CLSM). A dosimetry analysis revealed that the EMF exposure required to induce cell permeation such that the membrane was able to uptake 23 nm and 46 nm nanospheres was between three and six EMF doses with a specific absorption rate (SAR) of 5 kW/kg and 3 kW/kg per exposure, respectively, depending on the cell types being studied. This specific EMF bioeffects could not be duplicated using conventional heating methods under similar temperature conditions. The cells remained viable (85%) and permeable for at least nine minutes after EMF exposures. It is suggested that the taxonomic affiliation and cell wall/membrane structures (e.g., the presence of peptidoglycan layer, mannoprotein/β-glucan layer, phosphatidyl-glycerol and/or pentadecanoic fatty acid) may affect the extent of permeabilization to allow the uptake of 46 nm nanopsheres.

A new mechanism of cell permeability using 18 GHz EMF is postulated; EMF exposure disturbs the stability of the charged lipid bilayers within the cell membrane, thus affecting its fluidity. This mechanical stimulation, in turn, alters the tension of the membrane, causing it to deform, which results in an enhanced opportunity for transport through the membrane, which occurs via a quasi- exocytosis/endocytosis process.

ACKNOWLEDGMENTS

First and foremost, I would like to express my deepest and most sincere gratitude to my mentors and supervisors: Professor Elena Ivanova for her infinite support and encouragement in conducting research as well as writing for publications; Professor Russell J. Crawford for his supervision and guidance throughout my candidature. Without their support, none of this would have been possible and for which I am extremely grateful.

Secondly, I would like to thank Professor Rodney J. Croft from University of Wollongong, Dr. Vladimir A. Baulin from Universitat Rovira I Virgili, Professor Andrew W. Wood and Dr. Robert L. McIntosh from Swinburne University of Technology for their guidance and useful advice in this research.

I would like to sincerely thank my parents, siblings, relatives and friends for their perpetual love and unconditioned support during my PhD.

Finally, I would also like to extend my gratitude to Dr. Vi Khanh Truong, Dr. Yury Shamis, Dr. Hayden K. Webb, Dr. Jafar Hasan, Dr. Thi Song Ha Nguyen, Dr. Thi Hong Vy Pham for their immense academic and emotional support throughout my research.

Dr. James Wang from Faculty of Science, Engineering and Technology, Swinburne University of Technology assisted me in the use of Scanning Electron Microscope.

Dr. Igor Sbarski from Faculty of Science, Engineering and Technology, Swinburne University of Technology provided me technical support in the use of Peltier plate.

Mr. Phil Francis, Dr. Matthew Field, and Dr. Chaitali Dekiwadia from the RMIT Microscopy and Microanalysis Facility, RMIT University assisted me in the use of Transmission Electron Microscope and sample preparations.

Associate Professor Brian Phillips from Faculty of Health, Arts and Design, Swinburne University of Technology provided me valuable statistics advice and support in the use of SPSS software. DECLARATION

I, The Hong Phong Nguyen, declare that this thesis is my original work and contains no material which has been submitted for the award of any other degree or diploma from another university.

To the best of my knowledge, I certify that this thesis contains no material previously published or written by another person except where due reference has been made. Wherever contributions of others were involved, every effort has been made to acknowledge contribution of the respective workers or authors.

The Hong Phong Nguyen

LIST OF PUBLICATIONS

Peer-reviewed articles

Nguyen, T. H. P., Pham, T. H. V., Baulin, V., Croft, R. J., Crawford, R. J. & Ivanova, E. P. (2017) The effect of a high frequency electromagnetic field in the microwave range on red blood cells. Sci Rep 7, 10798.

Nguyen, T. H. P., Pham, T. H. V., Nguyen, S. H., Baulin, V., Croft, R. J., Phillips, B., Crawford, R. J. & Ivanova, E. P. (2016) The bioeffects resulting from prokaryotic cells and yeast being exposed to an 18 GHz electromagnetic field. PLoS ONE 11, e0158135.

Nguyen, T. H. P., Shamis, Y., Croft, R. J., Wood, A., McIntosh, R. L., Crawford, R. J. & Ivanova, E. P. (2015) 18 GHz electromagnetic field induces permeability of Gram-positive cocci. Sci Rep 5, 10980.

Pogodin, S., Hasan, J., Baulin, V. A., Webb, H. K., Truong, V. K., Nguyen, T. H. P., Boshkovikj, V., Fluke, C. J., Watson, G. S., Watson, J. A., Crawford, R. J. & Ivanova, E. P. (2013) Biophysical model of bacterial cell interactions with nanopatterned cicada wing surfaces. Biophys J 104, 835-840.

Conference poster presentations with published abstracts

Nguyen, T. H. P., Perera, P. G. T., Pham, T. H. V., Croft, R., Crawford, R. J. & Ivanova, E. P. Super High Frequency (18 GHz) electromagnetic field induced membrane permeability on erythrocytes and PC-12 neuronal cells. Science and wireless 2016, 2016 Melbourne, Australia.

Nguyen, T. H. P., Shamis, Y., Croft, R., Crawford, R. J. & Ivanova, E. P. Specific electromagnetic effects of microwave radiation on bacteria. 34th Annual Meeting of the Bioelectromagnetics Society, 2012 Brisbane, Australia.

TABLE OF CONTENTS

ABSTRACT ...... 2

ACKNOWLEDGMENTS ...... 4

DECLARATION ...... 6

LIST OF PUBLICATIONS ...... 7

LIST OF TABLES ...... 12

LIST OF FIGURES ...... 13

Chapter 1. Introduction ...... 22 1.1. Overview ...... 23 1.2. The aims ...... 25 Chapter 2. Literature review ...... 27 2.1. Overview ...... 28 2.2. The nature of electromagnetic fields (EMFs) ...... 28 2.2.1. Electromagnetic spectrum ...... 28 2.2.2. Electromagnetic fields ...... 29 2.2.3. Electromagnetic phenomena ...... 33 2.2.4. Dosimetry ...... 34 2.2.5. Operational health and safety standards ...... 35 2.2.6. Current understanding of the biological effects induced by microwave range electromagnetic fields ...... 36 2.2.6.1. Biological effects of microwave range electromagnetic fields at cellular level ...... 37 2.2.6.2. The combined biological effects resulting from exposure to microwave range electromagnetic fields and other agents ...... 43 2.2.6.3. Electromagnetic field effects on cell membranes resulting from exposure to microwave range radiation ...... 44 2.2.6.5. Proposed mechanisms responsible for the observed biological effects in cells resulting from exposure to microwave range EMF radiation . 44 2.3. Cell membrane permeability and drug delivery systems ...... 48 2.3.1. Mechanical stress ...... 49 2.3.2. Photoporation ...... 50 2.3.3. Sonoporation ...... 52 2.3.4. Electroporation ...... 54 2.4. Functional roles of lipids in membranes ...... 56 Chapter 3. Experimental Design ...... 60 3.1. Overview ...... 61 3.2. Experimental design...... 61 3.2.1. Temperature-dependent approach ...... 61 3.2.2. Power-variation approach ...... 61 3.3. Instrument set-up and dosimetry ...... 62 3.4. Heat treated controls ...... 67 3.5. Negative controls ...... 68 Chapter 4. Materials and Methods...... 69 4.1. Overview ...... 70 4.2. Sample preparation ...... 70 4.2.1 Bacterial and yeast strains, growth and maintenance ...... 70 4.2.2. Selected cell types ...... 72 4.2.2.1. Gram negative bacteria ...... 72 4.2.2.1.1. Branhamella catarrhalis ATCC 23246 ...... 72 4.2.2.1.2. Escherichia coli K 12 ...... 73 4.2.2.2. Gram positive bacteria ...... 73 4.2.2.2.1. Kocuria rosea CIP 71.15T ...... 73 4.2.2.2.2. Planococcus maritimus KMM 3738 ...... 74 4.2.2.2.3. Staphylococcus aureus ATCC 25923 and CIP 65.8T ...... 75 4.2.2.2.4. Staphylococcus epidermidis ATCC 14990T ...... 75 4.2.2.2.5. Streptomyces griseus ATCC 23915 ...... 76 4.2.2.3. Unicellular eukaryote ...... 77 4.2.2.3.1. Saccharomyces cerevisiae ATCC 287 ...... 77 4.2.2.4. Red blood cells ...... 78 4.3. Sample preparation ...... 78 4.4. Bacterial cell viability ...... 79 4.5. Scanning electron microscopy ...... 80 4.5.1. Sample preparation for analysis of cell morphology ...... 80 4.5.2. Sample preparation for cell rigidity analysis ...... 80 4.6. Confocal laser scanning microscopy ...... 81 4.7. Quantification of intake nanospheres ...... 83 4.7.1. Experimental procedure ...... 83 4.7.2. Theoretical consideration ...... 83 4.8. Transmission electron microscopy ...... 84 4.8.1. Grids and samples preparation ...... 84 4.8.1.1. Bacteria and yeast ...... 84 4.8.1.2. Erythrocytes ...... 85 4.8.1.3. Sectioning and examination ...... 86 4.9. Statistical analysis ...... 86 Chapter 5. EMF effects on Gram negative bacterial cells ...... 87 5.1. Overview ...... 88 5.2. Background ...... 88 5.3. Results ...... 90 5.3.1. EMF effects on cell membrane permeability ...... 90 5.3.2. EMF effects on cell morphology ...... 97 5.3.3. EMF effects on cell viability ...... 98 5.4. Discussion ...... 99 Chapter 6. EMF effects on Gram positive bacterial cells...... 101 6.1. Overview ...... 102 6.2. Background ...... 102 6.3. Results ...... 106 6.3.1. EMF effects on cell membrane permeability ...... 106 6.3.2. The effect of EMF exposure on cell morphology ...... 116 6.3.3. EMF effects on cell viability ...... 118 6.3.4. Confirmation of change in bacterial metabolic status ...... 120 6.3.5. Effects of multiple 18 GHz EMF exposures on Staphylococcus aureus strains 120 6.4. Discussion ...... 123 Chapter 7. EMF effects on the eukaryotic unicellular yeast Saccharomyces cerevisiae ...... 127 7.1. Overview ...... 128 7.2. Background ...... 128 7.3. Results and discussion ...... 130 7.3.1. EMF effects on cell viability ...... 130 7.3.2. EMF effects on cell membrane permeability ...... 132 7.3.3. EMF effects on cell morphology ...... 136 7.4. Further direction...... 137 Chapter 8. The effects of EMF exposure on red blood cells ...... 138 8.1. Overview ...... 139 8.2. Background ...... 139 8.3. Results and discussion ...... 140 8.3.1. EMF effects on cell morphology ...... 140 8.3.2. EMF effects on cell membrane permeability ...... 142 Chapter 9. General discussion ...... 150 9.1. Overview ...... 151 9.2. Bioeffects: 18 GHz EMF dosimetry requirements ...... 151 9.2.1. Temperature-dependent approach ...... 151 9.2.2. Power variation ...... 152 9.3. Proposed mechanism/s of the bioeffects arising from exposing cells to an 18 GHz EMF ...... 152 9.4. The potential applications of 18 GHz EMF exposures in drug delivery and gene therapy applications ...... 155 Chapter 10. Conclusions and Future Directions ...... 158 10.1. Conclusions ...... 159 10.2. Future directions ...... 160 References ...... 162

LIST OF TABLES

Table 2.1. A summary of the non-thermal biological effects of EMF in the microwave range...... 39 Table 4.1. Taxonomic affiliation of bacterial and yeast strains employed in this study...... 71 Table 6.1. Internalization of silica nanospheres by bacterial and yeast cells after EMF exposure...... 114 Table 6.2. Phospholipids compositions (%) of cell membranes in 18 GHz EMF exposure studies ...... 125 Table 6.3. Chain melting temperature from rippled lamellar gel to fluid lamellar phase of phospholipid chains in studied cell membranes...... 126 Table 8.1. Internalization of silica nanospheres by RBCs subjected to EMF irradiation...... 145

LIST OF FIGURES

Figure 1.1. Flow chart presenting the structure of the study...... 26 Figure 2.1. The electromagnetic spectrum...... 29 Figure 2.2. The molecular structure of the glycerol-based phospholipids headgroups...... 58 Figure 3.1. EMF apparatus. (a) The Vari-Wave Model LT 1500 (Lambda Technologies) EMF apparatus; (b) The EMF chamber containing the temperature control optic probes and (c) schematic diagram of the LT 1500 system...... 63 Figure 3.2. Modelling of the electric field and absorbed power using CST Microwave Studio 3D Electromagnetic Simulation Software. The EMF was uniformly distributed in the sample...... 64 Figure 3.3. Temperature control during EMF exposures. (a) overview of the LFOTU, micro Petri dish and the cell suspension layer; (b) Top view of the Petri dish with five different positions of measurement in the LFOTU; (c) Side view of the cell suspension, showing the three measured depth-levels of the LFOTU...... 66 Figure 3.4. The bulk temperature profiles of EMF exposed and Peltier plate heat-treated cell suspensions...... 67 Figure 3.4. Peltier heating stage (diameter = 65 mm, suspension depth = 0.6 mm) with position used for applying the cell suspension...... 68 Figure 4.1. Structural physiology of the forewing of Psaltoda claripennis, with major areas indicated. The cutting areas for experiment were all the M areas and C areas...... 81 Figure 5.1. Internalization of propidium iodide by Gram-negative bacterial cells after EMF exposure. Row 1: CLSM images showing the internalization of propidium internalization 1 min after EMF exposure. Row 3: no propidium iodide was observed to have been internalized in any tested cell types after 10 min. Row 2 and 4: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all fluorescence images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg- 1...... 91 Figure 5.2. Internalization of propidium iodide by the heat control groups. Row 1: CLSM images showing that no propidium iodide had been internalized by the Peltier heat treated cells (40°C). Row 3: internalization of propidium iodide was observed in the heat inactivated cells (boiling, 100°C). Row 2 and 4: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all fluorescence images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1...... 92 Figure 5.3. No propidium iodide internalization by the untreated control. Row 1: CLSM images showing that no propidium iodide was internalized by untreated bacterial cells. Row 2: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all fluorescence images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg- 1...... 93 Figure 5.4. Internalization of 23.5 nm and 46.3 nm nanospheres by the Gram-negative bacterial cells after EMF exposure. Row 1: CLSM images showing the internalization of 23.5 nm nanospheres (in green) by the EMF exposed cells. Row 2: internalization of 46.3 nm nanospheres was observed in the EMF- exposed cells. Row 3: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all fluorescence images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1...... 94 Figure 5.5. Internalisation of 23.5 nm and 46.3 nm nanospheres by the untreated and heat-treated controls. Row 1: CLSM images showing no internalization of either the 23.5 or 46.3 nm nanospheres by the control cells. Row 2: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all fluorescence images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1...... 96 Figure 5.6. Internalization of 23.5 nm nanospheres by Gram-negative Branhamella catarrhalis cells following EMF exposure. Scale bars in electron micrographs are 200 nm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1...... 97 Figure 5.7. Morphology of EMF-exposed, heat-treated and untreated Gram-negative Branhamella catarrhalis cells. Scale bars in all scanning electron micrographs are 1 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1...... 98 Figure 5.8. Effect of EMF exposure and bulk heat treatment on the viability of Gram-negative bacterial cells. Cells inactivated by boiling (100°C) were found to be non-viable. Data presented are the mean ± standard deviation and representative of 3 independent experiments with 10 replicates each. *p < 0.05 versus the corresponding controls. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1...... 99 Figure 6.1. The structure of teichoic acids on bacterial cell membrane. (a) Structure of glycerol teichoic acid, and (b) Structure of ribitol teichoic acid.104 Figure 6.2. Internalization of propidium iodide internalization by the Gram-positive bacterial cells after EMF exposure. Row 1: CLSM images showing the internalization of propidium iodide when placed in contact with the bacterial cells 1 min following EMF exposure. Row 3: no propidium iodide was observed in any tested cell types when exposed to the cells 10 min after irradiation. Row 2 and 4: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1...... 107 Figure 6.3. Internalization of propidium iodide by the heat control groups. Row 1: CLSM images showing that no propidium iodide was able to be internalized by the bacterial cells 1 min following heating using a Peltier plate (40°C). Row 3: Propidium iodide internalization was found to have occurred in the heat inactivated cells (boiling, 100°C). Row 2 and 4: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg- 1...... 108 Figure 6.4. Internalization of propidium iodide by untreated control samples. Row 1: CLSM images showing that no propidium iodide was able to be taken up by cells that were not exposed to EMF. Row 2: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg- 1...... 109 Figure 6.5. Uptake of 23.5 nm and 46.3 nm silica nanospheres by the Gram-positive bacterial cells after EMF exposure. Row 1: CLSM images showing the uptake of 23.5 nm nanospheres (in green) by the EMF exposed cells. Row 2: uptake of the 46.3 nm nanospheres by the EMF exposed cells. Row 3: phase contrast micrographs showing the bacterial cells in the same field of view. Scale bars in all images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1...... 111 Figure 6.6. Uptake of 23.5 nm and 46.3 nm silica nanospheres by the untreated and heat-treated controls. Row 1 and 3: CLSM images showing negligible internalization of the 23.5 and 46.3 nm nanospheres by the control (heat treated and untreated) cells. Row 2 and 4: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg- 1...... 112 Figure 6.7. Internalization of the 23.5 nm silica nanospheres by Gram- positive bacterial cells following EMF exposure. Row 1: TEM images showing the uptake of 23.5 nm silica nanospheres by the EMF exposed cells. Row 2 and 3: Location of the nanospheres in the control (Peltier plate heated to 40°C and untreated) cells, highlighting that the control samples did not internalize the nanospheres. Scale bars are 200 nm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1...... 115 Figure 6.8. The morphology of EMF exposed, heat-treated and untreated Gram-positive bacterial cells. Row 1: SEM images showing the dehydrated appearance of the surface and traces of leaked cytosolic fluid surrounding the EMF exposed cells. Row 2 and 3: the typical morphology of the control (Peltier plate heated to 40°C and untreated) cells. Scale bars are 400 nm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg- 1...... 117 Figure 6.9. Effect of EMF exposure and bulk heat on the viability of the Gram-positive bacterial cells. Cells inactivated by boiling (100°C) were found to be non-viable. The data presented are the mean ± standard deviation and representative of 3 independent experiments, each comprising 10 replicates each. *p < 0.05 versus the corresponding controls. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1...... 119 Figure 6.10. EMF effect on the cell rigidity of four selected Gram positive bacteria. Row 1: typical scanning electron micrographs of untreated Planococcus maritimus KMM 3738, Staphylococcus aureus ATCC 25923, Staphylococcus aureus CIP 65.8T and Staphylococcus epidermidis ATCC 14990T on wing membrane substrata of the insect Clanger cicada Psaltoda claripennis. Row 2: after EMF exposures, Gram positive cells have disturbed cell membrane rigidity and cellular inner pressure, thus are penetrated by nanopillars. Scale bars are 200 nm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg- 1...... 120 Figure 6.11. The effect of multiple 18 GHz EMF exposures on the morphology and permeability of Staphylococcus aureus cells. Row 1 and 4: typical scanning electron micrographs of S. aureus ATCC 25923 and S. aureus CIP 65.8T cells after multiple 18 GHz EMF exposure. No significant change of cell morphology was observed up to 7th exposure (insets). Scale bars are 10 μm, inset scale bars are 200 nm. Row 2 and 5: CLSM images showing intake of 23.5 nm nanospheres after the 2nd exposure. Row 3 and 6: the phase contrast images in the bottom row show the bacterial cells in the same field of view. Scale bars are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg- 1...... 121 Figure 6.12. The effect of multiple 18 GHz EMF exposures on the viability of Staphylococcus aureus cells. Staphylococcus aureus ATCC 25923 (shaded white) and Staphylococcus aureus CIP 65.8T (dotted grey) cell viability as a function of time and number of EMF exposures. The cell viability of the two Staphylococcus aureus strains is displayed in colony forming units (cfu) per 100 µL. The untreated cells preserved their viability throughout the 54 min period, declining slightly to 97 ± 1% for both strains. The x-axis represents the number of viable cells present (expressed as a percentage) after corresponding EMF exposures and the y-axis represents the number of EMF exposures. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1...... 122 Figure 7.1. Effect of EMF exposure and bulk heat on the Saccharomyces cerevisiae yeast cell viability. Cells inactivated by boiling (100°C) were found to be non-viable. Data are mean values ± standard deviation and are representative of 3 independent experiments, each with 10 replicates. *p < 0.05 versus the corresponding controls. Cell suspensions were subjected to three and six consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1...... 131 Figure 7.2. Internalization of propidium iodide by Saccharomyces cerevisiae yeast cells after EMF exposure. Row 1: CLSM images showing the internalization of propidium iodide following 6 consecutive cycles of EMF exposure. No propidium iodide was observed in any of the yeast cells after 3 consecutive cycles of EMF exposure. The internalization of propidium iodide was observed in the heat inactivated cells (boiling, 100°C). No propidium iodide uptake was observed in the cells subjected to Peltier heat heating to 33°C. CLSM images showing no propidium iodide uptake by the untreated cells. Row 2: phase contrast micrographs showing yeast cells in the same field of view. Scale bars are 5 μm. Cell suspensions were subjected to three and six consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1...... 133 Figure 7.3. Nanospheres (23.5 nm and 46.3 nm) internalization by the Saccharomyces cerevisiae yeast cells after 6 cycles of EMF exposure. Row 1: CLSM images showing 23.5 nm nanospheres (in green) being taken up by the EMF-exposed cells. Row 2: Internalization of the 46.3 nm nanospheres was not observed in the EMF-exposed and control cells. Row 3: Phase contrast micrographs showing the yeast cells in the same field of view. Arrows indicate young yeast cells, which were unable to take up any nanospheres. Scale bars are 5 μm. Cell suspensions were subjected to three and six consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1...... 135 Figure 7.4. Internalization of 23.5 nm nanospheres by the Saccharomyces cerevisiae yeast cells following 6 cycles of EMF exposure. Row 1: TEM images showing the internalization of 23.5 nm nanospheres by the EMF-exposed cells. Row 2: Location of the 23.5 nm nanospheres outside and around the cell membrane of both the untreated and heat-treated cells, showing a uniform cytosol with no nanospheres being present. Scale bars are 200 nm. Cell suspensions were subjected to three and six consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1...... 136 Figure 7.5. The morphology of EMF-exposed, heat-treated and untreated Saccharomyces cerevisiae yeast cells. Column 1: SEM image highlighting that no significant change in cell morphology occurred after exposure to the EMF. Column 2 and 3: Typical cell morphology of the control (heat treated to a temperature of 33°C) and untreated cells. Scale bars are 1 μm. Cell suspensions were subjected to three and six consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1...... 137 Figure 8.1. RBC morphology after 18 GHz EMF exposure. Typical SEM micrographs of rabbit RBCs after 18 GHz EMF radiation exposure, resulting in final temperatures of 33 and 37°C. Approximately 9% (33°C) and 14% (37°C) RBC vesicles (first row) and acanthocytes (first row) were observed. The morphology of the non-treated and control Peltier heat-treated RBCs remained unchanged in their morphology (second and third rows). Scale bars are 10 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 4.1 kW kg- 1 that resulted in temperature increases ranging from 20 to 37ºC (at a heating rate of 17ºC per min). Then the SAR was reduced to approximately 3.0 kW kg-1 which resulted in temperature increases ranging from 20 to 33ºC (at a heating rate of 13ºC per min)...... 141 Figure 8.2. Permeabilization of RBCs resulting from exposure to an 18 GHz EMF. CLSM images show an uptake of 23.5 and 46.3 nm nanospheres (first row). A lipophilic membrane stain, DiI (Life Technologies, Scoresby, VIC, Australia) was used to stain the entire population of RBCs for contrasting purposes (first row). The phase contrast images (second row) show erythrocytes in the same field. Scale bars are 2 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1 that resulted in temperature increases ranging from 20 to 33ºC (at a heating rate of 13ºC per min)...... 142 Figure 8.3. Permeabilization of RBCs resulting from exposure to an 18 GHz EMF. No uptake of 23.5 and 46.3 nm nanospheres by the control groups. CLSM and phase contrast images showing the appearance of the RBCs remained unchanged, and with no internalization of nanospheres. Scale bars are 2 μm (first and second rows). Typical TEM images of ultra-thin (70 nm) cross-sections of RBCs, showing the cell membrane of untreated and heat-treated RBCs with a uniform cytosol without any 46.3 nm nanospheres being present. Scale bars are 1 μm (fourth row). Inset scale bars are 200 nm (third row). Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1 that resulted in temperature increases ranging from 20 to 33ºC (at a heating rate of 13ºC per min).144 Figure 8.4. Quantification of the permeabilized RBCs resulting from 18 GHz EMF exposure. CLSM images show an uptake of 23.5 and 46.3 nm nanospheres (first row). The phase contrast images (second row) show RBCs in the same field. Only the RBCs with circular morphology in the phase contrast images were counted for the quantification. Scale bars are 10 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 4.1 kW kg-1 that resulted in temperature increases ranging from 20 to 37ºC (at a heating rate of 17ºC per min). Then the SAR was reduced to approximately 3.0 kW kg-1 which resulted in temperature increases ranging from 20 to 33ºC (at a heating rate of 13ºC per min).146 Figure 8.5. Internalization of 46.3 nm nanospheres into the EMF- exposed RBCs. Typical TEM images of ultra-thin (70 nm) cross-sections of EMF- exposed RBCs, showing the internalization of 46.3 nm nanospheres. The RBCs exposed to EMF and allowed to reach a temperature of 37°C were able to internalize a greater number of nanospheres than those achieving a maximum temperature of 33°C. Scale bars are 0.5 μm (second row). Inset scale bars are 200 nm (first row). Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 4.1 kW kg-1 that resulted in temperature increases ranging from 20 to 37ºC (at a heating rate of 17ºC per min). Then the SAR was reduced to approximately 3.0 kW kg-1 which resulted in temperature increases ranging from 20 to 33ºC (at a heating rate of 13ºC per min)...... 147 Figure 8.6. Internalization of 23.5 nm nanospheres into the EMF- exposed RBCs. Typical TEM images of ultra-thin (70 nm) cross-sections of EMF- exposed RBCs, showing the internalization of 23.5 nm nanospheres. It appeared that the 23.5 nm nanospheres were able to cross over the 2D spectrin network into the cytosol (indicated by arrows). Scale bars are 200 nm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1 that resulted in temperature increases ranging from 20 to 33ºC (at a heating rate of 13ºC per min).148

Chapter 1. Introduction

1.1. Overview

In Chapter 2, we provide literature on the electromagnetic spectrum and the physical nature of electromagnetic fields (EMFs) together with their effects on biological systems such as bacteria. The dosimetry is introduced and described as a function of the EMF power, frequency, and time of exposure as well as their relationship to operational health and safety standards. A summary of the possible thermal and non-thermal mechanisms taking place as a result of EMF exposure is reviewed together with limitations. This chapter also provides a rationale for cell membrane permeability being the main consequence of exposure of bacterial cells to microwave range EMF. Various techniques that induce cell membrane permeability, including mechanical stress, photoporation, sonoporation and electroporation are described in detail and discussed in relation to their role in drug delivery applications. The functional roles of lipids in membranes are summarised in order to justify the selection of various target cell types for determination of the mechanisms taking place in the bacterial cells as a result of the EMF interaction. In Chapter 3, the experimental design is described in detail, including the EMF exposure conditions, instrument set-up, dosimetry and controls. The scope of Chapter 4 is the description of the materials and methods employed for the study of the biological effects of exposing bacterial cells to 18 GHz EMF radiation, with an emphasis being placed on investigating the effects on membrane permeability, cell morphology and viability. Various cell types, including prokaryotes and eukaryotes are described in detail together with their taxonomic descriptions.

