PLGA) Nanoparticles Across the Nasal Mucosa

In vitro assessment of the transport of Poly D, L Lactic-Co-Glycolic Acid (PLGA) nanoparticles across the nasal mucosa

Albarki, Mohammed Abdulhussein Handooz https://iro.uiowa.edu/discovery/delivery/01IOWA_INST:ResearchRepository/12730608840002771?l#13730783320002771

Albarki, M. A. H. (2016). In vitro assessment of the transport of Poly D, L Lactic-Co-Glycolic Acid (PLGA) nanoparticles across the nasal mucosa [University of Iowa]. https://doi.org/10.17077/etd.cl2e1klm

IN VITRO ASSESSMENT OF THE TRANSPORT OF POLY D, L LACTIC-CO- GLYCOLIC ACID (PLGA) NANOPARTICLES ACROSS THE NASAL MUCOSA

Mohammed Abdulhussein Handooz Albarki

A thesis submitted in partial fulfillment of the requirements for the Master of Science degree in Pharmacy in the Graduate College of The University of Iowa

August 2016

Thesis Supervisor: Professor Maureen D. Donovan Graduate College The University of Iowa Iowa City, Iowa

CERTIFICATE OF APPROVAL

______

MASTER'S THESIS

______

This is to certify that the Master's thesis of

Mohammed Abdulhussein Handooz Albarki

has been approved by the Examining Committee for the thesis requirement for the Master of Science degree in Pharmacy at the August 2016 graduation.

Thesis Committee: ______Maureen D. Donovan, Thesis Supervisor

______Aliasger K. Salem

______Lewis L. Stevens

To my parents for their continuous guidance throughout my life and career

ii ACKNOWLEDGEMENTS

I would like to express my deepest appreciation to my advisor, Professor Dr.

Maureen Donovan. This thesis would not have been possible unless her thoughtful support and guidance. Thank you for your expert advices and for patiently teaching me everything I need to know along the way in my graduate education.

I would like to thank Professor Dr. Aliasger Salem for his generous gift of PLGA polymer used in this study and for allowing me to use instruments in his laboratory.

I would like to express my special thanks for Professor Dr. Lewis Stevens for serving in my thesis committee and for his time in reviewing this work.

I would like to thank Kareem Ebeid from Dr. Salem laboratory for his suggestions in nanoparticle preparation and for performing SEM images of my particles. I would also like to thank my labmates in Dr. Donovan Laboratory, Ana Ferreira, Wisam Albakri,

Namita Sawant, Laxmi Shanthi, Ammar Alkhafaji, Zainab Bakri, and Saikishore Meruva, for their support in this thesis work.

I would also like to thank to Higher Committee for Education Development in

Iraq (HCED) for their financial support.

Finally, I would like to thank my parents, my brother and sisters for their love and support, my work would not have come so far without them.

iii

ABSTRACT

The nasal mucosa provides a rapid, noninvasive route for drug administration to the systemic circulation and even potentially to the CNS. Nanoparticles made from the biodegradable polymer, PLGA, are of great interest for use in drug delivery systems due to PLGA’s relative safety and ease of surface modification. Nanoparticles may provide improved targeting and transport through the nasal mucosa. However, the optimal nanoparticle sizes and surface properties for intranasal delivery are unknown. In this study, we prepared PLGA nanoparticles within a size range of 50-70 nm containing the lipophilic fluorescent dye, Nile Red, using a surfactant-free nanoprecipitation method.

The resulting nanoparticles were evaluated using dynamic light scattering and scanning electron microscopy. Nanoparticle uptake into the nasal mucosa was determined by exposing the tissues to nanoparticle dispersions for 30 or 60 minutes. The in vitro uptake of the nanoparticles by the nasal mucosal tissues revealed that the Nile Red-loaded PLGA nanoparticles were transported across the epithelial layer and accumulated in the sub- mucosal connective tissues. Nanoparticle uptake in the full thickness tissues was time dependent where 2% of the total loads of nanoparticles exposed to the tissues were measured in the mucosal tissue after 30 minutes and 4% were present in the tissues after

60 minutes. The rapid and measurable transfer of PLGA nanoparticles into the nasal mucosal tissues indicate that they may be an efficient delivery vehicle for drugs with either local or systemic activities.