In Chapter 5, we report the biological effects resulting from exposure of Gram negative bacterial cells to 18 GHz EMF, with an emphasis being placed on investigation of the radiation effects on membrane permeability, cell morphology and viability. Branhamella catarrhalis ATCC 23246 (B. catarrhalis) and Escherichia coli ATCC 15034 (E. coli) bacterial cells were selected as representative samples of Gram negative bacteria. The scope of Chapter 6 is the biological effects resulting from exposing Gram-positive bacterial cells to 18 GHz EMF radiation are reported, including changes to the cell membrane permeability, morphology and viability. The Gram-positive Kocuria rosea CIP 71.15T, Planococcus maritimus KMM 3738, Staphylococcus aureus ATCC 25923, Staphylococcus aureus CIP 65.8T, Staphylococcus epidermidis ATCC 14990T and Streptomyces griseus ATCC 23915 bacterial cells were selected for analysis. The effects resulting from prolonged multiple EMF exposures using two strains of Staphylococcus aureus bacteria as model organisms were also studied for the first time. In Chapter 7, we thoroughly study the biological effects resulting from exposing a representative eukaryotic unicellular organism, yeast Saccharomyces cerevisiae ATCC 287 to 18 GHz EMF, including changes to the cell membrane permeability, morphology and viability. The optimisation of dosimetry is also performed and described in detail to obtain cell membrane permeability without compromising the cell viability and morphology. The scope of Chapter 8 is the determination of biological effects associated with red blood cells being exposed to 18 GHz EMF radiation, including changes in cell morphology and membrane permeability.

In Chapter 9, a number of general conclusions are drawn to explain the way in which 18 GHz EMF exposure interacts with cells, resulting in a number of bioeffects, particularly cell permeability, in various typical representatives of prokaryotic and eukaryotic taxa. These included two Gram-negative bacteria (Branhamella catarrhalis ATCC 23246 and Escherichia coli ATCC 15034), six Gram-positive bacteria (Kocuria rosea CIP 71.15T, Planococcus maritimus KMM 3738, Staphylococcus aureus CIP 65.8T, Staphylococcus aureus ATCC 25923, Staphylococcus epidermidis ATCC 14990T, and Streptomyces griseus ATCC 23915), a eukaryotic unicellular organism (yeast Saccharomyces cerevisiae ATCC 287) and red blood cells (obtained from a New Zealand rabbit). The role of the dosimetry parameters associated with the 18 GHz EMF is discussed in terms of the way in which it induces permeability in the cells. Finally, in Chapter 10, we provide a summary of our findings throughout the thesis together with the suggestions for future directions and in drugs/genes delivery applications.

1.2. The aims

Inclusive investigations of EMFs have revealed various biological effects (Celandroni et al. 2004; Cohen et al. 2010; Inhan-Garip et al. 2011; Shamis et al. 2011) upon genes (Aitken et al. 2005; Le Quément et al. 2012; Li et al. 2008; Ruediger 2009; Ruiz-Gómez and Martínez-Morillo 2009), proteins and enzyme kinetics (Bohr and Bohr 2000a; Bohr and Bohr 2000b; George et al. 2008; Gurisik et al. 2006; Laurence et al. 2000), that depended on the EMF strength, frequency, and time of interaction (Banik et al. 2003; Rai et al. 1999). Notwithstanding the sterilization/inactivation effects, many specific effects have been also observed that cannot be explained by the increase in bulk temperature that occurs through EMF exposure alone (Celandroni et al. 2004; Cohen et al. 2010; Inhan-Garip et al. 2011; Shamis et al. 2011).

Despite many studies having been undertaken, the mechanisms responsible for those specific EMF effects are not fully understood and have been the subject of debate (Banik et al. 2003; Celandroni et al. 2004; Cohen et al. 2010; George et al. 2008; Inhan-Garip et al. 2011; Laurence et al. 2000; Ruediger 2009; Shamis et al. 2011). In addition, to the best of our knowledge, the EMF dosage requirements for those specific EMF effects were not optimized systematically. Thus, it was aimed to investigate the specific bioeffects and dosage requirements of 18 GHz EMF on typical representatives of prokaryotic and eukaryotic taxa, including two Gram-negative bacteria (Branhamella catarrhalis ATCC 23246 and Escherichia coli ATCC 15034), six Gram-positive bacteria (Kocuria rosea CIP 71.15T, Planococcus maritimus KMM 3738, Staphylococcus aureus CIP 65.8T, Staphylococcus aureus ATCC 25923, Staphylococcus epidermidis ATCC 14990T, and Streptomyces griseus ATCC 23915), a eukaryotic unicellular organism (yeast Saccharomyces cerevisiae ATCC 287) and red blood cells obtained from a New Zealand rabbit. Three parameters of the EMF exposure (power, duration, and exposure number) were varied to determine their effect on the biological system being studied. Advanced microscopy techniques were employed, comprising Scanning Electron Microscopy (SEM), Transmission Electron Microscopy (TEM), and Confocal Scanning Laser Microscopy (CSLM) together with fluorescent probes, in order to allow a thorough examination of the cell membrane morphology and permeability following EMF exposure/s to be studied.

The overall structure of the thesis is shown in the flowchart of Figure 1.1.

Figure 1.1. Flow chart presenting the structure of the study.

Chapter 2. Literature review

2.1. Overview

This chapter provides an overview of the electromagnetic spectrum and the physical nature of electromagnetic fields (EMFs), and their effect on biological systems such as bacteria. The calculation of dosimetry is described as a function of the EMF power, frequency, and time of exposure. This information is used to evaluate the mechanisms taking place during various EMF induced phenomena as well as their relationship to operational health and safety standards. In the published research of EMF exposure at the cellular level, there are many instances of conflicting data concerning the biological effects of microwave range EMF as well as the proposed mechanisms of interaction. This chapter summarises the possible thermal and non-thermal mechanisms taking place as a result of EMF exposure, together with a review of their limitations. This chapter also provides a rationale for cell membrane permeability being the main consequence of exposure of bacterial cells to microwave range EMF. Various techniques that induce cell membrane permeability, including mechanical stress, photoporation, sonoporation and electroporation are described in detail and discussed in relation to their role in drug delivery applications. The functional roles of lipids in membranes are summarised in order to justify the selection of various target cell types for determination of the mechanisms taking place in the bacterial cells as a result of the EMF interaction.

2.2. The nature of electromagnetic fields (EMFs)

2.2.1. Electromagnetic spectrum

The electromagnetic spectrum covers the entire span of electromagnetic radiation as shown in Figure 2.1. The spectrum consists of ionizing radiation (gamma rays, x-rays, and extreme ultraviolet radiation, with wavelengths below approximately 10-7 m and frequencies above approximately 3 x 1015 Hz); non- ionizing visible radiation (wavelengths ranging from approximately 4 x 10-7 - 7 x 10-7 m and frequencies ranging from approximately 4.2 x 1014 - 7.7 x 1014 Hz); non- ionizing non-visible radiation (short wavelength radio waves and microwaves, with wavelengths ranging between approximately 10-3 - 105 m and frequencies ranging from approximately 3 x 1011 - 3 x 103 Hz; long wavelengths, ranging from approximately 105 - 108 m and frequencies ranging from 3 x 103 - 3 Hz) (Wangsness 1986).

Figure 2.1. The electromagnetic spectrum.

In this thesis, the focus is on the electromagnetic field (EMF) of radiation, with a frequency of 18 GHz, which falls in the microwaves region of the electromagnetic spectrum. According to the International Telecommunication Union (ITU) nomenclature for the frequency and wavelength bands, 18 GHz microwave EMF belongs to the Super High Frequency (SHF) bands. These SHF bands are used for most radar transmitters, wireless LANs, satellite communication, microwave radio relay links, and numerous short range terrestrial data links.

2.2.2. Electromagnetic fields

By definition, the EMF is produced by electrically charged objects that are capable of affecting the behaviour of other charged objects in the vicinity of the field (Wangsness 1986). There are two fields often described as being the sources of the EMF, which are electric and the magnetic fields (Cheng 1989). The electric field surrounds the charged objects and exerts forces on them. In the case of microwave radiation, based on Faraday’s Law of induction; “The induced electromotive force (EMF) in any closed circuit is equal to the time rate of change of the magnetic flux through the circuit”, the electric and magnetic fields are considered to coexist. As a result, one field cannot be considered as being the source of the other due to the high frequencies of microwave radiation (Banik et al. 2003; Cheng 1989; Wangsness 1986). The way in which charge and current interacts with the EMF is described by Maxwell's equations, which are a generalisation of previous laws describing the behaviour of electricity and magnetism (Cheng 1989; Wangsness 1986).

Classical EMF theory is based on the set of equations that are referred to as ‘Maxwell’s equations’. In free space, the differential forms are (Hand 2008):

�� ∇ (Faraday's law, Equation 2.1) × E = - �� ∇ (Ampère-Maxwell law, Equation 2.2) × B = μ0 ( J + ε0 �� ) ρ ∇ (Gauss' law, Equation 2.3) • E = ε0

∇ • B = 0 (Gauss' law for magnetism, Equation 2.4)

and the integral forms are (Hand 2008):

�� ∫ � l ∫ ( S (Equation 2.5) � • d = - � �� ) • d �� ∫ � l ∫ � S ∫ ( S � • d = μ0 � • d + ε0 μ0 � �� ) • d (Equation 2.6) 1 ∫ � S ∫ ρ (Equation 2.7) � • d = ε0 � dV

∫� � • dS = 0 (Equation 2.8) where E is the electric field (V m−1), B is the magnetic flux density (T), J is

−2 −3 the conduction current density (A m ), ρ is the charge density (C m ), μ0 is the

−7 −1 permeability of free space ( = 4π × 10 H m ) and ε0 is the permittivity of free �� space ( = 8.854 × 10−12 F m−1). The term in Equation 2.2 and Equation 2.6 ε0 �� is known as the displacement current density (Hand 2008).

The Equation 2.1, Faraday's law of induction, describes how a changing magnetic field can create an electric field (Hand 2008). For example, a mechanical force (such as the force of moving air rotate the blades of a wind turbines) can spin a huge magnet. The resulting changing magnetic field creates an electric field, which drives electricity through the power grid. In the case of Equation 2.2, Ampère's law with Maxwell's correction states that magnetic fields can be generated in two ways, either through the generation of an electrical current and/or by producing changing electric fields (Hand 2008). The idea that a magnetic field can be induced by a changing electric field follows from the modern concept of displacement current, which was introduced to maintain the solenoidal nature of Ampère's law in a vacuum capacitor circuit (Hand 2008). This modern displacement current concept has the same mathematical form as Maxwell's original displacement current (Hand 2008). Maxwell's original displacement current applies to a polarization current in a dielectric medium and it sits adjacent to the modern displacement current in Ampère's law (Hand 2008).

The Equation 2.3, Gauss's law, describes how electric charge can create and alter an electric field (Hand 2008). Electric fields tend to point away from positive charges, and towards negative charges (Hand 2008). Gauss's law is a primary explanation of why opposite charges attract, and like charges repel; the charges create certain electric fields, to which other charges respond, via an electric force (Hand 2008). In Equation 2.4, Gauss's law for magnetism, states that magnetism is unlike electricity; there are no distinct "north pole" and "south pole" particles that attract and repel in the way that positive and negative charges do (Hand 2008). Instead, north and south poles always exist as pairs (magnetic dipoles) (Hand 2008). In addition, unlike electric fields, which tend to point away from positive charges and towards negative charges, magnetic field lines always come in loops, coming away from the north pole to the outside of a bar magnet towards the south pole and then inside the magnet (Hand 2008).

Based on Maxwell’s equations, Poynting’s theorem was derived, which allows the sum of the electric and magnetic energies to be determined (Poynting 1920).The cross product of the electric and magnetic fields is called the Poynting vector, as defined below (Poynting 1920):

1 S = E x B (Equation 2.9) μ0 2 where S is the Poynting vector(W/m ), μ0 is the permeability of free space

(= 4π × 10−7 H m−1), E is the electric field (V m−1) and B is the magnetic flux density (T) (Poynting 1920).

The Poynting theorem can be summarised by the following formula:

�� ∇ (Equation 2.10) �� + • S = - J • E �� where S is the Poynting vector, is the change in energy density with time, ��

E is the electric field (V m−1) and J is the conduction current density (A m−2) (Poynting 1920).

The energy density u is defined by the following formula:

2 1 2 � (Equation 2.11) u = 2 ( ε0 E + μ0 )

where u is the energy density, ε0 is the permittivity of free space ( = 8.854

× 10−12 F m−1), E is the electric field (V m−1), B is the magnetic flux density (T)

−7 −1 and μ0 is the permeability of free space ( = 4π × 10 H m ) (Poynting 1920).

Therefore, the Poynting theorem can be expanded from Equation 2.2 and Equation 2.3 as:

�� ∇ • S + ε0 E • + • + J • E = 0 (Equation 2.12) �� μ0 ��

Integration of the Poynting vector from Equation 2.10 over a given surface allows the calculation of electromagnetic power in watts (W) (Poynting 1920):

� ∫ � ∮ � ∫ � (Equation 2.13) �� dV + �� dA = - � • E dV

If this power is negative, the net power is entering the sample, which means that the medium inside the sample has absorbing losses. The total power absorption can be obtained by integrating the losses over the whole volume of the sample.

2.2.3. Electromagnetic phenomena

Until now, no electromagnetic phenomenon has been identified that does not satisfy Maxwell’s equations (Cheng 1989; Wangsness 1986). In living systems, however, the number of electromagnetic phenomena is low compared to the extremely broad variety of phenomena that are known in physics and engineering (Banik et al. 2003). Therefore, Maxwell’s equations are generally not applied to describe the electromagnetic phenomena for living systems (Banik et al. 2003). Nonetheless, electromagnetic power is a very important factor for the evaluation of the biological effects of exposing biological systems to radiation. This is because the thermal effects of EMF radiation are known to be related to the power (Banik et al. 2003). In aqueous systems, the electrical component of the EMF can result in dipole interactions with polar molecules such as water, a major and very important constituent of biological cells, and lead to the alignment of the polar ends of these molecules, as well as oscillation. Collisions and friction between these moving molecules result in the production of heat, which can cause damage to bacterial cells (Ponne and Bartels 1995).

Interestingly, many studies have been performed that suggest that exposure of biological systems to an EMF can cause different biological effects. These effects are dependent on the EMF power, frequency, and time of exposure (Banik et al. 2003; Barnabas et al. 2010; Campanha et al. 2007; Dreyfuss and Chipley 1980; Fujikawa et al. 1992; Geveke and Brunkhorst 2008; Kim et al. 2009; Kozempel et al. 2000; Shamis et al. 2009; Shamis et al. 2008; Shazman et al. 2007; Vela and Wu 1979; Welt et al. 1994; Woo et al. 2000; Yeo et al. 1999; Zhou et al. 2010).

2.2.4. Dosimetry

When considering the interaction of an EMF with living systems, it is important to distinguish between the fields outside the body (the exposure) and field levels or absorbed energy within the biological tissue being exposed to the EMF (the dose) (Habash 2007). The exposure is measured as the electric (E) or magnetic (H) field strength, or as power density on the body (Habash 2007). In contrast, the dose absorbed depends on the exposure, as well as on sample geometry, size, orientation with respect to the field, and other factors (Habash 2007). The dose is usually quantified in terms of the specific absorption rate (SAR), which is a good predictor of thermal effects (Habash 2007). The vast majority of the non-thermal biological effects are determined by the minute amounts of energy/power absorbed by specific biomolecules, which cannot be calculated (Panagopoulos et al. 2013). Hence, SAR is usually used as a complementary measure for quality control and cross comparison of instrumentation, but not essentially related to any consequential biological effects; whereas the EMF intensity, along with additional physical parameters (such as frequency, modulation etc.), remains the primary measure for the prediction of biological effects (Habash 2007; Panagopoulos et al. 2013).

Mathematically, the SAR (W kg-1) is defined as:

2 E|| dT c = = SAR =  = c (Equation 2.14)  dt

dT where is the time derivative of the temperature (K s-1), σ is the dt electrical conductivity (S m-1), ρ is the mass density (kg m-3), and c is the specific heat capacity of the sample (J kg-1 K-1).

From Equation 2.14, the localized SAR is directly related to the internal electric field (Habash 2007). Calculation of the internal field is complicated due to many factors, including the nature (near or far field zone) and frequency of the incident field, shape and dimension of the sample, dielectric properties of the sample, and whether the sample is insulated (Habash 2007). Similarly, determination of SAR based on the time derivative of the temperature is complicated due to the presence of temperature heterogeneity and biological variations that exist within a sample in addition to the development of thermotolerance (Habash 2007). Unlike calculations involving the use of an electric field, for which an appropriate continuum physical model may exist, there is still no clear approach to accurately predict the specific heat capacity of a biological sample (Habash 2007). Variations in the specific heat within biological matter are usually much smaller than any corresponding variations in conductivity, which results in a much more uniform temperature than an electric field distribution (Panagopoulos et al. 2013). Thus, estimation of SAR based on the time derivative of the temperature is more accurate than making an estimation based upon the internal electric field (Panagopoulos et al. 2013).

2.2.5. Operational health and safety standards

In order to avoid possible health hazards and harmful interaction with EMFs, an independent scientific organization, the International Commission on Non- Ionizing Radiation Protection (ICNIRP), in conjunction with the Environmental Health Division of the World Health Organization (WHO), developed international guidelines to limit the exposure to EMFs (Ahlbom et al. 1998; ICNIRP 2009; Sienkiewicz et al. 2016). The exposure limits, also known as basic restrictions, are set with an additional reduction factor in relation to the threshold that has been shown to result in adverse effects to biological systems. This limit is designed to eliminate any possible scientific uncertainties with regard to exposure (Ahlbom et al. 1998; ICNIRP 2009; Sienkiewicz et al. 2016). Generally, the basic restrictions are expressed in terms of the SAR (Ahlbom et al. 1998; ICNIRP 2009; Sienkiewicz et al. 2016). At frequencies greater than about 10 GHz, SAR is not recommended for assessing absorbed energy due to the small depth of penetration into tissues (Ahlbom et al. 1998; ICNIRP 2009; Sienkiewicz et al. 2016). In these cases, the incident power density of the field (in W m-2) is a more appropriate dosimetric quantity (Ahlbom et al. 1998; ICNIRP 2009; Sienkiewicz et al. 2016). Between 10 and 300 GHz, the basic restrictions of power density for occupational exposure and exposure to the general public are up to 50 W m-2 and 10 W m-2, respectively, to prevent excessive heating in any biological tissue at or near the body surface (Ahlbom et al. 1998; ICNIRP 2009; Sienkiewicz et al. 2016).

2.2.6. Current understanding of the biological effects induced by microwave range electromagnetic fields

Studies on the biological effects of EMF radiation exposure at different frequencies in the microwave range can be approached several ways, such as epidemiology, human studies, animal studies, and cellular studies (Ahlbom et al. 1998; ICNIRP 2009; Sienkiewicz et al. 2016). Whilst epidemiological and human studies directly report results related to human health, it should be recognized that cellular and animal studies are of value in assessing causality and biological plausibility (van Deventer et al. 2011). Extensive research has been conducted with regard to EMF exposure below basic restriction levels. These studies have shown that no adverse health effects such as headaches, concentration difficulty, sleep quality, cognitive function, cardiovascular effects, brain tumours, etc. have arisen as a result of exposure (Ahlbom et al. 1998; ICNIRP 2009; Sienkiewicz et al. 2016; van Deventer et al. 2011). Cellular studies remain one of the best ways to elucidate any bioeffects resulting from EMF exposure (Ahlbom et al. 1998; ICNIRP 2009; Shamis et al. 2012a; Sienkiewicz et al. 2016; van Deventer et al. 2011).

2.2.6.1. Biological effects of microwave range electromagnetic fields at cellular level

At a cellular level, the ability for microwave range EMF exposure to induce any biological effects can be studied, together with investigations into the plausibility of existing mechanistic hypotheses to explain any resulting effects (van Deventer et al. 2011). The most significant advantage of cellular studies is that they are straightforward to perform, allow reproducibility to be determined under precisely controlled parameters, and allow good dosimetry studies to be performed that are able to be statistically assessed for their reliability because large sample sizes are being investigated (Michaelson et al. 2006).

There have been many studies of the biological effects of EMF radiation exposure to biological systems. These studies have been performed using different EMF frequencies in the microwave range (Aitken et al. 2005; Bohr and Bohr 2000a; Bohr and Bohr 2000b; Campanha et al. 2007; Celandroni et al. 2004; Cohen et al. 2010; Dreyfuss and Chipley 1980; George et al. 2008; Gurisik et al. 2006; Harris et al. 1990; Inhan-Garip et al. 2011; Kim et al. 2009; Laurence et al. 2000; Le Quément et al. 2012; Li et al. 2008; Ruediger 2009; Ruiz-Gómez and Martínez- Morillo 2009; Schelle 1996; Shamis et al. 2012b; Shamis et al. 2009; Shamis et al. 2011; Shamis et al. 2008; Woo et al. 2000; Zhou et al. 2010). It is generally accepted that EMF radiation causes dielectric heating of exposed tissues, cells, and cellular components (Ahlbom et al. 1998; ICNIRP 2009; Shamis et al. 2012a; Sienkiewicz et al. 2016; van Deventer et al. 2011). It was found that microwave range EMF radiation is capable of reducing the extent of microbial proliferation (Campanha et al. 2007; Dreyfuss and Chipley 1980; Harris et al. 1990; Kim et al. 2009; Schelle 1996; Shamis et al. 2009; Shamis et al. 2008; Woo et al. 2000; Zhou et al. 2010). Many microorganisms have been known to be inactivated as a result of exposure to EMF radiation in the microwave range, such as Candida albicans, Bacillus licheniformis, Bacillus subtilis, Acinetobacter calcoaceticus, Pseudomonas aeruginosa, Burkholderia cepacia, Clostridium perfringens, Escherichia coli, Streptococcus faecalis, Staphylococcus aureus, Staphylococcus epidermidis, Salmonella, and Listeria spp. (Campanha et al. 2007; Dreyfuss and Chipley 1980; Harris et al. 1990; Kim et al. 2009; Schelle 1996; Shamis et al. 2009; Shamis et al. 2008; Welt et al. 1994; Woo et al. 2000; Yeo et al. 1999; Zhou et al. 2010).

In addition to thermal damage, EMF radiation exposure at different frequencies in the microwave range were found to affect eukaryotic cells at subcellular levels, having an influence over genes, proteins and enzyme kinetics (Bohr and Bohr 2000a; Bohr and Bohr 2000b; Buttiglione et al. 2007; George et al. 2008; Gerner et al. 2010; Gurisik et al. 2006; Laurence et al. 2000; Le Quément et al. 2012; Li et al. 2008; Ruediger 2009; Ruiz-Gómez and Martínez-Morillo 2009; Shahbazi-Gahrouei et al. 2016; Shamis et al. 2012b; Shamis et al. 2011; Zhao et al. 2007; Zuo et al. 2014). A summary of the literature is presented in Table 2.1.

Table 2.1. A summary of the non-thermal biological effects of EMF in the microwave range. Experimental setup EMF Exposure Exposure Biological organism Effects Proposed mechanisms Reference frequency power/SAR temperature (°C) 900 MHz 1.0 W/kg 37.0 ± 1.0 Human neuroblastoma The EMF affected both Egr-1 gene The mechanism was a (Buttiglione cell expression and cell regulatory direct interaction with the et al. 2007) functions, involving apoptosis EMF. inhibitors like Bcl-2 and survivin. 900 MHz 354.6 37.0 ± 0.7 Human mesenchymal The EMF reduced cell viability and The mechanism was a (Shahbazi- µW/cm2 stem cells derived proliferation rates of human direct interaction with the Gahrouei et from adipose tissue mesenchymal stem cells. EMF. al. 2016) 1800 MHz 2.0 W/kg 37.0 ± 0.1 Cerebral cortical and After EMF exposure, 24 up- The mechanism was a (Zhao et al. hippocampal neuronal regulated genes and 10 down- direct interaction with the 2007) cultures regulated genes were identified EMF. which are associated with multiple cellular functions (cytoskeleton, signal transduction pathway, metabolism, etc.). 1800 MHz 2.0 W/kg 37.0 ± 0.1 Jurkat T-cells and The EMF caused a significant The mechanism was due to (Gerner et al. human fibroblasts increase in protein synthesis. the radiation induced 2010) excitation of hydrogen bonds.

1800 MHz 1209 37.0 Mouse NIH/3T3 and The EMF decreased significantly The mechanism was due to (Xing et al. mW/m2 human U-87 MG cells the cell viability and induced a direct interaction of EMF 2016) apoptosis-related events such as with the DNA and reactive oxygen species (ROS) mitochondrial ROS burst and more oxidative DNA generation. damage.

Table 2.1. Cont. Experimental setup EMF Exposure Exposure Biological organism Effects Proposed mechanisms Reference frequency power/SAR temperature (°C) 2.45 GHz 800 W 2 - 44 β-lactoglobulin protein EMF induced denaturation of β- The protein conformational (Bohr and lactoglobulin protein at lower changes were due to the Bohr 2000a; temperature than conventional resonant EMF energy Bohr and heating. absorption by intrinsic Bohr 2000b) protein modes. 2.45 GHz 4.85 25 Citrate synthase The EMF cause a significantly The effect was due to the (George et al. kW/kg protein higher degree of protein unfolding resonant absorption of EMF 2008) than conventional heating to the energy to some vibrational same maximum temperature. states of the protein or its bound water envelope. 2.856 GHz 30 mW/cm2 37.0 ± 0.5 PC12 After EMF exposure, cells formed The mechanism was a (Zuo et al. pheochromocytoma chromatin and apoptotic body with direct interaction with the 2014) cells decreasing mitochondria membrane EMF. potential and increasing DNA fragmentation. 18 GHz 1500 20 - 40 Escherichia coli The EMF induced cell permeability The mechanism was a (Shamis et al. kW/m3 and increased in the catalytic direct interaction with the 2012b; activity of two common bacterial EMF. Shamis et al. enzymes, lactate dehydrogenase 2011) and cytochrome c oxidase. 60.4 GHz 42.4 W/kg 37.0 ± 1.0 Primary human skin The cells expressed differentially No mechanism was given. (Le Quément cells five genes, namely CRIP2, et al. 2012) PLXND1, PTX3, SERPINF1, and TRPV2.

It has been reported that exposure at 900 MHz (SAR 1.0 W/kg) affected the regulatory function of human neuroblastoma cells, involving apoptosis and surviving (Buttiglione et al. 2007). In another exposure study at 900 MHz, with an intensity of 354.6 µW/cm2 square waves (217 Hz pulse frequency, 50% duty cycle), the authors observed a reduction in the viability and proliferation rates of human mesenchymal stem cells derived from adipose tissue (Shahbazi-Gahrouei et al. 2016). Zhao et al. (2007) reported some modifications in the expression of genes associated with cytoskeleton organization, signal transduction pathways, and cell metabolism of cerebral cortical and hippocampal neuronal cultures for samples exposed to 1800 MHz EMF. It was found that after 8 h of exposure (SAR 2.0 W/kg), with a temperature increase of less than 0.15 °C, there was a significant increase in the synthesis of proteins in Jurkat T-cells and human fibroblasts, and to a lesser extent in the activated primary human mononuclear cells (Gerner et al. 2010). Furthermore, Xing et al. (2016) reported that after exposing samples to 1800 MHz EMF at a power density of 1209 mW/m2, mouse NIH/3T3 and human U-87 MG cells significantly reduced their cell viability, inducing apoptosis-related events such as a ROS burst and more oxidative DNA damage. It was found that the 2.45 GHz EMF induced the denaturation of the β-lactoglobulin protein at a lower temperature than that obtained using conventional heating methods (Bohr and Bohr 2000a; Bohr and Bohr 2000b). Similarly, George et al. (2008) revealed that exposure to 2.45 GHz EMF radiation (SAR 4.85 kW/kg) caused a significantly higher degree of unfolding protein citrate synthase than observed using conventional heating methods to the same maximum temperature. After exposure to 2.856 GHz EMF, with an average power density of 30 mW/cm2, differentiated PC12 pheochromocytoma cells were reported to form chromatin and an apoptotic body with decreasing mitochondria membrane potential and increasing DNA fragmentation (Zuo et al. 2014). It was reported that exposing samples to an 18 GHz EMF (SAR 1500 kW/m3 with a resultant temperature of 40 °C) increased the catalytic activity of two common bacterial enzymes, lactate dehydrogenase (LDH) and cytochrome c oxidase (COX) from Escherichia coli cells (Shamis et al. 2012b). After exposure to 60.4 GHz with an average incident power density of 1.8 mW/cm2 and an average SAR of 42.4 W/kg, primary human skin cells were found to express

41 five genes to a different extent, namely CRIP2, PLXND1, PTX3, SERPINF1, and TRPV2 (Le Quément et al. 2012).

There have been some inconsistent results reported in the literature regarding the biological effects resulting from exposure of biological systems to EMF radiation (d'Ambrosio et al. 2002; Danese et al. 2017; Liu et al. 2012; Liu et al. 2015; Ruediger 2009; Schwarz et al. 2008). For example, it has been reported that exposing C6 glioma cells and human glioblastoma cells to 1950 MHz EMF (SAR 5 W/kg) for periods up to 48 h did not affect their proliferation or the gene expression profile of the cells (Liu et al. 2012; Liu et al. 2015). Luukkonen et al. (2009) reported the enhancement of ROS production and DNA damage in human SH-SY5Y neuroblastoma cells after exposure to the 872 MHz continuous waves EMF at 5 W/kg but not to the global system for mobile communications (GSM) modulated EMF at similar SAR.

In another case study, Schwarz et al. (2008) reported the genotoxic effects resulting from exposure of cells to 1,950 MHz EMF, which is below the specific absorption rate (SAR) safety limit of 2 W/kg on human fibroblasts. Similarly, d'Ambrosio et al. (2002) reported that no change in cell proliferation kinetics or cytogenetic damage of human lymphocytes was observed after exposing the samples to a continuous wave of 1.748 GHz EMF radiation (SAR 5 W/kg). In addition, Danese et al. (2017) stated that exposing human lymphocytes to 900 MHz EMF radiation for 30 min periods did not significantly impact the integrity of the DNA within the samples. In contrast, Belyaev et al. (2009) revealed that exposure to 1947.4 MHz and 915 MHz EMF radiation at non-thermal levels lower than the ICNIRP safety standards can affect chromatin and inhibit the formation of DNA double-strand breaks, co-localizing 53BP1/γ-H2AX DNA repair foci in human lymphocytes from hypersensitive and healthy people.

The reason for these inconsistent results may be a function of flaws in the experimental procedure and technical design of reaction systems being used (Ruediger 2009; van Deventer et al. 2011). Some studies lack sufficient experimental replication through the use of more precise quantitative and specific measurements (Ruediger 2009; van Deventer et al. 2011). In addition, the 42 magnitude of any changes being observed is usually small, making interpretation of the possible mechanisms in play being very difficult (van Deventer et al. 2011).

2.2.6.2. The combined biological effects resulting from exposure to microwave range electromagnetic fields and other agents

Exposure of cells to microwave range EMF radiation not only induces some biological effects, but also can act as a co-promoter or potentiator of other agents (Boga et al. 2015; Cao et al. 2009; Kostoff and Lau 2017). For example, it was observed that exposure to 872 MHz continuous waves EMF (with a SAR of 5 W/kg) enhanced the effect of menadione, a chemical that induces the production of intracellular reactive oxygen species, thus causing an increase in secondary DNA damage in human SH-SY5Y neuroblastoma cells (Luukkonen et al. 2009). Similarly, Cao et al. (2009) discovered that pre-exposure of cells to 900 MHz EMF radiation (with a power density of 6 mW/cm2) enhanced the γ-ray damage of SHG44 human glioma cells, as evidenced by a significant inhibition of cell proliferation, with increased apoptosis, and oxidative stress.

It is possible that exposure of biological systems to microwave range EMF radiation and other agents may result in no biological effect being observed. If combined, however, some biological effects may result (Boga et al. 2015; Kostoff and Lau 2017). For example, Boga et al. (2015) found that subjecting samples to an EMF exposure (at 900 MHz and 1800 MHz with a SAR of 1 W/kg) in the presence of nicotine sulfate resulted in dramatic abnormalities and death among the Xenopus embryos; whereas exposure to the EMF or nicotine sulfate alone did not result in such an effect.

2.2.6.3. Electromagnetic field effects on cell membranes resulting from exposure to microwave range radiation

It was recently reported that after exposing Escherichia coli cells to 18 GHz EMF, the cells developed reversible poration/permeability in their cell membrane. This phenomenon was termed ‘microwave-induced poration’ (Shamis et al. 2011). The most significant implication of cell membrane permeability is that it allows the passage of different type of impermeable ions, molecules and macromolecules through the membrane (Ibey et al. 2011; Karshafian et al. 2005; Napotnik et al. 2012; Pakhomov et al. 2009; Stevenson et al. 2010; Yang et al. 2011). This has implications in many biomedical engineering, cell drug delivery and gene therapy applications (Chen et al. 2006; Granot and Rubinsky 2008; Hofmann et al. 1999; Ibey et al. 2011; Song et al. 2007; Yang et al. 2011). The effect of EMF radiation exposure at 18 GHz on other prokaryotic and eukaryotic cells belonging to different taxonomic groups and having different lipid membrane compositions has not been investigated. In addition, a study of the cell membrane poration/permeability arising from EMF exposures at different frequencies would be required in order to gain a comprehensive understanding of the dose-specific characteristics and threshold levels required to produce this effect. In addition, the mechanism of EMF induced poration/permeability effect has not yet been elucidated, and hence studies that would allow a mechanism to be postulated would be of significant benefit.

2.2.6.5. Proposed mechanisms responsible for the observed biological effects in cells resulting from exposure to microwave range EMF radiation

Despite the many studies that have been undertaken, the mechanisms responsible for the observed biological effects resulting from exposure of cells to EMF radiation in the microwave range are not well understood. The mechanisms

44 responsible have been the subject of debate for more than 30 years (Banik et al. 2003; Barnabas et al. 2010; Campanha et al. 2007; Dreyfuss and Chipley 1980; Fujikawa et al. 1992; Geveke and Brunkhorst 2008; Kim et al. 2009; Kozempel et al. 2000; Shamis et al. 2009; Shamis et al. 2008; Shazman et al. 2007; Vela and Wu 1979; Welt et al. 1994; Woo et al. 2000; Yeo et al. 1999; Zhou et al. 2010). There is, however, a consensus that the EMF radiation must interact with the biological molecules or structures, changing the charge distribution, chemical state, or energy, which can induce other effects through the biological scale of complexity (Apollonio et al. 2013).

Initially it was thought that the biological effects of EMF radiation at different frequencies in the microwave range on biological systems may arise due to thermal mechanisms, also known as dielectric heating (Challis 2005; Fujikawa et al. 1992; Kozempel et al. 2000; Maktabi et al. 2011; Shazman et al. 2007; Vela and Wu 1979; Welt et al. 1994; Woo et al. 2000). Dielectric heating is caused by absorption of the EMF energy from the electrical conductivity of most biological systems (Challis 2005). The rapid transfer of the oscillating EMF electric current energy into the translational motion of molecules results in an increase in the local temperature (Challis 2005). The main contribution to the heating comes from hindered molecules such as water, the heating of which arises due to its large permanent dipole moment (Challis 2005).

By contrast, many EMF studies have either shown or proposed non-thermal mechanisms for the observed biological effects in terms of the EMF energy transferred to the sample, producing various types of molecular transformations and alterations inside the cells (Apollonio et al. 2013; Banik et al. 2003; Barnabas et al. 2010; Challis 2005; Dreyfuss and Chipley 1980; Gaestel 2010; Geveke and Brunkhorst 2008; Shamis et al. 2009; Shamis et al. 2008). Three principal strategies have been used to study the non-thermal effects of radiation exposure to cells (Apollonio et al. 2013; Gaestel 2010). The first approach is to employ a thermostatic regulation of the temperature within the exposed sample to avoid any thermal activation (Apollonio et al. 2013; Gaestel 2010). In cases where the EMF exposure leads to a significant degree of heating, the second strategy is to compare the 45 samples exposed to the EMF radiation to control samples that have been heated by a conventional method, then the differences in the cell response observed (Apollonio et al. 2013; Gaestel 2010). This process can only be successful if the temperature delivered to the sample can be carefully monitored and evenly distributed (Apollonio et al. 2013; Gaestel 2010). The final strategy is to deliver non-thermal doses with SAR values below 2.0 W/kg to the sample to ensure that the temperature of the sample is carefully controlled (Apollonio et al. 2013; Gaestel 2010).

When considering the non-thermal mechanisms taking place in a biological samples as a result of EMF exposure, it should be noted that the energy required to ionise a molecule, 1 eV, is 2 x 105 times larger than the photon energy of 1 GHz EMF, 4 µeV (Challis 2005). Hence, the non-thermal effects resulting from EMF exposure will not be a result of molecular ionization or excitation due to the absorption of a single photon, nor through the immediate absorption of 2 x 105 photons (Challis 2005). Another important aspect to be noted is the thermal noise, which is the random fluctuation of the electric and magnetic fields associated with the random motion of charges/ions (Apollonio et al. 2013; Challis 2005). It is suggested that in order for these fluctuations to cause biological effects, the applied EMF must transfer an amount of energy greater than the corresponding to the thermal noise energy of all biological systems that are well thermally coupled to their surroundings (Apollonio et al. 2013; Challis 2005).

One of the most studied hypotheses on the non-thermal mechanisms taking place in biological systems as a result of EMF exposure is the prospect of the induction of conformational changes in proteins such as enzymes, ionic channels, and pumps, which in turn results in changes in their activity (Apollonio et al. 2013; Challis 2005; Shamis et al. 2012b; Soghomonyan and Trchounian 2013; Torgomyan et al. 2012; Torgomyan et al. 2011; Torgomyan and Trchounian 2012). It has been hypothesised that the conformational changes taking place in proteins can be due to the direct interaction of the electric field on the protein dipole moments. For example, Sato et al. also proposed that EMF energy could either cause dipoles to rotate and line up rapidly (2450 million times per sec) and/or cause ions to accelerate and collide with other molecules, which could speed up various 46 enzyme activities (Sato et al. 1996). Astumian (2003) suggested that the EMF coupling to the dipole moments of the components present in the enzyme ATPases, which are proteins that span cell membranes, could be capable of inducing conformational changes. These changes then could then lead to ion movement across the cell membrane (Astumian 2003). In another study, English and Mooney (2007) noted changes in the lysozyme secondary structure in egg whites as a consequence of alignment of the protein’s total dipole moment with the applied EMF.

Another hypothesis for the conformational changes taking place in proteins upon EMF exposure is the resonant energy being absorbed by the intrinsic protein modes. In a series of studies, Bohr and Bohr suggested that the conformation adopted by a protein depends on the amplitudes of the dynamic excitations within is structure (Bohr and Bohr 2000a; Bohr and Bohr 2000b). By applying an EMF dose that is close to the frequency of the intrinsic protein mode, changes in the secondary and tertiary protein structure of various microorganisms can be achieved (Bohr and Bohr 2000a; Bohr and Bohr 2000b).

Other studies have been reported that suggest that the EMF-induced changes in the property of water molecules within micro-organisms can alter the conformation of their proteins, their degree of hydration, and other properties that result a change in their activity (Shamis et al. 2012b; Soghomonyan and Trchounian 2013; Torgomyan et al. 2012; Torgomyan et al. 2011; Torgomyan and Trchounian 2012). It was demonstrated that exposing bacteria to an EMF of 51.8, 53, 70.6, 73 and 90 GHz frequency resulted in changes in their enzymatic activities and ion (H+ and K+) transport processes through the plasma membrane (Soghomonyan and Trchounian 2013; Torgomyan et al. 2012; Torgomyan et al. 2011; Torgomyan and Trchounian 2012).

2.3. Cell membrane permeability and drug delivery systems

Cell membrane permeability can arise as a the result of cell membrane poration, which is known to be caused by a rearrangement in the molecular structure of the cell membrane, together with interactions taking place at the aqueous–lipid interface (Chen et al. 2010; Chen et al. 2006; Hosokawa et al. 2011; Ibey et al. 2010; Ibey et al. 2011; Karshafian et al. 2005; Koshiyama et al. 2010; Napotnik et al. 2012; Pakhomov et al. 2009; Pakhomova et al. 2011; Paterson et al. 2005; Shamis et al. 2011; Song et al. 2007; Stevenson et al. 2006; Stevenson et al. 2010; Torres- Mapa et al. 2010; Vernier and Ziegler 2007; Weaver 2003; Yang et al. 2011). According to classical nucleation theory (CNT), the formation of pores in membranes is controlled by the competition between the surface tension of the membrane and the line tension associated with the rim of the pore (Leontiadou et al. 2004; Tieleman et al. 2003; Wang and Frenkel 2005). In such simplified models, the free energy of formation (E) of a cylindrical pore with a radius (r) is approximated by Equation 2.15:

2 �(�) = 2�� − �� Γ (Equation 2.15)

where γ is the line tension that opposes the creation of the pore and Γ is the surface tension that lowers the energetic barrier for pore creation and expansion (Leontiadou et al. 2004).

Increases in the surface tension are often regarded as the main driving force for the cell membrane poration process (Farago and Santangelo 2005). In such cases, the cell membrane is believed to stretch and spontaneously form initial ruptures in the membrane in order to relax the surface tension of the membrane itself (Farago and Santangelo 2005; Leontiadou et al. 2004). When the radius of the initial ruptures reaches a critical value, they transform into cylindrical pores that become lined with phospholipid head groups. These continue increasing in size until a state of zero surface tension is reached (Farago and Santangelo 2005; Tieleman et al. 2003). Such membrane pores can either be temporary and reseal following an

48 elapsed period, or continue to expand and eventually rupture the membrane depending on the degree of the external shock, time of exposure and cell characteristics (Chen et al. 2006; van Uitert et al. 2010; Weaver 2003). The cell membrane can also initiate a change in the osmotic pressure due to the passage of internal and external components through the pores that are formed (Farago and Santangelo 2005; Ibey et al. 2010; Ibey et al. 2011; Pakhomov et al. 2009; Pakhomova et al. 2011).

Cell membrane poration can be physically induced through the application of external shocks such as mechanical stress (i.e., pipet aspiration) (Leontiadou et al. 2004; Tieleman et al. 2003), or the application of ultrasound (sonoporation) (Karshafian et al. 2005; Koshiyama et al. 2010; Song et al. 2007), electric fields (electroporation) (Chen et al. 2010; Chen et al. 2006; Ibey et al. 2010; Ibey et al. 2011; Napotnik et al. 2012; Pakhomov et al. 2009; Pakhomova et al. 2011; Vernier and Ziegler 2007; Weaver 2003; Yang et al. 2011), lasers (photoporation) (Hosokawa et al. 2011; Paterson et al. 2005; Stevenson et al. 2006; Stevenson et al. 2010; Torres-Mapa et al. 2010), and EMF (microwave-induced poration) (Shamis et al. 2011).

2.3.1. Mechanical stress

Application of mechanical stress as an anisotropic pressure (ie. pipet aspiration) to a membrane comprised of phospholipid bilayers increases its surface tension, which leads to an increase in membrane surface area through a membrane- stretching process (Farago and Santangelo 2005; Leontiadou et al. 2004; Tieleman et al. 2003). When the surface area reaches a critical value, an initial rupture occurs and grows rapidly, forming an approximately cylindrical pore that becomes lined with phospholipid headgroups on a nanosecond time scale (Tieleman et al. 2003). Once formed, the pore continues to increase in size until the bilayer is destabilized (Tieleman et al. 2003).

2.3.2. Photoporation

Photoporation utilizes tightly focused laser light to induce the reversible poration of the cellular membrane, allowing exogenous material to pass through the membrane and enter the cell (Hosokawa et al. 2011; Palumbo et al. 1996; Paterson et al. 2005; Schneckenburger et al. 2002; Stevenson et al. 2006; Stevenson et al. 2010; Torres-Mapa et al. 2010). Laser light with wavelengths in the ultraviolet (UV), visible (VIS), and infrared (IR) region, both as pulsed (nanosecond or femtosecond) and continuous wave (CW) applications, have been used for photoporation (Torres-Mapa et al. 2010). For CW lasers, the poration mechanism is most likely due to localized heating of the cellular membrane by the laser irradiation (Hosokawa et al. 2011; Palumbo et al. 1996; Schneckenburger et al. 2002; Stevenson et al. 2010). Energy absorption in the UV and visible range corresponds to the energy corresponding to the electronic transitions taking place in molecules, while infrared radiation is associated with the vibrational transitions taking place in the molecules (Xiong et al. 2016). The non-radiative relaxation to the ground state results in the production of heat (Xiong et al. 2016). The accumulated localised heat leads to a local disruption of the membrane, either by a local phase transition of the lipid bilayer and/or by the thermal denaturation of integral proteins, which in turn generates pores in the membrane (Delcea et al. 2012; Hosokawa et al. 2011; Ivanov et al. 2007; Xiong et al. 2016).

In the case of pulsed lasers, the response of the cell membrane heavily depends on the energy and duration of the pulse (Hosokawa et al. 2011; Stevenson et al. 2010). It has been suggested that gene delivery efficiency is controlled by the size of the focal point and pulse frequency of the laser (Kamimura et al. 2011). It was reported that for pulsed lasers, localized heating by the absorption of single photons alone is not sufficient to effectively form pores in cell membranes (Quinto- Su and Venugopalan 2007; Xiong et al. 2016). This is mainly because water, lipids, and proteins have a relatively low absorption in the 350–1100 nm wavelength range (Quinto-Su and Venugopalan 2007; Xiong et al. 2016). For this reason, dye molecules such as phenol red were used to enhance the absorption of light by the cell, achieving pore formation (Palumbo et al. 1996; Schneckenburger et al. 2002).

Long pulse duration times (nanoseconds) or equivalently, high pulse energy (exceeding 800 nJ/pulse) not only causes an increase in bulk temperature, but also generates reactive free electrons and, more importantly, the dispersion of cells surrounding the laser focal point due to the formation of shockwave and cavitation bubbles (Hosokawa et al. 2011; Stevenson et al. 2010; Xiong et al. 2016). The cavitation bubbles are formed by the local evaporation of the medium within the cell (which is mostly water) and the expansion of the plasma (Vogel et al. 2005; Xiong et al. 2016). When the cavitation bubbles have expanded to their maximum size, the bubbles then collapse as a result of the surrounding hydrostatic pressure, inducing liquid jets or shockwaves that can form pores in the cell membrane (Vogel et al. 2005; Xiong et al. 2016). For short pulse durations (femtoseconds) or equivalently low pulse energies (less than 800 nJ/pulse) below the threshold for optical breakdown or cavitation bubble formation, only free reactive electrons are generated at the cell membrane surface (Hosokawa et al. 2011; Stevenson et al. 2010; Tirlapur et al. 2001). The reactive free electrons produce highly reactive oxygen species (ROS) which can locally induce cell membrane damage (Tirlapur et al. 2001; Xiong et al. 2016).

Photoporation has been extensively studied in cell transfection with nucleic acids (DNA, mRNA, siRNA), proteins, ions, small molecules, and semiconductor nanocrystals into several cell types (Clark et al. 2006; Tirlapur and Konig 2002; Tsukakoshi et al. 1984; Xiong et al. 2016). Femtosecond laser-assisted direct photoporation was also used in cellular imaging, particularly, to deliver actin- staining fluorophores into rat cortical neurons for visualizing the cytoskeleton of dendrites (Dhakal et al. 2014). Various anti-cancer drugs were delivered into cancer cells both in vitro and in vivo by photoporation for an enhanced chemotherapeutic effect (Lukianova-Hleb et al. 2014; Lukianova-Hleb et al. 2012). While being capable to deliver compounds into cells, photoporation can also be used to released compounds from cells (Delcea et al. 2012).

2.3.3. Sonoporation

Sonoporation is the process that utilises ultrasounds and microbubbles as contrast agents to physically induce cell permeability by either generate reversible pores in the cell membrane or endocytosis (Karshafian et al. 2005; Koshiyama et al. 2010; Lentacker et al. 2014; Meijering et al. 2009; Song et al. 2007). The microbubbles are gas-filled structures stabilized by a lipid, protein or polymer shell and some of them have been clinically approved (Ferrara et al. 2007; Lentacker et al. 2014). Due to the gas-filled, the microbubbles are compressible in respond to the ultrasound pressure waves (Lentacker et al. 2014). The process of alternate growing and shrinking is called cavitation, which depends on the ultrasound intensities can be divided into stable and inertial cavitation (Lentacker et al. 2014). Occurring at higher ultrasound intensities, the inertial cavitation can lead to microbubble implosion which can result in much stronger biophysical effects (Lentacker et al. 2014). The sonoporation can be induced by three conditions, which are low intensity ultrasound leading to stable cavitation of microbubbles, high intensity ultrasound leading to inertial cavitation with bubble collapse, and ultrasound without microbubbles (Lentacker et al. 2014).

During stable cavitation, the microbubbles oscillate and create a liquid flow around called microstreams, which can exert shear stress (Lentacker et al. 2014; VanBavel 2007). When closely located to the cell membrane, this shear stress together with microbubbles can gently push and pull the cell membrane (Lentacker et al. 2014; van Wamel et al. 2006). In this way, the cellular membrane is disturbed as a result of mechanical stress (Lentacker et al. 2014; van Wamel et al. 2006). Another possible mechanism is the formation of free radicals, which play an important role in the increased cell membrane permeability for Ca2 + (Juffermans et al. 2009; Lentacker et al. 2014). Finally, it was reported that the generation of reactive oxygen species (ROS) can induce cellular injury via lipid peroxidation which can result in lipid bilayer rearrangement and membrane disruption (Lentacker et al. 2014). The size of the cell membrane pores arising was estimated

52 from several tens of nanometers to a few hundreds of nanometer (Karshafian et al. 2005; Karshafian et al. 2010; Lentacker et al. 2014).

In the case of inertial cavitation, the microbubbles can collapse due to the inertia of the inrushing fluid, which generate shock waves and liquid microjets (Lentacker et al. 2014; Postema et al. 2004). It has been shown that shock waves and microjets create very high forces that can puncture cell surfaces and create cell membrane pores (Lentacker et al. 2014; Postema et al. 2004). While stably cavitating microbubbles need to have direct contact with the cell to affect the membrane, the inertial cavitating microbubbles can reach over a larger distance (Lentacker et al. 2014; Zhou et al. 2012). However, the maximal distance between the inertial cavitating microbubbles and cell membrane should not exceed the microbubble diameter to cause an effective impact on the cellular membrane (Lentacker et al. 2014; Zhou et al. 2012). The pore sizes which have been reported as a consequence of inertial cavitation (hundreds of nanometer to micrometer range) are larger than pores reported during stable cavitation (few nm to hundreds of nanometers) (Lentacker et al. 2014).

For ultrasound without microbubbles, it is believed that the two factors that trigger poration in the membrane on a nanosecond time scale are the repetitive irradiation of ultrasounds causes the inclusion of water molecules in the hydrophobic region of cell lipid membrane, which increases the surface tension (Koshiyama et al. 2010); accompanied with acoustic streaming from continuous impacts of shock waves on membrane, which exerts shear stresses on cell membranes (Baker et al. 2001; Humphrey 2007; Lentacker et al. 2014).

It is believed that ultrasound is capable of inducing endocytosis via two possible mechanisms (Apodaca 2002; Juffermans et al. 2009; Lentacker et al. 2014). The first mechanism is that the ultrasound induced mechanical forces and/or shear stress from microstreamings/acoustic streaming cause changes in cell membrane tension which lead to plasma membrane deformation accompanied by cytoskeletal rearrangements (Apodaca 2002; Juffermans et al. 2009). These changes can be sensed by mechanosensors, such as integrins and stretch-activated ion channels, which in turn transduce these signals into downstream cellular 53 processes such as endocytosis and exocytosis (Apodaca 2002; Juffermans et al. 2009). It is believed that these processes of exocytosis and endocytosis relax the plasma membrane tension (Apodaca 2002; Juffermans et al. 2009).

In the second mechanism, the shear forces can result in a physical disruption of the cell membrane which will also lead to elevated intracellular Ca2 + levels due to a concentration driven passive diffusion of Ca2 + (Hassan et al. 2010; Reddy et al. 2001). This influx of Ca2 + through membrane wounds triggers exocytosis, with recruitment of intracellular vesicles such as lysosomes to the site of injury (Hassan et al. 2010; Reddy et al. 2001) or endocytosis (Draeger et al. 2011; Meijering et al. 2009). This endocytosis can be a consequence of exocytosis (Tam et al. 2010). The exocytotic release of lysosomal acid sphingomyelinase (ASM) converts sphingomyeline in the cell membrane to ceramide and promotes inward budding together with vesicle formation (Tam et al. 2010). The Ca2 + can also directly stimulate endocytosis independently of ASM release but via interaction with cholesterol rich cell membrane areas, which can spontaneously vesiculate and form endocytic vesicles (Lariccia et al. 2011).

The sonoporation has been studied extensively in the fields of gene therapy/drugs delivery to deliver macromolecules, monoclonal antibodies, nucleic acids (DNA, mRNA, siRNA), and anticancer drugs in to cells, tissues, and organs (Barreiro et al. 2009; Cochran and Wheatley 2013; Guo et al. 2007; Hayashi et al. 2009; Jordao et al. 2010; Ka et al. 2007; Kodama et al. 2010; Kotopoulis et al. 2013; Liao et al. 2012; Shen et al. 2008; Sheyn et al. 2008; Suzuki et al. 2010; Suzuki et al. 2008; Tinkov et al. 2010; Wang et al. 2013). However, additional efforts are needed to improve its efficiency, especially for in vivo applications (Kamimura et al. 2011).

2.3.4. Electroporation

Electroporation is the phenomenon that occurs when a cell is exposed to an electric field that exceeds a certain threshold on either a microsecond (i.e., pulsed

54 electric field or traditional electroporation) (Chen et al. 2010; Chen et al. 2006; Weaver 2003) or nanosecond time scale (i.e., nanosecond pulsed electric field) (Ibey et al. 2010; Vernier and Ziegler 2007). The electric field changes the electrochemical potential across the cell membrane and induces local instabilities by the formation of defects in one of the leaflets of the membrane (Chen et al. 2006; Granot and Rubinsky 2008; Piggot et al. 2011; Vernier and Ziegler 2007; Weaver 2003). The defect is a point in the membrane where the head groups of a few (usually around three to five) lipids are located closer to the centre of the membrane bilayer than other “bulk” membrane lipids (Piggot et al. 2011; Vernier and Ziegler 2007). This defect is accompanied by the insertion of water molecules into the membrane at this point, with the stabilization of water conformation taking place as a result of interactions with the protruding lipid head groups (Piggot et al. 2011). This is followed by the formation of a single-file water channel that spans the membrane, which is not accompanied by the entry of lipid head groups into the low dielectric region of the membrane (Piggot et al. 2011). After the formation of the water channel, lipid head groups are then observed to move into the core of the membrane, forming hydrogen-bonding interactions with the water molecules forming the channel (Piggot et al. 2011). Once the lipids have moved to stabilize the water channel, the pore rapidly increases in size, with the phospholipids lining the pore until the membrane collapses (Piggot et al. 2011).

It has been well-documented that cell membrane poration approach, based on electric fields (electroporation), has been developed to deliver therapeutic molecules to tissues and organs in vivo. A new cancer treatment modality, electrochemotherapy, has emerged (Belehradek et al. 1993; Gehl 2003; Gothelf et al. 2003; Mir 2001; Mir et al. 1991; Mir et al. 1998; Neumann et al. 1999; Rebersek et al. 2004; Serša et al. 1995; Serša et al. 1998). By using electric pulses, molecules that are otherwise non-permeant can gain direct access to the cytosol of cells. Highly toxic molecules such as bleomycin and cisplatin, once inside the cell, respectively, create single- and double-strand DNA breaks or adducts at very low extracellular concentrations (approximately 10-9 M) if the cell is electroporated, whereas under normal conditions the concentrations required would need to be several orders of magnitude higher to achieve the same level of cytotoxicity (Chen et al. 2006). This electrochemotherapy has been successfully used in clinical trials 55 for cancer treatment (Belehradek et al. 1993; Gehl 2003; Gothelf et al. 2003; Mir 2001; Mir et al. 1991; Mir et al. 1998; Neumann et al. 1999; Rebersek et al. 2004; Serša et al. 1995; Serša et al. 1998). Beside drugs, a wide range of potentially therapeutic agents, including proteins, oligonucleotides, RNA and DNA, can be introduced into the cells by an electrotransfer process (Bloquel et al. 2004; Chabot et al. 2013; Escobar-Chávez et al. 2009; Golzio et al. 2004; Miklavčič et al. 2000; Prausnitz et al. 1993; Scherman et al. 2002). In addition, cell membrane poration caused by electric fields can also induce different types of cells to fuse when they are in close contact, a phenomenon known as electrofusion (Neumann et al. 1980; Sukhorukov et al. 2005; Teissie et al. 1982; Usaj et al. 2013; Zimmermann 1982).

2.4. Functional roles of lipids in membranes

The primary role of lipids in cellular function is in the formation of the lipid bilayer, which is the permeability barrier of cells and subcellular organelles (Dowhan et al. 2016). The major lipid type which make up about 75% of almost all membranes is glycerol-based phospholipid (Dowhan et al. 2016). The other important lipids vary in their presence and amount depending on the organisms (Dowhan et al. 2016). For instant, the ceramide-based sphingolipids are present in all the eukaryotic membranes; whereas sterols are present in eukaryotic cytoplasmic membranes and in a few bacterial membranes (Dowhan et al. 2016). The major sterol of mammalian cells is cholesterol; whereas yeast contain ergosterol (Dowhan et al. 2016). Bacteria do not make sterols but some species incorporate sterols from the growth medium (Dowhan et al. 2016). Neutral glycerol-based glycolipids, also known as diacylglycerol (DAG) glycans, are major components in many Gram- positive bacterial and plant membranes, while Gram-negative bacteria utilise a glucosamine-based phospholipids, also known as saccharolipid (Lipid A), as a major structural component of the outer leaflet of the outer membrane (Dowhan et al. 2016).

The variety of hydrophobic tails of lipids results in additional diversity (Dowhan et al. 2016). In eukaryotes and eubacteria, these tails are saturated and unsaturated fatty acids or lesser amounts of fatty alcohols; whereas many Gram-

56 positive bacteria contain branched chain fatty acids (Dowhan et al. 2016). The headgroups of the glycerol-based phospholipids further classify lipids into phosphatidic acid (PA, with OH), phosphatidylcholine (PC), phosphatidylserine (PS), phosphatidylglycerol (PG), phosphatidylinositol (PI), and cardiolipin (CL) with the molecular structures as shown in Figure 2.2 (Dowhan et al. 2016).

It is thought that the membrane permeation is dependent on the membrane fluidity, which in turn is dependent on the hydrophobic fatty acid composition of the membrane lipids, cell microenvironment and the presence of charged phospholipid head groups (Lande et al. 1995; McLaughlin et al. 1970). Modulation of the membrane fluidity may arise due to the ease of movement of water molecules, and the dielectric constant of water, which is affected by the EMF (Shimanouchi et al. 2009). It has been reported that a temperature increase would cause an increase in the membrane fluidity, as confirmed by the diffusion of calcein molecules throughout the phosphatidylcholine bilayer membrane (Shimanouchi et al. 2009). Cells maintained their constant membrane fluidity by regulating the fatty acid composition and/or the phospholipid headgroups (Annous et al. 1997; Dowhan et al. 2016). It was suggested that charged phospholipid head groups developed a substantial potential at the lipid ˗ solution interface, influencing the concentration of ions at the interface and hence the permeability properties of the cell membrane (McLaughlin et al. 1970). Ca2+ and other divalent cations (Mg2+, Sr2+ but not Ba2+) can reduce the effective size of the negatively charged head groups of CL and PA, which give them nonbilayer properties (Dowhan et al. 2016). Low pH has a similar effect on the head group of PS (Dowhan et al. 2016). It has been shown that the microorganisms Acholeplasma laidlawii and Escherichia coli always try to maintain the physical state of their membrane lipids close to a bilayer–nonbilayer phase transition (Lindblom et al. 1993; Morein et al. 1996; Vikstrom et al. 2000). This allows the membrane to readily undergo temporal local rearrangements such as cell division, transbilayer transport of lipids and polar solutes, and membrane fusion (de Kruijff 1997; de Kruijff et al. 1985; Epand 1998; Siegel and Epand 1997; van den Brink-van der Laan et al. 2004).

Figure 2.2. The molecular structure of the glycerol-based phospholipids headgroups.

It was suggested that when E. coli mutants with membranes containing a high content of unsaturated fatty acids are grown at low temperature, they lyse when raised rapidly to high temperature (Dowhan et al. 2016). This is probably due to the increased membrane permeability of fluid membranes and a phase transition by PE and/or CL (Dowhan et al. 2016).

Since the fatty acid and phospholipid compositions vary between different cell types (Gunstone et al. 1994; Ratledge and Wilkinson 1989b), it is of considerable interest to understand whether exposure to 18 GHz EMF will induce cell permeability in typical representatives of the major microbial taxa possessing different compositions of membrane fatty acids and phospholipids.

Chapter 3. Experimental Design

3.1. Overview

As discussed in section 2.1.3, the biological effects of EMF on biological systems can be induced by changing the EMF frequency range, power, and time of exposure. In this chapter, the experimental design is described in detail, including the EMF exposure conditions, instrument set-up, dosimetry and controls. The Gram negative E. coli K12 and Gram positive S. aureus CIP 65.8T bacterial cells were selected as representative bacteria for investigation.

3.2. Experimental design

3.2.1. Temperature-dependent approach

Bacteria were subjected to both single and multiple exposures of EMF at two frequencies (18 GHz, 6.5 GHz), at a power of 17 W, within a sub-lethal temperature range of either 20 to 40°C or 20 to 33°C, depending on the taxonomic affiliation of the microbial species. These trials were conducted to determine the dose response characteristics and to obtain an insight into the biological effects that these EMF exposures would cause in the bacterial samples. Six combinations of exposure number and frequency (3 separate exposures at 2 different EMF frequencies) were conducted for each of the bacterial samples, with each of these combinations repeated independently 10 times to allow an appropriate statistical analysis to be performed. Samples were exposed to the EMF and then allowed to cool on ice in between exposures.

3.2.2. Power-variation approach

In this approach, bacteria were subjected to EMF exposures at a frequency of 18 GHz at different power levels (8, 9, 10, 15, 16 and 17 W) and a fixed duration (1 min). Experiments were conducted to determine the energy threshold at which 61 the EMF bio-effects on the bacterial samples commenced. Thirty-six combinations of exposure number and EMF power (6 treatments × 6 power levels) were conducted for each of the bacteria under investigation, with each of these combinations repeated independently 10 times to allow an appropriate statistical analysis to be performed. Samples were exposed to the EMF and then allowed to cool on ice in between exposures.

The bulk temperature rise of the bacterial suspension during EMF exposure was monitored and the specific absorption rate (SAR) was calculated as described in Section 3.3 below.

3.3. Instrument set-up and dosimetry

The exposure of the bacterial samples to the 18 GHz EMF was conducted in an LT 1500 resonant cavity device (Lambda Technologies Vari-Wave Model LT 1500) as shown in Figure 3.1. The Vari-Wave Model LT 1500 (Lambda Technologies) is capable of generating EMFs with a variable frequency, ranging from 6.5 to 18 GHz, which is well above the conventional frequency of a household microwave (2.45 GHz). The LT 1500 instrument is also known for being able to deliver an excellent level of EMF control, together with a uniformity of energy distribution. Moreover, using this instrument, both the amplitude and frequency of the EMF power could be varied, allowing a significant expansion of the parameter space within which the system could be optimized. A data logging option was also available, which allowed the processed data to be captured from the embedded computer system over a standard RS-232-C serial interface. The fiber optic probe within the microwave apparatus could be attached to the sample to monitor the internal temperature.

Figure 3.1. EMF apparatus. (a) The Vari-Wave Model LT 1500 (Lambda Technologies) EMF apparatus; (b) The EMF chamber containing the temperature control optic probes and (c) schematic diagram of the LT 1500 system.

A total volume of 2 mL of working suspension was transferred into a micro Petri dish (35 mm diameter, Griener Bio One, Frickenhausen, Germany). In order to achieve reproducible and reliable results, the identification of the area within the EMF chamber that provided a constant flow of radiation was critical. Therefore, to obtain a uniform temperature gradient in the multimode cavity and to avoid “hot spot” and “cold spot” standing wave effects, each micro Petri dish containing the cell suspensions was placed onto a ceramic pedestal (Pacific Ceramics Inc. PD160,

ε’ = 160, loss tangent < 10-3) in a position determined within the chamber using numerical EMF modelling from CST Microwave Studio 3D Electromagnetic Simulation Software, as shown in Figure 3.1 and Figure 3.2 (Nguyen et al. 2015; Shamis et al. 2009; Shamis et al. 2011; Shamis et al. 2008).

Figure 3.2. Modelling of the electric field and absorbed power using CST Microwave Studio 3D Electromagnetic Simulation Software. The EMF was uniformly distributed in the sample.

The bulk temperature rise of the bacterial suspension during EMF exposure was monitored by placing a built-in Luxtron Fiber Optic Temperature Unit (LFOTU) (1 mm diameter, LumaSense Technologies Inc., Santa Clara, CA, USA) at various positions, as shown in the Figure 3.3 (Nguyen et al. 2015). According to the manufacturer’s specifications, the tip of the probe is less than 1 mm thick and operates with an accuracy of ± 0.2 °C and with a 250 ms (in water) response time (Nguyen et al. 2015). The temperature was also confirmed using a portable infrared/thermal monitoring camera (Cyclopes 330S (Minolta, Osaka, Japan)) (Nguyen et al. 2015; Shamis et al. 2011). In total, 90 measurements were collected from 15 different positions in the Petri dish (five different locations (Figure 3.3b)

64 and three depth-levels (Figure 3.3c) of the cell suspension) (Nguyen et al. 2015). It was found that the temperature variation was small, and within the range ± 0.2 °C.

Over the 15 measurement locations, the specific absorption rate (SAR) in the medium was calculated using equation (Equation 3.1) (Nguyen et al. 2015; Panagopoulos et al. 2013).

�� × SAR = c �� |t = 0 (Equation 3.1)

�� where c is the specific heat capacity of the medium, and ��|t=0 is the time derivative of the temperature determined at t = 0 s.

Figure 3.3. Temperature control during EMF exposures. (a) overview of the LFOTU, micro Petri dish and the cell suspension layer; (b) Top view of the Petri dish with five different positions of measurement in the LFOTU; (c) Side view of the cell suspension, showing the three measured depth-levels of the LFOTU.

3.4. Heat treated controls

In order to ensure that any effects observed in the biological samples resulting from their exposure to EMF were not solely a result of thermal heating, a heat treated control sample was used. A Peltier plate heating/cooling system (TA Instruments) was employed to replicate the bulk temperature profiles experienced by the studied cells during the EMF exposures (Nguyen et al. 2015; Shamis et al. 2011). A 2 mL volume of working suspension was applied directly onto the Peltier plate sample platform, and subjected to the same bulk temperature change profiles experiences during the EMF exposures, as demonstrated in Figure 3.4 (Nguyen et al. 2015; Shamis et al. 2011).

Figure 3.4. The bulk temperature profiles of EMF exposed and Peltier plate heat-treated cell suspensions.

The diameter of the Peltier plate sample platform was 65 mm and the cell suspension layer thickness was calculated to be 0.6 mm, as shown in Figure 3.5.

All Peltier plate heated samples were performed in triplicate, and in parallel with the EMF exposure experiments. The bulk temperature changes during heating and cooling were monitored using a portable infrared/thermal monitoring camera (Cyclopes 330S).

Figure 3.4. Peltier heating stage (diameter = 65 mm, suspension depth = 0.6 mm) with position used for applying the cell suspension.

3.5. Negative controls

Working bacterial suspensions that were not exposed to either EMF exposures or heat treatment were used as the negative controls for all experiments.

Chapter 4. Materials and Methods

4.1. Overview

This chapter reports the materials and methods employed for the study of the biological effects of exposing bacterial cells to 18 GHz EMF radiation, with an emphasis being placed on investigating the effects on membrane permeability, cell morphology and viability. Various cell types, including prokaryotes and eukaryotes were studied.

4.2. Sample preparation

4.2.1 Bacterial and yeast strains, growth and maintenance

The taxonomic affiliation of all selected bacteria and yeast cells used in this study is shown in Table 4.1. The detailed description of each cell type is described in Section 4.2.2.

Bacterial and yeast cells were obtained from the American Type Culture Collection (ATCC, USA), the Culture Collection of the Pasteur Institute (CIP, France), and the Collection of Marine Microorganisms (KMM, Russian Federation). Pure cultures were stored at -80 ˚C in suitable medium, which are marine broth (MB, Becton Dickinson, Sparks, Nevada, USA) for marine strains, nutrient broth (NB, Oxoid Ltd., Basingstoke, Hampshire, England) for the rest of bacteria, and potato dextrose broth (PDB, Becton Dickinson) for yeast cells, with the addition of 20% (v/v) of glycerol (Sigma Aldrich, St. Louis, Missouri, USA). All strains were routinely grown on nutrient agar (NA, Oxoid Ltd.), brain-heart infused agar (BHIA, Becton Dickinson), marine agar (MA, Becton Dickinson), or potato dextrose agar (PDA, Becton Dickinson), depending on the requirements of a particular strain.

Table 4.1. Taxonomic affiliation of bacterial and yeast strains employed in this study. Cell types Taxonomic affiliation Branhamella Bacteria; Proteobacteria; Gammaproteobacteria; catarrhalis Pseudomonadales; Moraxellaceae; Moraxella; ATCC 23246 Branhamella Bacteria; Proteobacteria; Gammaproteobacteria; Escherichia coli Enterobacteriales; Enterobacteriaceae; K 12 Escherichia Bacteria; Actinobacteria; Actinobacteria; Kocuria rosea Actinobacteridae; Micrococcales; CIP 71.15T Micrococcaceae; Kocuria Planococcus Bacteria; Firmicutes; Bacilli; Bacillales; maritimus Planococcaceae; Planococcus KMM 3738 Staphylococcus Bacteria; Firmicutes; Bacilli; Bacillales; aureus Staphylococcaceae; Staphylococcus ATCC 25923 Staphylococcus Bacteria; Firmicutes; Bacilli; Bacillales; aureus Staphylococcaceae; Staphylococcus CIP 65.8T Staphylococcus Bacteria; Firmicutes; Bacilli; Bacillales; epidermidis Staphylococcaceae; Staphylococcus ATCC 14990T Bacteria; Actinobacteria; Actinobacteria; Streptomyces Actinobacteridae; Streptomycetales; griseus Streptomycetaceae; Streptomyces; Streptomyces ATCC 23915 griseus group; Streptomyces griseus subgroup Eukaryota; Opisthokonta; Fungi; Dikarya; Saccharomyces Ascomycota; saccharomyceta; Saccharomycotina; cerevisiae Saccharomycetes; Saccharomycetales; ATCC 287 Saccharomycetaceae; Saccharomyces Prior to each experiment, each strain was grown up to the stationary phase of growth as confirmed by growth curves (data not shown) at 25 ˚C (B. catarrhalis, K. 71 rosea, P. maritimus, Streptomyces griseus), 30 ˚C (Saccharomyces cerevisiae) or 37 ˚C (E. coli, Staphylococcus aureus, Staphylococcus epidermidis).

4.2.2. Selected cell types

4.2.2.1. Gram negative bacteria

4.2.2.1.1. Branhamella catarrhalis ATCC 23246

Initially described as Micrococcus catarrhalis, this species was then reclassified and assigned to the genus Neisseria (Catlin 1990). In 1970, the genus Neisseria was transferred to the genus Branhamella (Catlin 1970). B. catarrhalis is an aerobic, nonmotile Gram-negative diplococcus opportunistic human pathogen, which is commonly found in the upper respiratory tract of humans. The bacterium can grow well at temperatures as low as 22 °C (Catlin 1990). Cells are kidney bean shaped, with a diameter of 0.6 to 1.0 µm, often appearing in pairs or as tetrads (Murphy 1996). B. catarrhalis is saccharolytic, DNase, oxidase and catalase- positive with butyrate esterase activity (Verduin et al. 2002). Colonies grown on on blood agar are non-hemolytic round, opaque, convex and greyish white cells (Verduin et al. 2002). B. catarrhalis can cause respiratory infections, acute otitis media, sinusitis and infections such as endocarditis, meningitis and bacterial tracheitis (Verduin et al. 2002). The bacterium is β-lactamase positive and resistant to amoxicillin, ampicillin, piperacillin, and penicillin (Verduin et al. 2002). B. catarrhalis is capable of producing capsules like Neisseria gonorrhoeae, visible using electron microscopy as a fibrillar coat (Catlin 1990).

4.2.2.1.2. Escherichia coli K 12

E. coli was described in 1885 by German paediatrician Theodor Escherich (Hacker and Blum-Oehler 2007). E. coli is a Gram negative, facultatively anaerobic, rod-shaped bacterium, which is commonly found in the lower intestine of warm-blooded organisms (Madigan et al. 2003). Some strains are motile with flagella in a peritrichous arrangement (Darnton et al. 2007). The bacterium can survive over the temperature range 7 to 49 °C, with an optimum growth temperature of 37 °C (Herendeen et al. 1979). E. coli cells are about 2.0 μm long and 0.25 to 1.0 μm in diameter, with a cell volume of 0.6 to 0.7 μm3 (Kubitschek 1990). E. coli is oxidase negative, catalase positive, and reduces nitrates (Madigan et al. 2003). Enteric E. coli can be classified into six categories based on its virulence properties, such as enterotoxigenic E. coli (ETEC), enteropathogenic E. coli (EPEC), enteroinvasive E. coli (EIEC), enterohemorrageic E. coli (EHEC), enteroadherent aggregative E. coli (EAggEC), and verotoxigenic E. coli (VTEC) (Johnson 2002). Cultivated strains (such as the E. coli K 12) have adapted well to the laboratory environment, and lost their ability to thrive in the intestine. The bacterium can be easily manipulated and inexpensively grown; be a very versatile host for the production of heterologous and recombinant proteins which make E. coli being one of the best-studied prokaryotic model organisms in biotechnology and microbiology (Perna et al. 2002). This strain was obtained from the American Type Culture Collection (USA).

4.2.2.2. Gram positive bacteria

4.2.2.2.1. Kocuria rosea CIP 71.15T

Originally, K. rosea was named Micrococcus roseus by Flugge in 1886 (Stackebrandt et al. 1995). In 1995, Stackebrandt et al. split the genus Micrococcus

73 into four genera, Dermacoccus, Kocuria, Kytococcus, and Nesterenkonia. Therefore Kocuria species are placed in the genus Kocuria of the family Micrococcaceae (Stackebrandt et al. 1995). K. rosea is an aerobic, non-motile, non- encapsulated, non-sporulated Gram-positive coccus opportunistic human pathogen, which commonly found on the surface of human skin, mucous membranes, in the oral cavity, and the outer ear canal (Purty et al. 2013; Stackebrandt et al. 1995). The bacterium can also be found in freshwater, saltwater, and soil environments (Purty et al. 2013; Stackebrandt et al. 1995). Cells have a diameter of approximately 1.0 to 1.8 µm and occur in pairs, tetrads or clusters (Stackebrandt et al. 1995). K. rosea is non-haemolytic, catalase positive, coagulase negative with an optimum growth temperature of 25 to 37 °C (Stackebrandt et al. 1995). Colonies of K. rosea are slightly convex, smooth, and red and/or pink in colour (Stackebrandt et al. 1995). It was previously reported that K. rosea can cause catheter-related bacteremia and peritonitis (Purty et al. 2013).

4.2.2.2.2. Planococcus maritimus KMM 3738

P. maritimus bacteria were described by Yoon in 2003 (Yoon et al. 2003). P. maritimus is an aerobic, motile with a single polar flagellum, Gram-positive coccus which isolated from sea water of a tidal flat in Korea and from the degradation of the brown alga Fucus evanescens thalluses (Ivanova et al. 2006; Yoon et al. 2003). Cells have a diameter of approximately 1.0 to 1.4 µm (Yoon et al. 2003). P. maritimus produces carotenoid pigments, is chemo-organotrophic, alkaliphilic, halo-tolerant, and grows well on nutrient media containing up to 17% NaCl over a temperature range 5 to 45 °C (Ivanova et al. 2006; Yoon et al. 2003). Colonies are smooth, glistening, low-convex, circular and yellow to orange in colour (Yoon et al. 2003).

4.2.2.2.3. Staphylococcus aureus ATCC 25923 and CIP 65.8T

In 1880, Scottish surgeon Sir Alexander Ogston first described the masses of staphylococci like bunches of grapes of bacterial cells in pus from a surgical abscess in a knee joint (Cowan et al. 1954). Then in 1884, German physician Friedrich Julius Rosenbach differentiated these bacteria by the colour of their colonies into Staphylococcus aureus (Latin, aurum for gold) and Staphylococcus albus (Latin, for white) (Cowan et al. 1954; Harris et al. 2002). Staphylococcus aureus is a facultative anaerobic, nonmotile Gram-positive coccus which is frequently found in the respiratory tract of humans (Cowan et al. 1954). Cells are 1 µm in diameter and appears as grape-like clusters (Cowan et al. 1954). The bacterium is haemolytic, catalase positive, coagulase negative with a growth temperature range from 4 °C to 46 °C, and an optimum growth temperature of 37 °C (Harris et al. 2002). Staphylococcus aureus causes various infections such as highly contagious skin infections (boils and abscesses), meningitis, osteomyelitis, pneumonia, septic phlebitis, endocarditis, and toxic shock syndrome by release of super-antigens into the blood stream (Harris et al. 2002). Infections acquired outside hospitals can usually be treated with penicillinase-resistant β-lactams; whereas the hospital acquired infection is often caused by antibiotic resistant strains (MRSA) and can only be treated with vancomycin (Harris et al. 2002). The Staphylococcus aureus ATCC 25923 and CIP 65.8T strains were obtained from the American Type Culture Collection (USA), and the Culture Collection of the Pasteur Institute (France), respectively.

4.2.2.2.4. Staphylococcus epidermidis ATCC 14990T

After being differentiated from Staphylococcus aureus by Rosenbach, the bacterium Staphylococcus albus was later renamed Staphylococcus epidermidis

75 because of its main presence on human skin (Cowan et al. 1954; Harris et al. 2002). Similar to Staphylococcus aureus, the Staphylococcus epidermidis is also a facultative anaerobic, non-motile Gram-positive coccus with a diameter from 0.5 to 1.5 µm, is catalase positive, coagulase negative, and appears as grape-like clusters; but is not haemolytic (Otto 2009). Cells grow well within the temperature range 15 to 45 °C with an optimum growth temperature of 26 to 37 °C (Bergey and Holt 1994). As part of the human epithelial microflora, Staphylococcus epidermidis is believed to provide a probiotic function by preventing colonization of other pathogenic bacteria; but it is nowadays also seen as an important opportunistic pathogen due to its ability to form biofilms on the skin (Otto 2009). Staphylococcus epidermidis was reported to be the most frequent cause of nosocomial infections (Otto 2009). In addition to methicillin resistance, Staphylococcus epidermidis strains have acquired resistance to several other antibiotics, including rifamycin, fluoroquinolones, gentamicin, tetracycline, chloramphenicol, erythromycin, clindamycin, and sulphonamides (Otto 2009). Very rarely, there is resistance to streptogramins, linezolid, and tigecycline (Otto 2009). There is evidence suggesting that methicillin resistance cassettes were transferred from Staphylococcus epidermidis to Staphylococcus aureus (Otto 2009).

4.2.2.2.5. Streptomyces griseus ATCC 23915

Streptomyces griseus was first isolated from soil by Krainsky in 1914 but was named in 1915 by Dr Selman A. Waksman, a microbiologist at the Agricultural Department of Rutger’s University, and colleagues (Waksman et al. 1948). Streptomyces griseus was initially named as Actinomyces griseus and reclassified as Streptomyces griseus in 1943 (Waksman et al. 1948). S. A. Waksman together with Schatz and Bugie, found the antibiotic “streptomycin” produced by a variant of Streptomyces griseus to be particularly effective against the tuberculosis bacteria, Tubercle bacillus (Waksman et al. 1948). Feldman and Hinshaw, two physicians from the Mayo Clinic in Rochester, found streptomycin to be effective in curing two extreme classes of tuburculosis: tuberculous meningitis and military tuberculosis (Waksman et al. 1948). Streptomyces griseus is non-motile, aerobic,

Gram-positive filamentous, spore forming rod-shaped bacterium which commonly found in soil and sometimes in deep sea sediment (Waksman et al. 1948). Streptomyces griseus is alkaliphilic, produces an aerial mycelium, which has modes of branching that eventually leads the hyphae to form chains of spores called arthospores (McCormick and Flärdh 2012; Waksman et al. 1948). When grown as colonies, Streptomyces griseus produce grey spore masses and grey-yellow reverse pigments (Waksman et al. 1948). The optimum temperature for cell growth is in the range 25 to 35 °C (Waksman et al. 1948). Considered relatively harmless to human, Streptomyces griseus produces many useful secondary metabolites such as enzyme inhibitors and comprise 70% of naturally-occurring antibiotics (McCormick and Flärdh 2012).

4.2.2.3. Unicellular eukaryote

4.2.2.3.1. Saccharomyces cerevisiae ATCC 287

The yeast Saccharomyces cerevisiae has been known and used since ancient times in the fermentation process that converts sugar into alcohol, and the baking process as a leavening agent (Mortimer 2000). It was firstly isolated by Emil Mrak from rotten figs found in Merced, California (Mortimer and Johnston 1986). Natural yeast strains can be found on the surfaces of plants, the gastrointestinal tracts, body surfaces of insects and warm-blooded animals, soils from all regions of the world and even in aquatic environments (Mortimer 2000). Most often it is found in areas where fermentation occurs, such as the on the surface of fruit, storage cellars and on the equipment used during the fermentation process (Mortimer 2000). The optimum growth temperature is in the range 30 to 35 °C (Feldmann 2012). Saccharomyces cerevisiae cells are round to ovoid, 5 to 10 μm in diameter, grow as two forms, which are haploid and diploid (Feldmann 2012). Under normal conditions, both forms grow and reproduce by mitosis process known as budding (Feldmann 2012). Under stress conditions, only the diploid form undergoes sporulation, entering meiosis and producing four haploid spores, which can

77 subsequently mate to form daughter cells (Feldmann 2012). Saccharomyces cerevisiae is considered to be a "model organism" for scientists because it has a fast rate of growth, being both a unicellular and eukaryotic organism (Mortimer 2000). It is not a pathogenic but can cause vaginal yeast infections, and induce irritation in people with Crohn's disease, an autoimmune disorder (Main et al. 1988).

4.2.2.4. Red blood cells

Red blood cells were isolated from a 16 week-old New Zealand white rabbit’s fresh blood. The rabbit was kept at Monash University’s animal house under approval from Monash University’s Animal Ethics Committee. The whole blood was bled into plastic Vacutainer tubes spray-coated with K2EDTA (Becton Dickinson) and inverted several times to prevent blood clotting. After bleeding, the Vacutainer tubes were disinfected with 70% ethanol then put in a sealed plastic bag and transferred back to Swinburne University for further extraction. In order to ensure an appropriate aseptic cell suspension, all operations were carried out in a bio-safety cabinet class II (Thermo Fisher Scientific, Waltham, Massachusetts, USA). The fresh blood was gently homogenized with phosphate buffer saline (PBS) 10 mM, pH 7.4 at 1:1 ratio. The blood mixture was then carefully overlaid on top of a Histopaque 1077 solution (Sigma Aldrich) at 3:1 ratio and centrifuged at 400 g for 20 min in a Falcon tube (Becton Dickinson). The red blood cells layer at the bottom of the Falcon tube was recovered and the rest was discarded.

4.3. Sample preparation

Working cell suspensions were freshly prepared for each independent experiment. The bacterial cell density was adjusted to OD600 ≈ 0.1 using 10 mM phosphate buffered saline (PBS) solution at pH 7.4 using a spectrophotometer (Dynamica Halo RB-10 UV-Vis, Precisa Gravimetrics AG, Dietikon, Switzerland). The cell density of the yeast Streptomyces cerevisiae ATCC 287 cells, erythrocytes and PC-12 cells were also adjusted to 1 × 106 viable cells mL-1, 5 × 105 viable cells

78 mL-1 and 3 × 105 viable cells mL-1, respectively, using 10 mM PBS solution using the Neubauer-improved haemocytometer. The working suspensions were subjected to EMF exposures and heat treatments as described in Chapter 3. Untreated working suspensions were used as untreated controls.

4.4. Bacterial cell viability

Initially, 100 µL of the working suspension was mixed with 900 µL of 10 mM PBS solution to make a 1:10 dilution suspension. Further 1:10 dilutions were then performed using the same method until a tenth 1:10 serial dilution was obtained. All of the suspensions were streaked on suitable agar plate using a volume of 100 µL of each diluted suspension per plate and then incubated at suitable temperature and time to allow sufficient growth, depending on the requirements of a particular strain. After inspection of the plates, it was determined that each working suspension should be subjected to four 1:10 serial dilution steps (dilution factor 104) and one 1:2 dilution step to obtain the recovery of about 500 colony- forming units (cfu) from 100 µL of bacterial suspension spread per plate, assuming that each viable cell would give rise to a single colony. This optimised dilution process was applied to each of the working bacterial suspensions after EMF exposures and heat treatments to determine the degree of cell viability using Equation 4.1.

CFU formed after EMF exposures(or heat treatments) Viability (%) = × 100 CFU formed of untreated suspension (Equation 4.1).

4.5. Scanning electron microscopy

Analysis of the cells was performed using Scanning electron microscopy (SEM) in order to identify any morphological change taking place as a result of heating or EMF exposures, and to determine whether these changes were reversible.

4.5.1. Sample preparation for analysis of cell morphology

A field emission scanning electron microscope FeSEM – ZEISS SUPRA 40VP (Carl Zeiss, Jena, Germany) with primary beam energy of 3 kV was used to obtain high-resolution images of the morphology of the cells. A 100 µL aliquot of cell suspension was placed on a glass cover slip (ProSciTech, Kirwan, Australia) in triplicate for each condition (untreated control, Peltier (conventional) heat treated control, EMF exposed sample). After one minute, the glass cover slips were washed -1 with MilliQ H2O (with a resistivity of 18.2 M cm ), dried with 99.99% purity nitrogen gas, then subjected to gold sputtering (6 nm thick gold film) using a NeoCoater MP-19020NCTR (JEOL, Tokyo, Japan). The remainder of the samples, which were left to stand for 9 and 10 min, were subjected to the same preparation process. Approximately ten SEM images were obtained at 5,000× and 70,000× magnifications for subsequent analysis.

4.5.2. Sample preparation for cell rigidity analysis

Clanger cicada (Psaltoda claripennis) specimens were collected from the greater Brisbane, Australia parkland areas (typically on flora such as eucalypts). The wings exhibiting nanopillars with dimensions of 200 nm tall, 100 nm in diameter at the base, 60 nm in diameter at the cap, and spaced 170 nm apart from center to center, were employed to observe the interactions between EMF exposed Gram positive cocci and nanopillars. All experiments were performed on the region

80 between veins C (Costa) and M (Media), and the branches on the dorsal side of the forewing as shown in Figure 4.1, for consistency.

Figure 4.1. Structural physiology of the forewing of Psaltoda claripennis, with major areas indicated. The cutting areas for experiment were all the M areas and C areas.

The wings were cut into pieces of approximately 0.5 cm × 0.5 cm and were attached by conductive adhesive tapes onto circular metallic discs. The latter was briefly rinsed with copious nanopure H2O and dried with nitrogen gas. Immediately following EMF exposures, the Gram positive cocci were seeded onto the wings and left for 10 minutes. The samples were then washed with nanopure H2O, dried with nitrogen gas and sputter-coated with 10 nm of gold (Ivanova et al. 2010). The negative control for this experiment consisted of untreated bacterial suspension that was seeded onto the wings for 10 minutes and then subjected to the same analysis. Approximately 40 SEM images taken at ×5000 and ×70000 magnifications for each sample were analysed.

4.6. Confocal laser scanning microscopy

Confocal laser scanning microscopy (CLSM) was employed in order to determine whether the cell membranes had been permeabilized as a result of exposure to the EMF radiation. These studies were performed using a Fluoview FV10i-W inverted microscope (Olympus, Tokyo, Japan) with several fluorescent probes, as described below.

Propidium iodide (1.0 mg mL-1 solution in water; Life Technologies Australia, Mulgrave, Australia) was used at a concentration of 500 nM to determine the viability of the bacterial cells 33. Untreated cells, cells inactivated by boiling (100 °C) and Peltier heat treated cells (40 °C) were used as three different types of control. Heat inactivated cells were prepared by boiling the bacterial suspension for 15 min, followed cooling in a 25 °C water bath for 30 min. PI was added to all the bacterial suspensions at 1 min and 10 min after EMF exposures. The PI remained in contact with the bacteria throughout the experiment.

Two types of fluorescent, hydrophilic, neutrally charged silica nanospheres, 23.5 ± 0.2 nm (FITC) and 46.3 ± 0.2 nm (Rhodamine B) (Corpuscular, Cold Spring, NY, USA) were used. These silica nanospheres were selected because hydrophilic nanoparticles rarely freely translocate through an intact lipid bilayer (Pogodin et al. 2012) and neutrally charged nanoparticle surfaces prevent any nonspecific interactions taking place with the membrane (Verma and Stellacci 2010). The nanospheres were added to the cell suspensions 1, 9 and 10 min after the EMF exposure at a concentration of 25 µg mL-1, incubated for 10 min then washed twice by centrifugation at 4500 rpm for 5 min. The controls for these experiments comprised of an untreated cell suspension mixed with the fluorescent silica nanospheres and Peltier heat treated cell suspensions (40 °C) that was mixed with the fluorescent silica nanospheres in parallel with the EMF exposed samples.

A 100 µL aliquot of each sample was then analyzed using a Fluoview FV10i- W inverted microscope. Approximately 10 CLSM images were obtained for each sample, with each containing at least 10 bacterial cells per image.

4.7. Quantification of intake nanospheres

4.7.1. Experimental procedure

This section of the study was designed to evaluate the approximate number of fluorescent silica nanospheres that could be taken into the cytosol of the target bacteria after exposure to EMF radiation. A POLARstar Omega microplate reader (BMG Labtech) was used to measure the fluorescence intensity of nanospheres in each bacterial suspension for use in CLSM analysis.

A standard curve was also constructed to determine the correlation of fluorescence intensity as relative fluorescence units (RFU) as a function of the concentration of silica nanospheres. A total of seven nanosphere concentrations were prepared (0.005, 0.05, 0.5, 1, 5, 19 and 15 µg mL-1). Approximately 3 measurements were conducted for each type of nanosphere, with the average result being taken.

4.7.2. Theoretical consideration

The mass of a silica nanosphere was determined using Equation 4.1.

  Vm (Equation 4.1) where m is the mass of a silica nanosphere (g), ρ is the mass density of silica (g cm-3) and V is the volume of a silica nanosphere (cm-3).

The volume of a silica nanosphere can be calculated using Equation 4.2. 4  rV 3 3 (Equation 4.2)

83 where V is the volume of a silica nanosphere (cm3) and r is the radius of a silica nanosphere (cm).

The radii of the 23.5 nm and 46.3 nm nanospheres are 11.75 × 10−7 cm, 23.15 × 10−7 cm, respectively (Corpuscular Inc.). According to Equation 4.2, the volumes of the 23.5 nm and 46.3 nm silica nanosphere were 6.8 × 10−18 cm3 and 5.2 × 10−17 cm3, respectively. By substituting the volumes of each nanosphere into Equation 4.1, the masses of each were determined to be 1.8 × 10−17 g and 1.38 × 10−16 g, respectively. The mass of a single nanosphere could be used to calculate the total number of nanosphere remaining in the cytosol of the bacterial cells.

4.8. Transmission electron microscopy

Transmission electron microscopy (TEM) analysis was conducted in order to reconfirm the cell permeabilization phenomenon and determine locations of nanospheres relatively to the cell membranes after EMF exposures.

4.8.1. Grids and samples preparation

4.8.1.1. Bacteria and yeast

The cell suspensions after the addition of the 23.5 nm nanospheres were pelleted by centrifugation at 4800 rpm for 5 min at 25 °C. The cells were then washed twice with phosphate buffer saline (PBS, 10 mM, pH 7.4) in order to remove any unbound nanospheres. The pellets were then suspended in 2 mL of 4% glutaraldehyde in PBS for 30 min, and washed twice in PBS for 5 min. After the final washing step, the cells were mixed thoroughly in 0.5 mL of 5% agarose gel by stirring. The agar was immediately cooled to 4 °C by refrigeration for 30 min, 3 then cut into 1 mm cubes and fixed with 1 mL of 1% osmium tetroxide (OsO4) for

1 h. The cubes were washed twice in nanopure H2O (with resistivity of 18.2 MW

84 cm-1) for 15 min. The cells were dehydrated by passing the cubes through a graded ethanol series (20%, 40%, and 60%) (2 mL) for 15 min and stained for 8 h with 2% uranyl acetate in 70% ethanol (2 mL). After staining, the cells were further dehydrated by passing the cubes through another graded ethanol series (80%, 90% and 100%) for 15 min (2 mL) (Dekiwadia et al. 2012; Zhao et al. 2011).

The embedding medium was prepared using araldite, dodecenyl succinic anhydride (DDSA), and benzyldimethylamine (BDMA) (ProSciTech) and stirred thoroughly(Luft 1961). In order to embed the samples, each cube was washed twice with 100% acetone (2 mL) for 20 min, then incubated in 2 mL of acetone and embedding medium (1:1 ratio) for 8 h, followed by transfer to acetone and embedding medium (1:3 ratio) for 8 h and finally transferred into the pure embedding medium for 8 h. The cube was then transferred into an embedding mould containing fresh pure embedding medium, which was then polymerized for 24 h at 60 °C (Mu et al. 2012).

4.8.1.2. Erythrocytes

The cell suspensions were added with 46.3 nm nanospheres and process similarly to the protocol for bacteria and yeast described above with some modification. Briefly, the cell suspensions with nanospheres were pelleted by centrifugation at 1300 rpm for 5 min at 25 °C. The cells were then washed twice with phosphate buffer saline (PBS, 10 mM, pH 7.4) in order to remove any unbound nanospheres. The pellets were then suspended in 2 mL of 1% glutaraldehyde in PBS for 30 min, and washed twice in PBS for 5 min. After the final washing step, the cell suspensions were added to 0.5 mL of molten 3% agarose gel by stabbing with a pipette tip. The agar was immediately cooled to 4 °C by refrigeration for 30 min, then trimmed down to the area containing cell suspension and fixed with 1 mL of

1% osmium tetroxide (OsO4) for 1 h. The agars containing cell suspension were -1 washed twice in nanopure H2O (with resistivity of 18.2 MW cm ) for 15 min. The cells were dehydrated by passing the agars through a graded ethanol series (20%, 40%, and 60%) (2 mL) for 15 min and stained for 8 h with 2% uranyl acetate in

70% ethanol (2 mL). After staining, the cells were further dehydrated by passing the agars through another graded ethanol series (80%, 90% and 100%) for 15 min (2 mL).

The embedding medium was prepared using LR gold resin (ProSciTech). In order to embed the samples, each agar containing cell suspension was incubated in 2 mL of 100% ethanol and LR gold monomer (1:1 ratio) for 8 h, followed by transfer to 100% ethanol and LR gold monomer (1:3 ratio) for 8 h and finally transferred into the pure LR gold monomer for 8 h. Each agar was then transferred into a gelatin capsule containing fresh LR gold monomer mixed with 1% of dry benzoyl peroxide, which was then polymerized for 24 h at 4 °C.

4.8.1.3. Sectioning and examination

The final block was trimmed, then cut into ultrathin sections (70 nm thickness) using a Leica EM UC7 ultramicrotome (Leica Microsystems, Wetzlar, Germany) with a diamond knife (Diatome, Pennsylvania, USA). Sections were placed onto 200 mesh copper grids and examined using a JEM 1010 instrument (JEOL). Approximately 40 TEM images taken at ×5000 and ×10000 magnifications for each sample were analysed.

4.9. Statistical analysis

All statistical data processing was performed using the SPSS 21.0 software (SPSS Inc., Chicago, Illinois, USA) and Microsoft Office Excel 2010.

Chapter 5. EMF effects on Gram negative bacterial cells

5.1. Overview

This chapter reports the biological effects resulting from exposure of Gram negative bacterial cells to 18 GHz EMF, with an emphasis being placed on investigation of the radiation effects on membrane permeability, cell morphology and viability. Branhamella catarrhalis ATCC 23246 (B. catarrhalis) and Escherichia coli ATCC 15034 (E. coli) bacterial cells were selected as representative samples of Gram negative bacteria. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1 that resulted in temperature increases ranging from 20 to 40ºC (at a heating rate of 20ºC per min). After exposure, samples were allowed to cool to 20°C on ice (at a rate of 10ºC per min) between exposures.

5.2. Background

To date, an abundant amount of research has reported the existence of certain bio-effects resulting from exposing Gram negative E. coli cells (as a representative species) to high frequency EMF radiation in the microwave (MW) region. The bioeffects that result from these exposures were found to be dependent on the strength, frequency, and time of interaction of the EMF radiation (Banik et al. 2003; Culkin and Fung 1975; Fujikawa et al. 1992; Rai et al. 1999; Shamis et al. 2012b; Shamis et al. 2011; Woo et al. 2000; Zhou et al. 2010). Some of the observed bio-effects likely arose because of the heat generated through the exposure, however many studies have also provided evidence of specific effects taking place in the cell membranes that cannot be explained solely by the bulk temperature increases (Banik et al. 2003; Barnabas et al. 2010; Dreyfuss and Chipley 1980; Geveke and Brunkhorst 2008; Shamis et al. 2009; Shamis et al. 2008).

Previous studies have reported that exposing E. coli cells to 18GHz EMF radiation prompted changes in the cell membrane that allowed the internalization

88 of large macromolecules such as dextran (150 kDa) and an increase in the enzyme activity levels of lactate dehydrogenase and cytochrome c oxidase (Shamis et al. 2012b; Shamis et al. 2011). Such effects were not observed in the thermally-heated control group (Shamis et al. 2012b; Shamis et al. 2011).

Since only E. coli cells were studied in these previous works, the aim of the current research was to investigate the membrane permeability, cell morphology and viability of an additional Gram negative bacterial type, B. catarrhalis, after being exposed to 18 GHz EMF radiation. Different exogenous materials were also used to test whether the EMF exposure resulted in membrane permeability. These materials included propidium iodide and two types of silica nanospheres (23.5 and 46.3 nm in diameter). A propidium iodide assay was used as the standard technique to confirm electroporation (formation of pores) of the bacterial membranes (Nesin et al. 2011; Pakhomova et al. 2011).

Propidium iodide does not normally pass through intact membranes (Nesin et al. 2011; Pakhomova et al. 2011), however, when a cell membrane is disrupted, the propidium cation (Pr2+) can pass through the membrane and bind to the nucleic acids within the cell, eventually fluorescing (Nesin et al. 2011). These types of silica nanospheres were selected because translocation of these nanoparticles (being hydrophilic and spherical) through a lipid bilayer is not usually favourable (Pogodin et al. 2012). It has also been previously reported that neutrally charged nanoparticle surfaces prevent any nonspecific interactions taking place within the membrane (Verma and Stellacci 2010).

The specific absorption rate (SAR) was determined experimentally because it has been suggested that it is a more accurate estimation of the energy absorbed by biological material (Gabriel et al. 1996; Haemmerich et al. 2005; Panagopoulos et al. 2013). This is because any variation in the specific heat within a sample of biological matter is usually much smaller than the corresponding variation in conductivity, resulting in a much more uniform temperature than that determined using an electric field distribution approach (Gabriel et al. 1996; Haemmerich et al. 2005; Panagopoulos et al. 2013).

5.3. Results

5.3.1. EMF effects on cell membrane permeability

The level of cell membrane permeability in the bacteria exposed to the EMF radiation was determined using confocal laser scanning microscopy (CLSM). This analysis demonstrated that membrane permeability was present in both of the Gram-negative strains tested, as confirmed by the uptake of propidium iodide 1 minute after the EMF exposure. The results are presented in Figure 5.1. The permeability effect lasted for up to 10 minutes, since no propidium iodide was observed to be taken into any of the bacterial cells 10 minutes after exposure (third row). The phase contrast micrographs show the bacterial cells in the same field of view (second and fourth rows).

As a control, bacterial suspensions were heated to 40°C using a Peltier plate heating system. For these heat control groups, no internalization of propidium iodide by the bacterial cells was observed (first and second rows) in Figure 5.2. Cells inactivated by boiling (100°C) were seen to internalize the propidium iodide (third and fourth rows).

Figure 5.1. Internalization of propidium iodide by Gram-negative bacterial cells after EMF exposure. Row 1: CLSM images showing the internalization of propidium internalization 1 min after EMF exposure. Row 3: no propidium iodide was observed to have been internalized in any tested cell types after 10 min. Row 2 and 4: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all fluorescence images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg- 1.

Figure 5.2. Internalization of propidium iodide by the heat control groups. Row 1: CLSM images showing that no propidium iodide had been internalized by the Peltier heat treated cells (40°C). Row 3: internalization of propidium iodide was observed in the heat inactivated cells (boiling, 100°C). Row 2 and 4: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all fluorescence images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

Untreated cells did not internalize the propidium iodide, as expected, since no membrane permeation was present, as shown in Figure 5.3.

Figure 5.3. No propidium iodide internalization by the untreated control. Row 1: CLSM images showing that no propidium iodide was internalized by untreated bacterial cells. Row 2: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all fluorescence images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg- 1.

Figure 5.4. Internalization of 23.5 nm and 46.3 nm nanospheres by the Gram-negative bacterial cells after EMF exposure. Row 1: CLSM images showing the internalization of 23.5 nm nanospheres (in green) by the EMF exposed cells. Row 2: internalization of 46.3 nm nanospheres was observed in the EMF- exposed cells. Row 3: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all fluorescence images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

Internalization of 23.5 nm and 46.3 nm nanospheres into the bacterial cells subjected to EMF exposure was investigated. It was found that approximately 98 ± 4% of the Escherichia coli cells, and 98 ± 4% of the B. catarrhalis cells were able to internalize the 23.5 nm silica nanospheres, as shown in Figure 5.4 (first row). The extent of uptake of the 46.3 nm nanospheres was found to be quite different for the two bacterial strains. While the B. catarrhalis cells could not internalize any of the 46.3 nm nanospheres, 49 ± 4% of the E. coli cells were found to be able to internalize these nanospheres after EMF exposure (second row).

None of the Peltier place heated control or untreated bacterial cells were found to have the capacity to internalize the nanospheres, as shown in Figure 5.5.

It should be noted that up to approximately 5% of the untreated and heat- treated control bacterial cells were observed to be capable of internalizing the nanospheres. This is most likely due to the presence of damaged or dead cells, which are often present in cell populations (Nystrom 2004).

The ability to internalize the nanospheres was found to continue for up to approximately 9 minutes after the initial EMF exposure (data not shown), after which time no further uptake of the nanospheres was detected.

Figure 5.5. Internalisation of 23.5 nm and 46.3 nm nanospheres by the untreated and heat-treated controls. Row 1: CLSM images showing no internalization of either the 23.5 or 46.3 nm nanospheres by the control cells. Row 2: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all fluorescence images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1. 96

A transmission electron microscopy (TEM) analysis of the EMF-exposed bacterial cells confirmed the uptake of the 23.5 nm nanospheres, as shown in Figure 5.6. Within the cross-sectioned cells, it can be seen that some of the nanospheres were located around the cell membrane, whilst the others were found to be present within the cells themselves. The majority of nanospheres, however, were found to be present in the cytosol. In contrast, bacterial cells that had not been subjected to EMF exposure remained intact, with the majority of the cells (95%) showing no capacity to internalize either diameter nanospheres.

Figure 5.6. Internalization of 23.5 nm nanospheres by Gram-negative Branhamella catarrhalis cells following EMF exposure. Scale bars in electron micrographs are 200 nm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

5.3.2. EMF effects on cell morphology

The SEM analysis of EMF exposed bacterial cells did not reveal any major changes in the morphology of the cells resulting from the EMF exposure, however

97 the cells did appear to be somewhat dehydrated, with some traces of leaked cytosolic fluid surrounding the cells being detected, as seen in Figure 5.7. The morphology of the heat-treated bacterial cells was found to be unchanged in comparison to the untreated cells.

Figure 5.7. Morphology of EMF-exposed, heat-treated and untreated Gram-negative Branhamella catarrhalis cells. Scale bars in all scanning electron micrographs are 1 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

5.3.3. EMF effects on cell viability

Cell viability experiments were performed via the direct counting of colony forming units. These experiments were conducted for the EMF-exposed, heat- treated and non-treated bacterial cells. The results showed that 96 ± 7% of the Branhamella catarrhalis and 88 ± 4% of the Escherichia coli cells remained viable after EMF exposure (Figure 5.8). The Peltier plate heated cells maintained their viability (99% ± 6% Branhamella catarrhalis and 99% ± 4% of the Escherichia coli cells). A statistical analysis of the data did not reveal any statistically significant difference between the viability of the Peltier heated and untreated cells (Branhamella catarrhalis (p > 0.05) and Escherichia coli (p > 0.05)) (Figure 5.8). Although the cell viability of the EMF-exposed Escherichia coli cells appeared only slightly affected by the EMF exposure, this difference was found to be statistically significant in comparison to the controls (p < 0.05).

Figure 5.8. Effect of EMF exposure and bulk heat treatment on the viability of Gram-negative bacterial cells. Cells inactivated by boiling (100°C) were found to be non-viable. Data presented are the mean ± standard deviation and representative of 3 independent experiments with 10 replicates each. *p < 0.05 versus the corresponding controls. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

5.4. Discussion

The results reported in this chapter provide evidence that the exposure of Gram-negative bacterial cells to 18 GHz EMF radiation, with a specific energy absorption rate of approximately 5.0 kW kg-1, induced permeability in the cell membranes. The permeability was confirmed by the uptake of propidium iodide into the cells. In addition, TEM and CLSM microscopy confirmed the propidium iodide uptake through the visualization of nanospheres that had been able to permeate through the cell membranes. Further examination of the CLSM and TEM micrographs revealed that after the cells had been exposed to 18 GHz EMF radiation, the cells were capable of internalizing both the propidium iodide and the nanospheres.

The SEM analysis allowed the postulation that the EMF was causing a disruption to the cell membrane, which allowed the leakage of cytosolic fluids from the cells and the internalization of propidium iodide and nanospheres. This effect

99 appeared to be temporary and reversible for a period of up to 10 minutes, after which time no further nanospheres were able to be internalized by the EMF- exposed bacteria. The large sizes of the nanospheres passing through the cell membrane (23.5 nm and 46.3 nm) are important characteristics; it appears that this is the first time the physical internalization of such large exogenous materials by Gram-negative bacterial cells has been observed. The variation in the extent of internalization of the 46.3 nm nanospheres between the two bacterial types studied may be due the differences in the cell membrane components (such as lipids and/or proteins).

Since no internalization of either the propidium iodide or nanospheres was observed for the bacterial cells subjected to conventional heat treatment (Peltier plate heating), it can be assumed that the EMF-induced permeabilization of the cell membrane could not be attributed simply to the bulk temperature rise of the Gram- negative bacterial cells. Therefore, the permeabilization must have taken place as a direct result of either the interaction of the EMF with the Gram-negative bacterial cell membrane and its components (e.g., phospholipids, membrane proteins, etc), or microthermal changes in the cell membrane that were not detectable at the macro level.

Further studies are required to investigate the EMF effects on other cell types, such as Gram-positive bacterial and eukaryotic cells, due to their differences in their structure and cell membrane components. This will allow us to postulate the mechanism of which EMF interacts with bacterial cells.

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Chapter 6. EMF effects on Gram positive bacterial cells

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6.1. Overview

In this chapter, the biological effects resulting from exposing Gram-positive bacterial cells to 18 GHz EMF radiation are reported, including changes to the cell membrane permeability, morphology and viability. The Gram-positive Kocuria rosea CIP 71.15T, Planococcus maritimus KMM 3738, Staphylococcus aureus ATCC 25923, Staphylococcus aureus CIP 65.8T, Staphylococcus epidermidis ATCC 14990T and Streptomyces griseus ATCC 23915 bacterial cells were selected for analysis. The cell suspensions containing these separate bacteria were subjected to three consecutive EMF exposures with SAR doses of approximately 5.0 kW kg- 1. This exposure resulted in a temperature increase between 20 to 40ºC (at a heating rate of 20ºC per min). Samples were then allowed to cool to 20°C on ice (at a rate of 10ºC per min) between exposures. The effects resulting from prolonged multiple EMF exposures using two strains of Staphylococcus aureus bacteria as model organisms were also studied for the first time.

6.2. Background

In Chapter 5, it was proposed that permeabilization of the cell membrane must have taken place as a direct result of the EMF interaction with the Gram- negative cell wall and its components (e.g., phospholipids, membrane proteins, etc). Based on their response to the Gram staining technique, bacteria are classified into two classes, these being Gram positive and Gram negative (Madigan et al. 2003). The cell membrane of Gram positive bacteria is comprised of a different structure and components in comparison to Gram negative bacteria (Madigan et al. 2003).

The outer layer of the Gram positive bacterial membrane is comprised of a thick layer of peptidoglycan embedded with teichoic acid, a unique component that cannot be found in the outer membrane of Gram negative bacteria (Madigan et al. 2003; Royet and Dziarski 2007; Vollmer et al. 2008). The rigidity of the bacterial cell wall is normally considered to be controlled by the thickness of peptidoglycan

102 layer, which is a polymer of β(1-4)-linked N-acetyl glucosamine (GlcNAc) and N- acetyl muramic acid (MurNAc), with all lactyl groups of the MurNAc substituted with stem peptides, typically containing four alternating L- and D-amino acids (Royet and Dziarski 2007). Peptidoglycan accounts for approximately half of the mass of Gram positive bacterial cell walls (Madigan et al. 2003; Meroueh et al. 2006; Royet and Dziarski 2007).

The actual three-dimensional arrangement of the peptidoglycan in the cell wall has been the topic of debate. One model proposes a parallel arrangement of the glycan strands in relation to the cell surface (Vollmer et al. 2008; Vollmer and Höltje 2004), whereas a second model proposes a perpendicular arrangement of the glycan strands relative to the bacterial surface (Dmitriev et al. 2004). The second model, which also postulates the presence of pores in the cell wall with sizes ranging from about 50 to about 500 Å, has been supported by data obtained using atomic force microscopy (AFM) to visualise such pores, as well as through nuclear magnetic resonance (NMR) analysis of the synthetic peptidoglycan fragments (Meroueh et al. 2006; Touhami et al. 2004).

Teichoic acids have been known since 1959 to be a unique component of the Gram positive bacteria outer membrane, which cannot be found in Gram negative bacteria (Knox and Wicken 1973; Madigan et al. 2003; Royet and Dziarski 2007; Vollmer et al. 2008). Although the occurrence can be variable between different genera of Gram positive bacteria, teichoic acids appear to be virtually ubiquitous in the cell membrane (Knox and Wicken 1973). Teichoic acids are polymers, which mostly consist of a linear backbone of poly-glycerolphosphate or poly- ribitolphosphate in which the linkage occurs through phosphodiester groups in positions one and three of adjacent glycerol residues or in positions one and four of adjacent ribitol residues, as demonstrated in Figure 6.1. Structural variation in teichoic acids is determined by the nature and extension of glycosyl substitution in the secondary hydroxyl groups of the glycerol units or of glycosyl substitution in the tertiary hydroxyl groups of the ribitol units. D-Alanine ester residues are usually also found as substituents of either glycerol or glycosyl hydroxyl groups (Knox and Wicken 1973).

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Figure 6.1. The structure of teichoic acids on bacterial cell membrane. (a) Structure of glycerol teichoic acid, and (b) Structure of ribitol teichoic acid.

Most membrane teichoic acids appear to be covalently linked to membrane glycolipids, with the term ‘lipoteichoic acid’ being given to the resulting complexes. Lipoteichoic acids are immunogenic, which specifically depends on the teichoic acid component. The teichoic acid component alone, although reacting with antibodies in certain systems, will not induce antibody formation i.e. hapten (Knox and Wicken 1973). Moreover, the main function of teichoic acids is to provide extra rigidity to the Gram positive bacterial cell through the attraction of cations such as magnesium and sodium (Knox and Wicken 1973).

Since it remained unclear as to whether other bacterial taxa with different cell wall structures and compositions to that of Gram negative E. coli cells (Banik et al. 2003; Barnabas et al. 2010; Campanha et al. 2007; Celandroni et al. 2004; Dreyfuss

104 and Chipley 1980; Kim et al. 2009) (e.g., Gram positive bacteria) would be affected in a similar way by EMF exposures, the aim of this chapter is to investigate whether the 18 GHz EMF exposures would induce permeability in the membranes of Gram positive cocci; Kocuria rosea CIP 71.15T, Planococcus maritimus KMM 3738, Staphylococcus aureus ATCC 25923, Staphylococcus aureus CIP 65.8T, Staphylococcus epidermidis ATCC 14990T and Streptomyces griseus ATCC 23915 bacterial cells. In this work, propidium iodide (Nesin et al. 2011; Pakhomova et al. 2011), large (23.5 nm and 46.3 nm) silica nanosphere uptake assays, Confocal Laser Scanning Microscopy (CLSM), TEM and SEM were employed to assess whether the cells could be made permeable under certain carefully defined experimental conditions.

As there were traces of leaking materials surround EMF exposed Gram negative bacterial cells in the scanning electron micrographs in Figure 5.7 of Chapter 5, possible change of the inner pressure (defined as turgor pressure) of bacterial cells was also investigated. Earlier studies have demonstrated that the turgor pressure may indicate the change of bacterial cell membrane integrity (Cabeen and Jacobs-Wagner 2005; Jiang and Sun 2010). Recently, a theoretical model was developed to describe the bacterial cells and cicada wing surface interactions based on the geometry of the nanopillars. The model suggested that the nanopillars do rupture certain bacteria with lower turgor pressure (Pogodin and Baulin 2010; Pogodin et al. 2011). Our hypothesis was that since the rigidity of cellular membrane had decreased due to the EMF exposures, bacterial cell membranes are more vulnerable to be ruptured by the nanopillars. Hence, the change in metabolic status (i.e. the cellular turgor pressure) of EMF exposed coccoid bacterial Planococcus maritimus KMM 3738, Staphylococcus aureus ATCC 25923, Staphylococcus aureus CIP 65.8T and Staphylococcus epidermidis ATCC 14990T cells were studied using wing membrane of the insect Clanger Cicada Psaltoda claripennis.

Due to the lack of data on the effect the multiple 18 GHz EMF exposures has on bacterial cells, two Staphylococcus aureus strains were subjected to 18 treatments over a one-hour period to monitor the permeability of the cells, together with the number of viable cells that could be grown on the nutrient agar (NA) plates. 105

6.3. Results

6.3.1. EMF effects on cell membrane permeability

CLSM analysis of the bacterial samples exposed to the EMF radiation revealed that the exposure induced permeability in the membranes of the Gram- positive bacterial strains tested. This was confirmed by the uptake of propidium iodide into the cells 1 min (fourth column) following EMF exposure in Figure 6.2. The duration of the permeability effect was determined to be up to 10 min, since no propidium iodide was observed in any tested cell types when exposed to the bacterial cells 10 min after irradiation (second column). Phase contrast micrographs are included in Figure 6.2, highlighting the bacterial cells in the same field of view (first and third columns).

Cells that were inactivated by boiling at 100°C were seen to internalize the propidium iodide (first and second columns), Figure 6.3. The Peltier plate heated control groups (40°C) were not able to internalize the propidium iodide (third and fourth columns). Untreated bacterial cells were not able to internalize the propidium iodide, as shown in Figure 6.4.

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Figure 6.2. Internalization of propidium iodide internalization by the Gram-positive bacterial cells after EMF exposure. Row 1: CLSM images showing the internalization of propidium iodide when placed in contact with the bacterial cells 1 min following EMF exposure. Row 3: no propidium iodide was observed in any tested cell types when exposed to the cells 10 min after irradiation. Row 2 and 4: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

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Figure 6.3. Internalization of propidium iodide by the heat control groups. Row 1: CLSM images showing that no propidium iodide was able to be internalized by the bacterial cells 1 min following heating using a Peltier plate (40°C). Row 3: Propidium iodide internalization was found to have occurred in the heat inactivated cells (boiling, 100°C). Row 2 and 4: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

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Figure 6.4. Internalization of propidium iodide by untreated control samples. Row 1: CLSM images showing that no propidium iodide was able to be taken up by cells that were not exposed to EMF. Row 2: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

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In order to determine an estimate of the pores being formed in the cell membrane upon EMF exposure, treated bacterial cells were exposed to silica nanospheres of different sizes, 23.5 nm and 46.3 nm. The extent of uptake into the cells for each particle size was found to vary according to the EMF exposed bacterial cells, as shown in Figure 6.5 (second and third columns). Phase contrast micrographs, showing bacterial cells in the same field of view, are also included in Figure 6.5 (first column).

Internalization of the nanospheres was found to continue for up to approximately 9 min after the initial EMF exposure, whereas no uptake of the nanospheres was detected when they were placed in contact with the bacterial samples 10 min after the EMF exposure (data not shown).

It should be noted that up to approximately 5% of the untreated and heat- treated control bacterial cells were able to internalize the nanospheres, however this is most likely to have occurred due to the presence of damaged or dead cells, which are often present in cell populations (Nystrom 2004).

In contrast, the control cells remained intact, with the vast majority of the cells (95%) showing no internalisation of either particle size of nanosphere, as shown in Figure 6.6.

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Figure 6.5. Uptake of 23.5 nm and 46.3 nm silica nanospheres by the Gram-positive bacterial cells after EMF exposure. Row 1: CLSM images showing the uptake of 23.5 nm nanospheres (in green) by the EMF exposed cells. Row 2: uptake of the 46.3 nm nanospheres by the EMF exposed cells. Row 3: phase contrast micrographs showing the bacterial cells in the same field of view. Scale bars in all images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

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Figure 6.6. Uptake of 23.5 nm and 46.3 nm silica nanospheres by the untreated and heat-treated controls. Row 1 and 3: CLSM images showing negligible internalization of the 23.5 and 46.3 nm nanospheres by the control (heat treated and untreated) cells. Row 2 and 4: phase contrast micrographs showing bacterial cells in the same field of view. Scale bars in all images are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

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A statistical analysis of the data revealed that there was no statistically significant difference between the internalization of propidium iodide or silica nanospheres in the Peltier heated and untreated cells (K. rosea (p > 0.05), P. maritimus (p > 0.05), Staphylococcus aureus ATCC 25923 (p > 0.05), Staphylococcus aureus CIP 65.8T (p > 0.05), Staphylococcus epidermidis (p > 0.05), and Streptomyces griseus (p > 0.05)). There were, however, statistically significant differences existing between the uptake of propidium iodide and the nanospheres in the EMF exposed cells and the control samples (p < 0.05).

The number of bacterial cells that were able to internalize the nanospheres, together with the loading capacity of each bacterial strain were quantified. The results are summarized in Table 6.1. The corresponding concentration of 23.5 nm nanospheres taken into the cells as a function of relative fluorescence units (RFU) was found to be approximately 0.25 µg mL-1 for K. rosea, 0.31 µg mL-1 for P. maritimus, 0.29 µg mL-1 for S. aureus ATCC 25923, 0.47 µg mL-1 for S. aureus CIP 65.8T, 0.38 µg mL-1 for S. epidermidis and 0.36 µg mL-1 for Streptomyces griseus cells. By dividing the concentration of the nanospheres by the total cell concentration in the cell suspension (108 bacterial cells mL-1), the mass of the internalized nanospheres was found to be approximately 2.5 fg cell-1 (K. rosea), 3.1 fg cell-1 (P. maritimus), 2.9 fg cell-1 (S. aureus ATCC 25923), 4.7 fg cell-1 (S. aureus CIP 65.8T), 3.8 fg cell-1 (S. epidermidis), and 3.6 fg cell-1 (Streptomyces griseus). Divided by the mass of a single 23.5 nm nanosphere, the number of internalized nanospheres per single bacterium was able to be determined as being 139 spheres cell-1 (K. rosea), 172 spheres cell-1 (P. maritimus), 161 spheres cell-1 (S. aureus ATCC 25923), 261 spheres cell-1 (S. aureus CIP 65.8T), 211 spheres cell-1 (S. epidermidis) and 200 spheres cell-1 (Streptomyces griseus). The number of 46.3 nm nanospheres that could be internalized by the bacteria was similarly quantified (Table 6.1).

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Table 6.1. Internalization of silica nanospheres by bacterial and yeast cells after EMF exposure. Silica nanospheres

23.5 nm 46.3 nm

Loading Loading Bacterial strains Nanospheres Nanospheres capacity * uptake cells (%) capacity * uptake cells (%) Kocuria rosea 139 ± 8 99 ± 5 62 ± 8 83 ± 8 CIP 71.15T Planococcus maritimus 172 ± 8 97 ± 5 75 ± 8 80 ± 9 KMM 3738 Staphylococcus aureus 161 ± 8 99 ± 4 81 ± 8 40 ± 7 ATCC 25923 Staphylococcus aureus 261 ± 8 99 ± 3 114 ± 8 44 ± 7 CIP 65.8T Staphylococcus epidermidis 211 ± 8 99 ± 5 Not detected Not applicable ATCC 14990T Streptomyces griseus 200 ± 8 99 ± 5 109 ± 8 55 ± 8 ATCC 23915 * nanospheres per single cell

The silica nanosphere loading capacity was calculated using the fluorescence intensity of the nanospheres. The number of bacterial cells that were able to internalize the nanospheres, expressed as a percentage, was calculated by counting the fluorescent cells present in the CLSM images. The data presented are the mean ± standard deviation and are representative of 3 independent experiments.

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Figure 6.7. Internalization of the 23.5 nm silica nanospheres by Gram-positive bacterial cells following EMF exposure. Row 1: TEM images showing the uptake of 23.5 nm silica nanospheres by the EMF exposed cells. Row 2 and 3: Location of the nanospheres in the control (Peltier plate heated to 40°C and untreated) cells, highlighting that the control samples did not internalize the nanospheres. Scale bars are 200 nm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

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The TEM analysis of the cells confirmed the uptake of 23.5 nm nanospheres by the EMF exposed cells, as shown in Figure 6.7 (first column). Within the cross- sectioned cells, some of the nanospheres were seen to be located around the cell membrane, with others being located within the cells themselves. The majority of nanospheres, however, were found to be located in the cytosol. There was no evidence of any nanospheres being located within the cytosol of the control group cells. Here, the nanospheres were observed to cluster around the cell membrane, as shown in Figure 6.7 (second and third columns).

6.3.2. The effect of EMF exposure on cell morphology

An SEM analysis of the EMF exposed bacterial cells did not reveal any significant changes in the morphology of the cells, as shown in Figure 6.8 (first column), however, as seen with the Gram-negative bacterial cells, the Gram- positive bacterial cells exhibited a somewhat dehydrated appearance. Traces of leaked cytosolic fluid can be seen surrounding the EMF-exposed P. maritimus and Staphylococcus aureus ATCC 25923 bacterial cells. The morphology of the non- treated and heat-treated bacterial cells remained constant, as shown in Figure 6.8 (second and third columns).

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Figure 6.8. The morphology of EMF exposed, heat-treated and untreated Gram-positive bacterial cells. Row 1: SEM images showing the dehydrated appearance of the surface and traces of leaked cytosolic fluid surrounding the EMF exposed cells. Row 2 and 3: the typical morphology of the control (Peltier plate heated to 40°C and untreated) cells. Scale bars are 400 nm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

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6.3.3. EMF effects on cell viability

The viability of the EMF treated cells was assessed. The cell viability results showed that after the EMF exposures with the SAR doses of approximately 5.0 kW kg-1, most of the studied Gram-positive bacterial strains had a cell viability rate of more than 80%, as shown in Figure 6.8, however, a lower viability rate was observed for the Kocuria rosea bacterial cells, this being 62 ± 10%.

The Peltier plate heated control cells maintained their viability above 85% (87 ± 8% Kocuria rosea, 99 ± 6% Planococcus maritimus, 98 ± 7% Staphylococcus aureus ATCC 25923, 99 ± 9% Staphylococcus aureus CIP 65.8T, 99 ± 8% Staphylococcus epidermidis and 99 ± 7% Streptomyces griseus cells). A statistical analysis of the data did not reveal any statistically significant difference between the viability of the Peltier heated and untreated cells (Planococcus maritimus (p > 0.05), Staphylococcus aureus ATCC 25923 (p > 0.05), Staphylococcus aureus CIP 65.8T (p > 0.05), Staphylococcus epidermidis (p > 0.05), and Streptomyces griseus (p > 0.05)), with the exception of the Kocuria rosea bacterial cells (p < 0.05) (Figure 6.8). Although the cell viability of the EMF exposed cells was only slightly affected, this difference was found to be statistically significant in comparison to the controls (p < 0.05).

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Figure 6.9. Effect of EMF exposure and bulk heat on the viability of the Gram-positive bacterial cells. Cells inactivated by boiling (100°C) were found to be non-viable. The data presented are the mean ± standard deviation and representative of 3 independent experiments, each comprising 10 replicates each. *p < 0.05 versus the corresponding controls. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

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6.3.4. Confirmation of change in bacterial metabolic status

The EMF exposed Gram positive bacterial cells collapsed on nanopillar- structured insect wing surfaces as shown in Figure 6.10. This finding confirmed that the EMF disturbs the cell membrane integrity and hence changing cellular inner pressure of Gram-positive bacterial cells.

Figure 6.10. EMF effect on the cell rigidity of four selected Gram positive bacteria. Row 1: typical scanning electron micrographs of untreated Planococcus maritimus KMM 3738, Staphylococcus aureus ATCC 25923, Staphylococcus aureus CIP 65.8T and Staphylococcus epidermidis ATCC 14990T on wing membrane substrata of the insect Clanger cicada Psaltoda claripennis. Row 2: after EMF exposures, Gram positive cells have disturbed cell membrane rigidity and cellular inner pressure, thus are penetrated by nanopillars. Scale bars are 200 nm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg- 1.

6.3.5. Effects of multiple 18 GHz EMF exposures on

Staphylococcus aureus strains

The CLSM analysis of EMF exposed Staphylococcus aureus cells highlighted that after the second EMF exposure, the cells of both strains had become permeable (up to 99% of the treated cells, p < 0.05; Figure 6.10) and remained so for the remainder of the subsequent EMF exposures.

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Figure 6.11. The effect of multiple 18 GHz EMF exposures on the morphology and permeability of Staphylococcus aureus cells. Row 1 and 4: typical scanning electron micrographs of S. aureus ATCC 25923 and S. aureus CIP 65.8T cells after multiple 18 GHz EMF exposure. No significant change of cell morphology was observed up to 7th exposure (insets). Scale bars are 10 μm, inset scale bars are 200 nm. Row 2 and 5: CLSM images showing intake of 23.5 nm nanospheres after the 2nd exposure. Row 3 and 6: the phase contrast images in the bottom row show the bacterial cells in the same field of view. Scale bars are 5 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg- 1.

As the number of EMF exposures was increased up to the seventh and eighth exposure, a gradual decline in bacterial cells that could be recovered on nutrient agar plates was observed, decreasing to 54 ± 5% (p < 0.05) for Staphylococcus aureus ATCC 25923 (8th exposure) and 46 ± 5% (p < 0.05) for Staphylococcus

121 aureus CIP 65.8T cells (7th exposure). After subsequent exposures, however, the cell number was found to increase periodically, up to 78 ± 5% (p < 0.05) for Staphylococcus aureus ATCC 25923 (9th exposure) and 60 ± 5% (p < 0.05) for Staphylococcus aureus CIP 65.8T (8th exposure). Hence, despite the overall decline in the viable cell numbers over continuous EMF exposures, it is noteworthy that after the seventh/eighth, thirteenth/fourteenth, and sixteenth/seventeenth EMF exposures, the number of cells recovered on the plates was found to increase for both strains (p < 0.05, Figure 6.11).

Figure 6.12. The effect of multiple 18 GHz EMF exposures on the viability of Staphylococcus aureus cells. Staphylococcus aureus ATCC 25923 (shaded white) and Staphylococcus aureus CIP 65.8T (dotted grey) cell viability as a function of time and number of EMF exposures. The cell viability of the two Staphylococcus aureus strains is displayed in colony forming units (cfu) per 100 µL. The untreated cells preserved their viability throughout the 54 min period, declining slightly to 97 ± 1% for both strains. The x-axis represents the number of

122 viable cells present (expressed as a percentage) after corresponding EMF exposures and the y-axis represents the number of EMF exposures. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 5.0 kW kg-1.

6.4. Discussion

The results discussed in this chapter indicated that three 18 GHz EMF exposures with 1 min SAR doses of approximately 5.0 kW kg-1 (at 40°C) consistently induced cell wall permeabilisation in all of the studied bacterial cells, regardless of the differences in their cell wall/ membrane structures. The TEM analysis of the ultra-thin (90 nm) cross-sections of the cells confirmed the intracellular location of the 23.5 nm nanospheres. It was observed that the number of nanospheres around the Gram positive bacterial cells were considerably more than that observed around the Gram negative bacterial cells. It could be speculated that the peptidoglycan layer of the Gram positive bacterial cell walls may have played a role in filtering and/or trapping the nanospheres.

Some bacteria such as Gram negative Branhamella catarrhalis (B. catarrhalis) and Gram positive Staphylococcus epidermidis bacteria cells were not able to take up the larger (46.3 nm) nanospheres, yet Staphylococcus aureus cells were able to do so. A comparative analysis of the phospholipid and fatty acid compositions (Table 6.2) indicated that Staphylococcus epidermidis cells have greater proportion of phosphatidyl-glycerol (PG) and less lysyl-PG compared to those levels reported for Staphylococcus aureus (Lechevalier 1977). It is likely that the PG head-groups contributed to the membrane fluidity, affecting the uptake of the 46 nm nanospheres. Some variation in the uptake of the large nanospheres was noted for the Gram negative B. catarrhalis and Escherichia coli (E. coli) bacterial cells. For example, a substantial proportion of PG has been reported to be present in the B. catarrhalis membrane phospholipids; this higher proportion of PG did not allow the internalisation of 46.3 nm nanospheres. The E. coli cell membranes, however, contained a greater proportion of phosphatidyl-ethanolamine (PE) and were able to internalise the 46 nm nanospheres. It is important to note that other components of the cell membrane may also play a role in the permeabilisation of

123 the cell membrane. For example, it appeared that although PG comprises up to 30% of the membrane phospholipids in P. maritimus cells, this bacterium was found to be able to internalise a majority of the 46.3 nm nanospheres.

Amongst the strains tested, it is likely that the greater proportion (up to 65% of total fatty acids) of 15:0 fatty acid, which has the lowest melting temperature (33.5 °C, Table 6.2 and 6.3), might increase the susceptibility of P. maritimus cells to the EMF exposure. Hence, it is suggested that the amount of the phospholipid head-group (PG) and C 15:0 saturated fatty acid present in the cell membrane may play an important role in the destabilisation of lipid bilayer of the membrane by EMF exposure, such that the cells are then able to internalise the 46 nm nanospheres.

The periodic increases in cell number (inferred from the direct counting of viable cells) throughout the multiple 18 GHz EMF exposures is a notable phenomenon, which does not appear to have been previously reported. Copty et al. suggested that the enzyme kinetics could be changed by heating the water molecules that are attached to the cellular proteins (Copty et al. 2006). This perhaps may contribute to the observed increases in cell number.

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Table 6.2. Phospholipids compositions (%) of cell membranes in 18 GHz EMF exposure studies Phospholipids Phosphatidyl Phosphatidyl- Phosphatidyl Diphosphatidyl Phosphatidic Lysyl Phosphatidyl Species -choline ethanolamine -glycerols -glycerol Others acid (PA) -PG -inositol (PI) (PC) (PE) (PG) (DPG) Branhamella catarrhalis (Beebe 0 3 38 41 18 0 0 0 and Wlodkowski 1976) Escherichia coli (Lugtenberg and 0 0 91 3 6 0 0 0 Peters 1976) Kocuria rosea (Whiteside et al. 0 0 0 74 26 0 0 0 1971) Planococcus maritimus (Miller 0 0 8-14 23-38 44-45 0 0 0 1985; Thirkell and Summerfield 1977) Staphyloccoccus aureus (Haest et al. 0 0 0 57 5 38 0 Traces 1972) Staphyloccoccus epidermidis 0 0 0 90 1 0 0 9 (Komaratat and Kates 1975) Streptomyces griseus 0 0 + 0 + 0 + 0 (Lechevalier 1977)

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Table 6.3. Chain melting temperature from rippled lamellar gel to fluid lamellar phase of phospholipid chains in studied cell membranes (Ratledge and Wilkinson 1989a)

Phospholipids Phosphatidyl- Phosphatidyl- Diphosphatidyl Phosphatidic Phosphatidyl- Lysyl- Phosphatidyl- Phosphatidyl- ethanolamine glycerols -glycerol acid (PA) choline (PC) PG inositol (PI) serine (PS) (PE) (PG) (DPG) 15:0 ND 34.7 58.4 33.5 ND ND ND ND

16:0 65 41.4 63.1 to 64.9 41.3 39.5 to 54.3 40.4 40.9 51.4

16:1 ND -36 to -4 -33.5 to 20.7 ND ND ND ND ND

17:0 ND 49.8 70.5 ND ND ND ND ND of fatty acids of fatty

18:0 75.4 55.3 70.4 to 75.8 54.4 ND ND ND 63.7 (°C) m

T 18:1 37 -21 to 41 -16 to 39.7 -21.5 ND ND ND 25

20:0 ND 66.4 81.1 to 83.4 ND ND ND ND ND

ND: no data.

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Chapter 7. EMF effects on the eukaryotic unicellular yeast Saccharomyces cerevisiae

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7.1. Overview

In this chapter, the biological effects resulting from exposing a representative eukaryotic unicellular organism, yeast Saccharomyces cerevisiae ATCC 287 to 18 GHz EMF were thoroughly studied, including changes to the cell membrane permeability, morphology and viability. The cell suspensions were subjected to three consecutive EMF exposures with SAR doses of approximately 5.0 kW kg-1 that resulted in a temperature increase ranging from 20 to 40ºC (at a heating rate of 20ºC per min), followed by allowing the sample to cool to 20°C on ice (at a rate of 10ºC per min) between exposures. After the cell viability test, the maximum temperature was decreased to 33°C (with a heating rate of 13°C per min for 1 min), allowing the sample to cool to 20°C on ice for 1 min (at a rate of 10°C per min) between exposures. The number of EMF exposures was also increased from 3 to 6 cycles to enable the cell permeability assay to be performed.

7.2. Background

In Chapter 5 and Chapter 6, it was reported that the permeabilization of the cell membrane must be a direct result of the EMF interacting with the bacterial cell wall and its components (e.g., phospholipids, membrane proteins, etc). It was further reported that charged phospholipid head groups developed a substantial potential at the lipid ˗ solution interface, influencing the concentration of ions at the interface and hence the permeability properties of the cell membrane (McLaughlin et al. 1970). Hence, it would be of considerable interest to investigate whether exposure to 18 GHz EMF would induce cell permeability in eukaryotic cells possessing different compositions of membrane phospholipids to those of bacterial cells.

The composition of phospholipids of the Saccharomyces cerevisiae yeast cell wall/membrane are mainly phosphatidylcholine (35%), phosphatidylethanolamine (24%), and phosphatidylinositol (35%) (Rank and Robertson 1978; van der Rest et

128 al. 1995). Both the phosphatidylcholine and phosphatidylinositol are present in trace amounts, comprising less than 5% of the total membrane phospholipid composition of, for example, Branhamella catarrhalis (Beebe and Wlodkowski 1976), Staphyloccoccus aureus (Haest et al. 1972), and Streptomyces griseus (Lechevalier 1977) bacteria. Phosphatidylcholine is the major lipid component of most eukaryotic membranes, including Saccharomyces cerevisiae, comprising up to 50% of the total phospholipids present (de Kroon 2007; Kent and Carman 1999). Phosphatidylcholine is the bulk structural element of eukaryotic membranes, which has the ability to spontaneously organize into bilayers due to its overall cylindrical molecular shape (Cullis and De Kruijff 1979). Phosphatidylinositol is mainly concentrated at the cytosolic surface of eukaryotic cell membranes (Di Paolo and De Camilli 2006). It is synthesized primarily in the endoplasmic reticulum and is then delivered to outer membranes either by vesicular transport or via cytosolic phosphatidylinositol-transfer proteins (Di Paolo and De Camilli 2006). It can be reversibly phosphorylated to generate seven species, each being phosphoinositides, which play an important part in controlling membrane–cytosol interfaces (Di Paolo and De Camilli 2006). Their main functions, besides classical signal transduction at the cell surface, include the regulation of membrane traffic, the cytoskeleton, nuclear events and the permeability and transport functions of membranes (Botelho et al. 2004; Di Paolo and De Camilli 2006; Gaidarov and Keen 1999; Odorizzi et al. 2000; Roth 2004). The eukaryotic yeast Saccharomyces cerevisiae has been considered a "model organism" for scientists due to its fast growth rate and for being a unicellular eukaryotic organism (Mortimer 2000).

In light of these findings, the first aim of the work reported in this chapter was to investigate whether the 18 GHz EMF exposures would induce permeability in the membranes of eukaryotic Saccharomyces cerevisiae yeast cells. Propidium iodide, large (23.5 nm and 46.3 nm) silica nanosphere uptake assays, Confocal Laser Scanning Microscopy (CLSM), Transmission Electron Microscopy (TEM) and Scanning Electron Microscopy (SEM) were employed to assess whether the cells could be made permeable under certain carefully defined experimental conditions. The second aim was to determine the minimum EMF dose that is required to induce permeability in the bacterial cell membranes without compromising their viability. The number of viable cells remaining after exposure 129 was measured using a direct plate counting technique for the bacteria present on the potato dextrose agar (PDA) plates.

7.3. Results and discussion

7.3.1. EMF effects on cell viability

The results obtained from the cell viability tests showed that after the yeast cells had been subjected to EMF exposures with SAR doses of approximately 5.0 kW kg-1 (causing a temperature rise to 40°C), only 47 ± 8% of the yeast cells remained viable, as shown in Figure 7.1. When the SAR doses were reduced to approximately 3.0 kW kg-1 (temperature up to 33°C), even after 6 dosage cycles, most of the yeast cells had retained a cell viability rate of greater than 94%.

The Peltier plate heated control cell samples were found to not be able to sustain a temperature increase to 40°C, with only 57 ± 8% cells remaining viable following the heating cycle. At temperature increases to 33°C, the yeast cells were observed to maintain a viability of 98 ± 7%. A statistical analysis of the data did not reveal a statistically significant difference between the viability of the Peltier heated and untreated cells (p > 0.05), although the viability of EMF exposed cells was slightly affected. This difference was found to be statistically significant to that observed for the control samples (p < 0.05).

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Figure 7.1. Effect of EMF exposure and bulk heat on the Saccharomyces cerevisiae yeast cell viability. Cells inactivated by boiling (100°C) were found to be non-viable. Data are mean values ± standard deviation and are representative of 3 independent experiments, each with 10 replicates. *p < 0.05 versus the corresponding controls. Cell suspensions were subjected to three and six consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1.

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7.3.2. EMF effects on cell membrane permeability

The Saccharomyces cerevisiae yeast cells that exposed to EMF with SAR doses of approximately 3.0 kW kg-1 (temperature up to 33°C) were selected for cell membrane permeability testing due to their high recovery rate (more than 94%). The CLSM analysis demonstrated that exposing the yeast cells to 6 consecutive EMF doses consistently induced permeability in the membrane of the yeast cells, regardless of the differences in cell wall/membrane structures, as confirmed by the uptake of propidium iodide (first column) following EMF exposure in Figure 7.2. This specific EMF bioeffect could not be duplicated using conventional heating methods under similar temperature conditions.

The 3 consecutive exposures of EMF were not able to induce membrane permeability since no propidium iodide was observed to have been taken up into any of the cells being tested. Cells inactivated by boiling (100°C) allowed the propidium iodide to be taken into the cells. The heat control groups and the untreated cells were also not found to take up the propidium iodide. Phase contrast micrographs showed the yeast cells being present in the same field of view (second column).

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Figure 7.2. Internalization of propidium iodide by Saccharomyces cerevisiae yeast cells after EMF exposure. Row 1: CLSM images showing the internalization of propidium iodide following 6 consecutive cycles of EMF exposure. No propidium iodide was observed in any of the yeast cells after 3 consecutive cycles of EMF exposure. The internalization of propidium iodide was observed in the heat inactivated cells (boiling, 100°C). No propidium iodide uptake was observed in the cells subjected to Peltier heat heating to 33°C. CLSM images showing no propidium iodide uptake by the untreated cells. Row 2: phase contrast micrographs showing yeast cells in the same field of view. Scale bars are 5 μm. Cell suspensions were subjected to three and six consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1.

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Permeability assays performed using the two diameter nanospheres revealed that only the 23.5 nm nanospheres were able to be internalized up by the EMF- exposed yeast cells, as shown in Figure 7.3 (first and second row). None of the control groups (untreated and heat-treated cells) had the capacity to internalize either diameter nanosphere samples. Phase contrast micrographs show the yeast cells in the same field of view (third row). The lack of permeabilization of young yeast cells (Figure 7.3, indicated by arrows) was observed, however due to the lack of data pertaining to the composition of the young yeast cell wall, reasons for this phenomenon being observed remain unclear.

It should be noted that up to approximately 5% of the untreated and control heat-treated yeast cells were observed to be capable of internalizing the nanospheres, most likely due to the presence of damaged or dead cells.

A statistical analysis of the data revealed that there was no statistically significant difference between the Peltier heated control and the untreated cells (p > 0.05). There were, however, statistically significant differences observed between the EMF-exposed cells and the control samples (p < 0.05).

It was found that 97 ± 5% of the yeast cells had the ability to internalize the 23.5 nm nanospheres, with a loading capacity of 27,778 ± 8 nanosphere per single cell.

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Figure 7.3. Nanospheres (23.5 nm and 46.3 nm) internalization by the Saccharomyces cerevisiae yeast cells after 6 cycles of EMF exposure. Row 1: CLSM images showing 23.5 nm nanospheres (in green) being taken up by the EMF-exposed cells. Row 2: Internalization of the 46.3 nm nanospheres was not observed in the EMF-exposed and control cells. Row 3: Phase contrast micrographs showing the yeast cells in the same field of view. Arrows indicate young yeast cells, which were unable to take up any nanospheres. Scale bars are 5 μm. Cell suspensions were subjected to three and six consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1.

A TEM analysis of ultra-thin (80 nm) cross-sections of the cells confirmed the intracellular location of the 23.5 nm nanospheres. Although the nanospheres were observed entering the cells themselves, they were also found to be present on the surface of the cell walls on the EMF exposed cells, as shown in Figure 7.4. It could be therefore postulated that the mannoprotein/β-glucan layer of the yeast cell wall (Lipke and Ovalle 1998) may have played a role in filtering and/or trapping

135 the nanospheres. In contrast, the non-EMF exposed cells remained intact, with most the cells (95%) showing no internalization of either size of nanosphere.

Figure 7.4. Internalization of 23.5 nm nanospheres by the Saccharomyces cerevisiae yeast cells following 6 cycles of EMF exposure. Row 1: TEM images showing the internalization of 23.5 nm nanospheres by the EMF-exposed cells. Row 2: Location of the 23.5 nm nanospheres outside and around the cell membrane of both the untreated and heat-treated cells, showing a uniform cytosol with no nanospheres being present. Scale bars are 200 nm. Cell suspensions were subjected to three and six consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1.

7.3.3. EMF effects on cell morphology

A SEM analysis of the EMF-exposed yeast cells did not reveal any significant change in their morphology, as shown by the images presented in Figure 7.5.

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Figure 7.5. The morphology of EMF-exposed, heat-treated and untreated Saccharomyces cerevisiae yeast cells. Column 1: SEM image highlighting that no significant change in cell morphology occurred after exposure to the EMF. Column 2 and 3: Typical cell morphology of the control (heat treated to a temperature of 33°C) and untreated cells. Scale bars are 1 μm. Cell suspensions were subjected to three and six consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1.

7.4. Further direction

A further study of the effect that exposure of multicellular eukaryotes and liposomes (with or without proteins) to an EMF is required in order to fully evaluate the biological effects resulting from such multiple EMF exposures. This information will allow EMF exposure to be assessed with regard to its applicability as a new drug delivery/release technique. If successful, this technique would have distinct advantages over conventional systems in that it is non-invasive, can be applied locally, and is cost-effective.

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Chapter 8. The effects of EMF exposure on red blood cells

138

8.1. Overview

In this chapter, the biological effects associated with red blood cells being exposed to 18 GHz EMF radiation were determined. The red blood cells were obtained from a representative eukaryotic multicellular organism, the New Zealand rabbit. The resulting effects on the cells including changes in cell morphology and membrane permeability, were determined. Cell viability testing was performed by subjecting cell suspensions to three consecutive 4.1 kW kg-1 EMF exposures that resulted in temperature increases ranging from 20 to 37ºC (at a heating rate of 17ºC per min). The samples were then allowed to cool to 20°C on ice (at a rate of 10ºC per min) between exposures. Following the cell viability testing, the maximum heating temperature was decreased to 33°C (with a heating rate of 13°C per min for 1 min), with the samples then allowed to cool to 20°C on ice for 1 min (at a rate of 10°C per min) between exposures. The number of EMF exposures was increased from 3 to 6 cycles to study whether cell permeability was achieved after these exposures.

8.2. Background

Gaining an understanding of whether EMF-induced permeability of RBCs can be achieved is of particular interest in the context of RBC preservation and freeze-drying. This understanding is also beneficial, from a fundamental point of view, when inducing pinocytosis and endocytosis in RBCs in the development of RBC-targeted drug delivery platforms (Stoll and Wolkers 2011). This interest arises because RBCs are biocompatible, biodegradable, non-immunogenic in nature, and less prone to aggregation and fusion (Magnani 2003; Tan et al. 2015). In order to target the reticuloendothelial system21 or to reduce the extent of allergenic reactions, exogenous materials and/or drugs can be protected from the extracellular environment by encapsulation within RBCs (Bourgeaux et al. 2016; Tan et al. 2015). The plasma membrane of RBCs can protect encapsulated drugs from inactivation, resulting in a prolonged and controllable lifespan (Magnani 2003; Tan

139 et al. 2015). Current approaches for the production of RBCs that contain encapsulated drugs exploit the techniques of osmotic diffusion and electroporation; these processes have a minimal effect on the structure and morphology of the RBCs (Bourgeaux et al. 2016).

The aim of this chapter was to determine whether exposing RBCs to 18 GHz EMF could induce cell permeability. If achieved, this technique would offer an additional and potentially important means of drug delivery.

8.3. Results and discussion

8.3.1. EMF effects on cell morphology

The morphologies of RBCs that were subjected to EMF exposures that resulted in final temperatures of 33 and 37°C was examined in order to confirm that the exposure to EMF does not cause the physiological death of the RBCs. Analysis of the SEM images of the RBCs after exposure showed that the EMF-treated RBCs preserved their original morphology, as shown in Figure 8.1, with no statistically significant difference in comparison to the controls (p > 0.05). Only 9 ± 1% and 14 ± 1% (at 33 and 37°C, respectively) of the EMF-exposed RBCs turned into vesicles and acanthocytes, respectively (top row). According to previous reports, spherisation, partial fragmentation and vesiculization of RBCs could be the result of damaged lipid membrane and/or skeletal proteins such as spectrin (Ivanov et al. 2007; Parshina et al. 2013; Vodyanoy 2015). In previous work, the morphological changes in heat-treated human RBCs were recorded at 44°C, (Zarkowsky 1979) while 49°C was regarded as the critical temperature for RBC fragmentation (Zarkowsky 1979). As such, EMF exposures resulting in final cell temperatures within the range of 33 to 37°C, should not affect the morphology of the RBCs. Indeed, it can be seen, from the images presented in Figure 8.1, that the morphology of the non-treated and control Peltier heat-treated RBCs remained unchanged as a result of EMF exposure (second and third rows). This is a manifestation of the

140 change in the area to volume ratio, which in turn, reflects the microscopic changes in membrane elasticity and/or the local osmotic pressure. It was also found that approximately 1.5% of the total water present was evaporated from the samples during the EMF exposures, which was considered to be negligible and should not affect the overall osmotic pressure.

Figure 8.1. RBC morphology after 18 GHz EMF exposure. Typical SEM micrographs of rabbit RBCs after 18 GHz EMF radiation exposure, resulting in final temperatures of 33 and 37°C. Approximately 9% (33°C) and 14% (37°C) RBC vesicles (first row) and acanthocytes (first row) were observed. The morphology of the non-treated and control Peltier heat-treated RBCs remained unchanged in their morphology (second and third rows). Scale bars are 10 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 4.1 kW kg-1 that resulted in temperature increases ranging from 20 to 37ºC (at a heating rate of 17ºC per min). Then the SAR was reduced to approximately 3.0 kW kg-1 which resulted in temperature increases ranging from 20 to 33ºC (at a heating rate of 13ºC per min).

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8.3.2. EMF effects on cell membrane permeability

A confocal laser scanning microscopy (CLSM) and transmission electron microscopy (TEM) analysis of the EMF-exposed RBCs showed that the EMF exposure consistently induced permeability in the RBC membrane, as confirmed by the uptake of silica nanospheres of two different sizes, as shown in Figure 8.2. There were statistically significant differences between the EMF exposed RBCs and the control samples (p < 0.05).

Figure 8.2. Permeabilization of RBCs resulting from exposure to an 18 GHz EMF. CLSM images show an uptake of 23.5 and 46.3 nm nanospheres (first row). A lipophilic membrane stain, DiI (Life Technologies, Scoresby, VIC, Australia) was used to stain the entire population of RBCs for contrasting purposes (first row). The phase contrast images (second row) show erythrocytes in the same field. Scale bars are 2 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1 that resulted in temperature increases ranging from 20 to 33ºC (at a heating rate of 13ºC per min).

The CLSM analysis also indicated that internalization of the nanospheres could continue for up to approximately 9 min after the EMF irradiation (data not

142 shown); whereas no uptake of the nanospheres was detected when the RBCs were exposed to the nanospheres 10 min after the EMF irradiation. This result indicated that the cells remained permeable for 9 min, in agreement with previously reported observations for other cell types (Nguyen et al. 2016; Nguyen et al. 2015; Shamis et al. 2011). This may be due to protracted relaxation processes in lipid membranes after mechanical disturbance.

The silica nanospheres were chosen for this study because they are hydrophilic and thus do not spontaneously cross through the lipid bilayer (Pogodin et al. 2012); the sizes of 23.5 nm and 46.3 nm where chosen to be smaller than the spectrin mesh size (Liu et al. 2003; Parshina et al. 2013). It has also been reported that the neutrally charged surface of the nanoparticles prevents any nonspecific interactions taking place within the membrane (Verma and Stellacci 2010). Thus, this type of nanosphere could be used as the negative control group. The results obtained for both the Peltier heated and untreated RBCs confirmed that in both cases, no nanospheres could be taken into the cell membrane in Figure 8.3, which rules out bulk temperature change as the cause. Further, there was no statistically significant difference between the control samples (p > 0.05). However, the effect of instantaneous localized temperature elevation (Ti) (Shamis et al. 2012a) elevation cannot be ruled out as a potential cause. This is because Ti can be greater than the bulk temperature (TB) in order to satisfy the Arrhenius equation (� =

� − � �ⅇ ��, where EA is the activation energy, R is the gas constant, and T is the temperature) (Shamis et al. 2012a), which means that it is possible that Ti differed between the EMF and Peltier plate conditions. This possibility cannot be determined from the present study because Ti is a function of EMF energy input and is not directly measurable due to its short existence and molecular nature (Shamis et al. 2012a).

It was further found here that the loading efficiency of the 23.5 nm nanospheres was over 96% for both 33 and 37°C temperature conditions with no statistically significant difference (p > 0.05). The loading efficiency of the 46.3 nm nanospheres was, however, 46 and 58% for samples subjected to 33 and 37°C temperatures, respectively, as demonstrated by the data presented in Table 8.1.

143

Figure 8.3. Permeabilization of RBCs resulting from exposure to an 18 GHz EMF. No uptake of 23.5 and 46.3 nm nanospheres by the control groups. CLSM and phase contrast images showing the appearance of the RBCs remained unchanged, and with no internalization of nanospheres. Scale bars are 2 μm (first and second rows). Typical TEM images of ultra-thin (70 nm) cross-sections of RBCs, showing the cell membrane of untreated and heat-treated RBCs with a uniform cytosol without any 46.3 nm nanospheres being present. Scale bars are 1 μm (fourth row). Inset scale bars are 200 nm (third row). Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1 that resulted in temperature increases ranging from 20 to 33ºC (at a heating rate of 13ºC per min).

144

Table 8.1. Internalization of silica nanospheres by RBCs subjected to EMF irradiation. Silica nanospheres

23.5 nm 46.3 nm

Loading Loading Loading Loading Temperature capacity efficiency capacity efficiency (°C) (fg)* (%) (fg)* (%)

33 16 ± 1 96 ± 5 14 ± 1 46 ± 6

37 22 ± 1 98 ± 5 24 ± 1 58 ± 6

* per single RBC The nanosphere loading capacity was calculated using the fluorescence intensity of the nanospheres. The number of RBCs that were able to internalize the nanospheres, expressed as a percentage, was calculated by counting the total number of fluorescent cells in the CLSM images. Data are mean ± standard deviation, and are representative of 3 independent experiments.

145

Overall, the number of RBCs that were able to internalize the 46.3 nm nanospheres increased (up to 12% of total RBCs) when the temperature was increased from 33 to 37 °C with a statistically significant difference (p < 0.05) (Table 8.1), which may indicate that changes in the cellular membrane and spectrin network occurred as a result of the heat exposure. Moreover, in the samples that were subjected to the 33°C and 37°C temperatures achieved as a result of the EMF exposures, a single RBC was estimated to have the capacity to internalize an additional 6 and 10 fg of the 23.5 and 46.3 nm nanospheres, respectively, with a statistically significant difference (p < 0.05), as shown in Table 8.1.

It should be noted that only the RBCs with circular morphology in the phase contrast images were used for the quantification of the loading efficiency, as shown in Figure 8.4.

Figure 8.4. Quantification of the permeabilized RBCs resulting from 18 GHz EMF exposure. CLSM images show an uptake of 23.5 and 46.3 nm nanospheres (first row). The phase contrast images (second row) show RBCs in the same field. Only the RBCs with circular morphology in the phase contrast images were counted for the quantification. Scale bars are 10 μm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 4.1 kW kg-1 that resulted in temperature increases ranging from 20 to 37ºC (at a heating rate of 17ºC per min). Then the SAR was reduced to approximately 3.0 kW kg-1 which resulted in temperature increases ranging from 20 to 33ºC (at a heating rate of 13ºC per min).

146

The results obtained in this work, regarding the temperature-dependent internalization of RBCs, were in agreement with the observations reported by Harisa et al. (2011). These authors reported that a greater degree of loading of the drug pravastatin into RBCs could be achieved at 37°C compared to that obtained at 25°C.

A TEM analysis of ultra-thin (70 nm) cross-sections of the RBCs revealed the intracellular location and the stages of nanosphere translocation. It can be seen from the TEM micrographs that the internalized nanospheres, while being in close proximity to the membrane, appeared to be engulfed by the cell membrane itself, and then translocated into the cytosol, as seen in Figure 8.5 and indicated by arrows. There were also noticeable clusters of nanospheres trapped within the membrane.

Figure 8.5. Internalization of 46.3 nm nanospheres into the EMF- exposed RBCs. Typical TEM images of ultra-thin (70 nm) cross-sections of EMF- exposed RBCs, showing the internalization of 46.3 nm nanospheres. The RBCs exposed to EMF and allowed to reach a temperature of 37°C were able to 147 internalize a greater number of nanospheres than those achieving a maximum temperature of 33°C. Scale bars are 0.5 μm (second row). Inset scale bars are 200 nm (first row). Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 4.1 kW kg-1 that resulted in temperature increases ranging from 20 to 37ºC (at a heating rate of 17ºC per min). Then the SAR was reduced to approximately 3.0 kW kg-1 which resulted in temperature increases ranging from 20 to 33ºC (at a heating rate of 13ºC per min).

It was also found, however, that the 23.5 nm-nanospheres were able to cross through the 2D spectrin network into the cytosol, as seen by the images presented in Figure 8.6, indicated by arrows. The untreated and control Peltier plate heated RBCs were found not to possess any internalized nanospheres, as highlighted by the images presented in Figure 8.3, highlighting the lack of cell permeability in these samples.

Figure 8.6. Internalization of 23.5 nm nanospheres into the EMF- exposed RBCs. Typical TEM images of ultra-thin (70 nm) cross-sections of EMF- exposed RBCs, showing the internalization of 23.5 nm nanospheres. It appeared that the 23.5 nm nanospheres were able to cross over the 2D spectrin network into the cytosol (indicated by arrows). Scale bars are 200 nm. Cell suspensions were subjected to three consecutive electromagnetic field (EMF) exposures with specific absorption rate (SAR) doses of approximately 3.0 kW kg-1 that resulted in temperature increases ranging from 20 to 33ºC (at a heating rate of 13ºC per min).

It should be noted that the translocated large nanospheres often remained in close proximity to the membrane on the inside of the RBCs, as seen in Figure 8.5. The 46.3 nm-nanospheres were most likely trapped between the membrane and the cytoskeletal network, which is located below the membrane of the RBCs. This could be due to the unique organisation of the cell membrane (Ohvo-Rekila et al. 2002). The asymmetrical phospholipid-cholesterol bilayer, composed of phosphatidylserine, is enriched in the inner leaflet (Muzykantov 2010). Underneath

148 the phospholipid-cholesterol bilayer is a two-dimensional (2D) cytoskeleton that is approximately 7.9 nm in thickness (Liu et al. 2003). This cytoskeleton network is anchored to the phospholipid bilayer, which has the dimensions of an approximately 162 nm × 65 nm (length × width) mesh (Liu et al. 2003; Parshina et al. 2013). The 2D cytoskeleton consists of spectrin heterodimers and actin bundles and is involved in the process of cell volume regulation (Parshina et al. 2013). This skeletal network allows RBCs to undergo significant extensional deformation whilst maintaining their structural integrity (Hansen et al. 1997). The results obtained in this work suggested that while the 46.3 nm nanospheres were able to be translocated through the phospholipid-cholesterol bilayer, only the 23.5 nm nanospheres had the ability to pass through the 2D cytoskeleton network.

It has been reported that RBCs may undergo invagination of their cell membrane, which can then be ‘pinched off’ and sealed, forming intracellular vacuoles (Ginn et al. 1969). This internalization of the cell membrane is reminiscent of the pinocytosis that has been observed in other cell types (Ginn et al. 1969). Ginn at el. suggested that this invagination process would result in a decrease in surface area of cell membranes under tension, thus reducing the critical haemolytic volume of the cell (Ginn et al. 1969). The consequences of membrane internalization are, therefore, similar to those produced by fragmentation, in that both processes will result in a decrease in the overall area of the cell surface membrane (Ginn et al. 1969).

The results of this study have demonstrated, for the first time, that exposing RBCs to 18 GHz EMF has the capacity to induce cell permeability without compromising cell viability. This effect was not found in the Peltier heating control condition, which shows that it was not due to bulk temperature rise. However, whether more spatially and temporally localised temperature changes may account for the effect cannot be determined from the present study. Upon EMF exposure, the RBC membranes are thought to become permeable due to mechanical membrane disturbance. It is proposed here that the EMF-induced nanoparticle uptake is a unique and universal phenomenon, because diverse cell types could be equally affected in a similar way. The elucidation of the molecular mechanism/s of the EMF-induced permeability will require further investigation.

149

Chapter 9. General discussion

150

9.1. Overview

In this chapter, a number of general discussions are drawn to explain the way in which 18 GHz EMF exposure interacts with cells, resulting in a number of bioeffects, particularly cell permeability, in various typical representatives of prokaryotic and eukaryotic taxa. These included two Gram-negative bacteria (Branhamella catarrhalis ATCC 23246 and Escherichia coli ATCC 15034), six Gram-positive bacteria (Kocuria rosea CIP 71.15T, Planococcus maritimus KMM 3738, Staphylococcus aureus CIP 65.8T, Staphylococcus aureus ATCC 25923, Staphylococcus epidermidis ATCC 14990T, and Streptomyces griseus ATCC 23915), a eukaryotic unicellular organism (yeast Saccharomyces cerevisiae ATCC 287) and red blood cells (obtained from a New Zealand rabbit). The role of the dosimetry parameters associated with the 18 GHz EMF is discussed in terms of the way in which it induces permeability in the cells.

9.2. Bioeffects: 18 GHz EMF dosimetry requirements

9.2.1. Temperature-dependent approach

The results obtained in this study suggested that the EMF frequency required to induce cell membrane permeability is 18 GHz. Membrane permeability was not found to result from the application of Peltier heating, highlighting that the bioeffect did not arise as a result of the bulk temperature rise alone. It was demonstrated that exposing bacteria to an EMF frequency of 51.8, 53, 70.6, 73 and 90 GHz resulted in changes in the enzymatic activities and degrees of ion (H+ and K+) transport processes through the plasma membrane (Soghomonyan and Trchounian 2013; Torgomyan et al. 2012; Torgomyan et al. 2011; Torgomyan and Trchounian 2012). While oscillations of EMF in the GHz frequency range are too fast to move larger molecules, such as proteins and other biomolecules, they can induce the rotation of water dipoles (English and MacElroy 2003). The rotation and reorientation of the dipoles are responsible for the energy loss in the consequent heating that was

151 induced by the EMF, however, whether more localised temperature changes may account for the cell membrane permeability effect cannot be determined from the present study; this will require further investigation to enable a conclusion to be drawn in this area.

9.2.2. Power variation

In the power variation experiments, bacteria were subjected to 18 GHz EMF exposures at different power levels (8, 9, 10, 15, 16, and 17 W) and for a fixed duration (1 min). It was found that at lower power levels (8, 9, and 10 W), the temperature of the sample altered to inconsistent degrees. In addition, the application of the EMF at these low power levels was not sufficient to generate a uniform EMF, as the samples were unable to absorb a consistent and even amount of EMF, causing the creation of localised “hot spots”. In addition, the energy threshold at which the EMF bio-effects commenced occurring was found to be dependent on the taxonomic affiliation of the sample cells rather than the differences in cell wall/membrane structures. It was established that the threshold level of EMF exposure that was required to induce such cell permeability as to allow the passage of large nanospheres (46 nm) through the cell membrane was between three to six EMF exposure doses, with a specific absorption rate (SAR) of 3 kW/kg and 5 kW/kg per exposure (a power level of 15 W and 17 W, respectively), depending on the type of cell.

9.3. Proposed mechanism/s of the bioeffects arising from exposing cells to an 18 GHz EMF

Throughout this study, it has been demonstrated that the application of an 18 GHz EMF to cell samples has the capacity to induce cell permeability in representative organisms of prokaryotic and eukaryotic taxa, regardless of differences in their cell wall and/or membrane structures. Cell permeability did not occur in the control samples subjected to Peltier plate heating, which highlighted that the cell permeability did not occur as a result of a simple bulk temperature rise.

152

There is a possibility that localised te mperature changes within the sa mples m a y a c c o u nt f or t h e e m er g e n c e of c ell p er m e a bilit y, h o w e v er t his w o ul d n e e d t o b e t h e s u bj e ct of a f urt h er st u d y. It is p ost ul at e d h er e t h at t h e 1 8 G H z E M F a cts dir e ctl y o n w at er m ol e c ul es , i n d u ci n g t h eir r ot ati o n a n d r e ori e nt ati o n , wit h t h e ass o ci at e d p o l ari z ati o n la g b ei n g r es p o nsi bl e f or t h e dissi p ati o n of e n er g y a n d h e ati n g. It w o ul d n ot b e e x p e ct e d t h at t h e l arger molecules i n t h e s a m pl es w o ul d be affected by such r a pi dl y alt er n ati n g fi el ds, si n c e t h eir r ot ati o n al fr e q u e n ci es ar e s uffi ci e ntl y l o w er, w hil e t h e vi br ati o n al fr e q u e n ci es of i n di vi d u al s u bstit u e nt gr o u ps p ossess l o w energy co mpared to t h at of t h e ki n eti c e n er g y generated by the E MF exposure ( Adair 2002) . The s mall water molecules present in the sa mples would receive energy fro m the 18 G Hz E M F e x p os ur es at a hi g h er r at e t h a n c a n b e dissi p at e d, thus leading to an increase in the instantaneous te mperature ( T i) ( S h a mis et al.

2012a) . T h e T i must be much greater than the bulk te mperature ( T B ); T B ( T i> T B ), t o − E a/ R T satisfy the Arrhenius equation (k = Ae ) ( S h a mis et al. 2 0 1 2 a) . T h e T i is a function of the E M F energy input and is not directly measurable due to its short period of existence and molecular nature ( S h a mis et al. 2 0 1 2 a) , h o w e v er, d es pit e any instantaneous te mperature increases taking place in the sa mples during E MF e x p os ur es, t h e c ell vi a bilit y a n d m or p h ol o g y w er e n ot aff e ct e d. As a r es ult, it c a n be concluded that the T i is li k el y t o b e wit hi n t h e p h ysi ol o gi c al t e m p er at ur e r a n g e of t h e c ells.

T h e r ot ati o n of w at er m ol e c ul es disr u pts t h e h y drogen bonding net work , aff e ct i n g t h e w at er pr o p erti es. As a r es ult, t h e di el e ctri c c o nst a nt of w at er, e , is decreased with frequency v a c c or di n g t o E q u ati o n 9. 1 :    )(  s    1   22  ( Equation 9 . 1) w h er e a n d ar e t h e di el e ctri c c o nst a nts of w at er at z er o (st ati c) a n d e s e ¥ i nfi nit e ( o pti c al li mit) fr e q u e n ci es. T h e r el a x ati o n ti m e of t h e w at er m ol e c ul e s, 4 p h a 3 t = , depends on te mperature ( T ), t h e r a di us of t h e m ol e c ul e ( a ) a n d t h e k T vis c osit y ( h ).

It has been r e p ort e d t h at t h e m a xi m u m E M F a bs or pti o n b y w at er at 2 0° C a n d 40° C occurred at 18 G Hz ( Clark and Sutton 1996) a n d 2 5 G H z ( Hasted 1972) ,

1 5 3 r es p e cti v el y. A p art fr o m t h e eff e ct of h e ati n g t h e w at er, t h e d e cr e as e d di el e ctri c c o nst a nt aff e ct s the interaction bet ween charges and Debye (screening) length, which is a measure of the screening of the electrostatic interactions in polarizable a n d s alt y m e di a. I n t h e a bs e n c e of a n E MF, the Debye length c a n b e c al c ul at e d usi n g E q uatio n 9.2: e k T l = s D 4 p e 2 r (E q u ati o n 9. 2 ) w h er e e is the ele mentary charge and r is t h e d e nsit y of t h e c o u nt eri o ns a n d s alt m ol e c ul es.

T h e c h a n g e i n t h e di el e ctri c c o nst a nt aff e cts t h e el e ctr ost ati c i nt er a cti o n b et w e e n c h ar g es i n w at er m e di a a n d t h us, it c a n aff e ct t h e st a bilit y of t h e charged bil a yers of t h e c ell w all, w hi c h ar e co mposed of polar li pi ds. T heref ore, t h e p h ysi c o c h e mi c al st at e ( i.e ., fl ui dit y) of t h e c ell m e m br a n es mi g ht c h a n g e a n d c o ul d render the m more sensitive to me mbrane defor mation ( G u nst o n e et al. 1 9 9 4; L a n d e et al. 1 9 9 5; L e nt a c k er et al. 2 0 1 4) . T his m o d ul ati o n of t h e m e c h a ni c al sti m ul ati o n , in turn , changes the me mbrane tension, causing it to defor m, resulting i n a n enhanced degree of me mbrane traffic king via exocytosis/endocytosis ( Apodaca 2002; Sheetz and Dai 1996) . The me mb rane re mained per meable for approxi mately ni n e mi n ut es aft er E M F e x p os ur e, t h e n r et ur n e d t o its ori gi n al st at e aft er 1 0 mi n ut es, as c o nfir m e d f or t h e diff er e nt c ell t y p es dis c uss e d i n C h a pt e rs 5, 6, 7, a n d 8. T his ti m e m a y b e a f u n cti o n of t h e l o n g relaxation processes in lipid me mbranes after mechanical disturbance.

Endocytosis is an endo me mbrane dyna mic feature of eukaryotes that has not been previously reported for bacteria, with the exception of recently discovered s u b c ell ul ar c o m p art m e nt ali z ati o n i n t w o b a ct eri al t a x a, Plancto mycetes a n d Verruco microbia ( F u erst 2 0 0 5; F u erst a n d W e b b 1 9 9 1; L e e et al. 2 0 0 9; Li n ds a y et al. 2 0 0 1; L o n hi e n n e et al. 2 0 1 0) . F or e x a m pl e, L o n hi e n n e et al. h a v e s h o w n a n endocytosis -like green fluorescence protein ( GFP) bei n g t a k e n u p b y G e m m at a obscuriglobus b a ct eri al c ells ( L o n hi e n n e et al. 2 0 1 0) . In the endocytosis process, localized regions of the plas ma me mbrane warp around the exogenous materials to b e i nt er n ali z e d, f ol d i n a n d d et a c h t o f or m e n d o c yt oti c v esi cl es ( Pana riti et al. 2 0 1 2) .

1 5 4

It has been reported that RBCs may undergo invagination of their cell membrane, which can then be ‘pinched off’ and sealed, forming intracellular vacuoles (Ginn et al. 1969). This internalization of the red blood cell membrane is reminiscent of the pinocytosis that has been observed in other cell types (Ginn et al. 1969). Ginn at el. suggested that this invagination process would result in a decrease in the surface area of cell membranes under tension, thus reducing the critical haemolytic volume of the cell (Ginn et al. 1969). The consequences of membrane internalization are, therefore, similar to those produced by fragmentation, in that both processes will result in a decrease in the overall area of the cell surface membrane (Ginn et al. 1969). The results presented in this work thus raise the possibility that EMF exposure might induce subcellular compartmentalization.

Other studies have suggested that the EMF-induced changes in water properties can alter the conformation of proteins, their degree of hydration, and other properties that results a change in their activity (Shamis et al. 2012b; Soghomonyan and Trchounian 2013; Torgomyan et al. 2012; Torgomyan et al. 2011; Torgomyan and Trchounian 2012).

9.4. The potential applications of 18 GHz EMF exposures in drug delivery and gene therapy applications

The goal of any drug delivery system is to deliver therapeutic amounts of a given drug to a targeted site of the body, followed by its prompt release by the application of appropriate stimuli (Rodriguez-Devora et al. 2012). Over the past decade, a variety of different physical methods have been studied that apply not only in drug delivery but also to gene therapy, such as photo-, sono-, and electro- based techniques (Rodriguez-Devora et al. 2012; Wells 2004; Wells et al. 2010). These physically controlled drug delivery/gene therapy systems offer many advantages, such as them being non-invasive, flexible in design, and tuneable (Kubiak et al. 2011; Paini et al. 2015; Pattni et al. 2015; Rodriguez-Devora et al. 2012; Wells 2004; Wells et al. 2010). In the same way, it is believed that the 18 GHz EMF can be applied as an innovative physical method to induce cell membrane permeability, which can facilitate the absorption of drugs/genes in 155 treating diseases/infections and of cosmetics in beauty therapy applications (Chen et al. 2006; Granot and Rubinsky 2008; Hofmann et al. 1999; Ibey et al. 2011; Rodriguez-Devora et al. 2012; Song et al. 2007; Wells 2004; Wells et al. 2010). Throughout this study, this effect was demonstrated directly through the use of transmission electron microscopy (TEM) and indirectly through the uptake of propidium iodide as well as large (23.5 nm and 46 nm) nanospheres and large macromolecules such as 150 kDa dextran, regardless of the cell types. The membrane permeability effect is fully reversible, with all of the studied cells fully recovering to their original state 9 minutes after exposure to the EMF. The EMF dosages can also be tuned to achieve cell membrane permeability, with minimal damage being caused to cells (attained a high cell viability rate of over 84% and preserved the original morphology). Thus, the application of 18 GHz EMF doses to cells appears to not only be safe, effective, minimally invasive and highly focussed, but also reproducible, titratable and can be used in both in vitro and in vivo studies. Nonetheless, it is essential to study the capability of 18 GHz EMF exposures in bacterial transformation as well as eukaryotic transfection with plasmids or non- permeable molecules, such as drugs, proteins, oligonucleotides, and RNA for gene therapy applications.

In order to apply the application of 18 GHz EMF doses in cell drug delivery applications, two important factors need to be considered; dosimetry and the taxonomic affiliation of the cells/tissues. In dosimetry, although the specific absorption rate (SAR) calculation alone is inadequate to describe the observed non- thermal effects, it can be used in conjunction with a consideration of the characteristics of EMF (such as modulation and frequency) in predicting the resulting EMF-induced biological effects (Panagopoulos et al. 2013). As described by Equation 3.1, SAR is directly proportional to c, the specific heat capacity of the medium. Thus, the specific heat capacity of the medium and/or the tissues needs to be determined relatively before applying the doses of 18 GHz EMF. This can be done using differential scanning calorimetry (Jin and Wunderlich 1990). Based on the taxonomic affiliation of the cells/tissues, the unwanted thermal effects can be minimized by adjusting the final bulk temperature rise, as demonstrated with the bacterial Kocuria rosea cells (Chapter 6), yeast Saccharomyces cerevisiae cells (Chapter 7), and red blood cells (Chapter 8). Although the EMF bio-effects are independent of the differences in cell wall/ membrane structure, these differences 156 may play a role in filtering and/or trapping drugs based on the size, such as the large 46.3 nm nanospheres. It was speculated in this study that the failure of cells to take up the 46.3 nm nanospheres was due to the peptidoglycan layer in the cell walls of bacterial Staphylococcus epidermidis cells (Chapter 6), and the mannoprotein/β- glucan layer in the cells walls of the yeast Saccharomyces cerevisiae (Chapter 7). In the case of red blood cells (Chapter 8), the 46.3 nm nanospheres were most likely trapped between the membrane and the 2D cytoskeletal network underneath. Hence, with an understanding of the cell wall/ membrane structures of targeted cells/tissues, a range of EMF dosages can be carefully estimated and applied to maximize the drug uptake in drug delivery systems without causing significant destruction/inactivation to nearby non-targeted cells/tissues.

157

Chapter 10. Conclusions and Future Directions

158

10.1. Conclusions

The results of this study have demonstrated, for the first time that exposing a range of cells to doses of 18 GHz EMF consistently induced cell permeability. Cells studied in this work included typical representatives of prokaryotic and eukaryotic taxa. These were two Gram-negative bacteria (Branhamella catarrhalis ATCC 23246 and Escherichia coli ATCC 15034), six Gram-positive bacteria (Kocuria rosea CIP 71.15T, Planococcus maritimus KMM 3738, Staphylococcus aureus CIP 65.8T, Staphylococcus aureus ATCC 25923, Staphylococcus epidermidis ATCC 14990T, and Streptomyces griseus ATCC 23915), a eukaryotic unicellular organism (yeast Saccharomyces cerevisiae ATCC 287) and red blood cells obtained from a New Zealand rabbit. It appeared that the 18 GHz EMF exposure induced cell permeability in all cell types studied, dependent on the taxonomic affiliation of the sample cells rather than the differences in cell wall/membrane structures. The dielectric heat generated by EMF might play a significant role in predicting EMF biological effects, particularly when the oscillating EMF electric current energy is rapidly transferred into the translational motion of molecules which results in an increase in the local temperature. According to this premise, the conventional heating might also induce similar biological effects to the EMF exposure at the same temperature raise conditions. In order to test this hypothesis, a Peltier plate heating/cooling system was employed to replicate the bulk temperature profiles experienced by the studied cells during the EMF exposures. The maximum temperatures were kept in the range of 33 – 40 °C, which is the sub-lethal temperature range for the studied cells. The results of this study clearly indicated that cell permeability was not observed for samples subjected to conventional heating with a similar heating profile using a Peltier heating as a control, highlighting that the cell permeability did not occur simply as a result of a bulk temperature rise. Nevertheless, there is a possibility that localised temperature changes brought about by exposure to the EMF may account for the permabilisation, however, this effect cannot be conclusively determined due to the lack of relevant experimental techniques.

In this study, the specific absorption rate (SAR) was determined along with the EMF power intensity not only for quality control and cross comparison of

159 instrument but also to predict the cell membrane permeability effect. It was established that the threshold level of EMF exposure required to induce cell permeability to such a point that large nanospheres (46 nm) were able to pass through the cell membrane was in the range of three and six EMF exposure doses, corresponding to a SAR of 3 kW/kg and 5 kW/kg per exposure (a power level of 15 W and 17 W, respectively), depending on the type of cell. This information is of great importance in drugs/genes delivery systems and beauty therapy applications since the unwanted thermal destruction/inactivation effects were minimized under such treatment conditions. It was demonstrated throughout this study that by careful tuning the EMF dosages, the cell viability of EMF exposed cells remained high (over 84%) while the cells were able to permeabilise the nanospheres. That could be a manifestation of the change of the area to volume ratio, which in turn, reflects the microscopic changes in membrane elasticity and/or local osmotic pressure.

It is likely that the 18 GHz EMF acts directly on water molecules, inducing their rotation and reorientation, with the associated polarization lag being responsible for the dissipation of energy and heating. The rotation of water molecules disrupts the hydrogen bonding network, affecting the water properties and the dielectric constant. This affects the electrostatic interaction between charges in water media and thus, it can affect the stability of the charged bilayers of the cell wall, which are composed of polar lipids. As a result, after EMF exposure, the physicochemical state (i.e., fluidity) of the cell membranes may change and can render them more sensitive to membrane deformation. This modulation of the mechanical stimulation, in turn, changes the membrane tension, causing it to deform, resulting in an enhanced degree of membrane trafficking via a quasi- exocytosis/endocytosis process. We believe that our study highlights the unique and universality of the EMF-induced nanoparticle uptake phenomenon, because diverse cell types could be equally affected in a similar way.

10.2. Future directions

It is anticipated that the results of this work will attract a wide range of interest from many areas, particularly from researchers involved in the development of innovative alternative cell permeability techniques, which may have applicability 160 in many biomedical fields, such as facilitating the absorption of drugs/genes in treating diseases/infections, and in beauty therapy applications.

Further study of the effect of EMF exposure has on radiation resistant brain tumour cancer cells (e.g. CD133-expressing glioma cells) and normal brain cell line (e.g. pheochromocytoma PC12 neuronal cells) is required in order to fully evaluate the unique effects of such multiple EMF exposures. In order to carefully estimate the EMF dosages, an investigation of the cell wall/ membrane structures of targeted cells/tissues is required to maximize the drug uptake in drug delivery systems without causing significant destruction/inactivation to nearby non-targeted cells/tissues. In addition, the specific heat capacity of the medium and/or the tissues needs to be determined relatively before applying the doses of 18 GHz EMF.

Furthermore, the application of an 18 GHz EMF to biological systems to induce membrane permeabilisation could be applied to systems where therapeutic molecules could be loaded into liposomal/biomimetic drug delivery systems, such as liposomes and cell membrane-coated nanoparticle systems. It was suggested that the presence of low melting temperature phospholipid chains (from rippled lamellar gel to fluid lamellar phase) present in the cell membrane may play an important role in the destabilisation of the lipid bilayer of membranes by their exposure to the EMF, such that the cells would then have the capability to internalize 46 nm nanospheres (Chapter 6). This information is of great importance when designing a liposomal/biomimetic drug delivery system using high frequency EMFs as the loading/releasing method, since the major components are phospholipids. The liposomal/biomimetic drug delivery system can be constructed in such a way that the drugs/genes could readily be released without causing significant destruction/inactivation to nearby non-targeted cells/tissues.

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