PUBLIC ABSTRACT

Intranasal drug administration provides a rapid, noninvasive route for drug administration directly to the blood and even potentially to the brain. Nanoparticles are particles with diameter < 100 nm that can be used as drug carriers due to their abilities to bypass various biological barriers due to their small sizes. Nanoparticles used in drug delivery are usually made from biocompatible, biodegradable polymers such as poly D, L lactic- co-glycolic acid (PLGA) which can deliver drug without causing long term damage or toxicity. PLGA nanoparticles are of great interest for use as drug carriers due to their relative safety and ease of surface modification.

Nanoparticles may provide improved targeting and transport through the nasal mucosa, however, their optimal size and surface properties for effective intranasal delivery are unknown. In this study, PLGA nanoparticles within a size range of 50-70 nm were prepared using a surfactant-free nanoprecipitation method. The resulting nanoparticles were characterized with dynamic light scattering and scanning electron microscopy for size and shape and by Nano Zeta Sizer for surface charge. Nanoparticle uptake into the nasal mucosa was determined by exposing the tissues to nanoparticle dispersions for

30 or 60 minutes. The in vitro uptake of the nanoparticles by the nasal mucosal tissues revealed that Nile Red-loaded PLGA nanoparticles were transported across the epithelial layer and accumulated in the sub-mucosal connective tissues. The rapid and quantitative transfer of PLGA nanoparticles into the nasal mucosal tissues indicated that these ~50 nm particles may be an efficient delivery vehicle for drugs for either local or potentially systemic activities.

TABLE OF CONTENTS

LIST OF TABLES ...... vii LIST OF FIGURES ...... viii CHAPTER 1: INTRODUCTION ...... 1 1.1 Nanoparticles as Drug Carrier...... 1 1.2 Biodegradable Polymers in Pharmaceutical Drug Delivery System ...... 2 1.3 Intranasal Drug Delivery ...... 4 1.4 Nasal Anatomy and Histology ...... 5 1.5 Nanoparticle Internalization in the Nasal Mucosa ...... 9 1.6 Effect of Nanoparticle Size on the Uptake in the Nasal Mucosa ...... 12 CHAPTER 2: OBJECTIVES ...... 14 CHAPTER 3: MATERIALS AND METHODS ...... 15 3.1 Materials ...... 15 3.2 Fabrication of PLGA Nanoparticles Using Nanoprecipitation Method ...... 16 3.3 Preparation of Nanoparticle Using Surfactant-free Nanoprecipitation Method ...... 19 3.4 Preparation of Nanoparticle Dispersion for Tissue Uptake ...... 23 3.5 Nile Red Fluorescent Dye ...... 28 3.6 Nanoparticle Uptake Studies ...... 28 3.7 Calculation of the Number of PLGA Nanoparticles ...... 32 3.8 Quantification of Nile Red Mass Per Single Nanoparticle ...... 33 3.9 Data Analysis ...... 34 CHAPTER 4: RESULTS AND DISCUSSION ...... 35 4.1 Nanoparticle Preparation Using Nanoprecipitation Method ...... 35 4.2 Nanoparticle Preparation with the Surfactant-Free Nanoprecipitation Method ...... 39 4.3 Lucifer Yellow VS Transport ...... 50 4.4 Quantification of Nanoparticle Uptake ...... 52 4.5 Conclusion ...... 58 APPENDIX A: STANDARD CURVES ...... 60 APPENDIX B: CALCULATION OF THE PLGA POLYMER DENSITY ...... 67 APPENDIX C: CALCULATION OF PLGA NANOPARTICLES NUMBER, YIELD AND LOADING ...... 68 APPENDIX D: PREPARATION OF BUFFER SOLUTIONS ...... 71 APPENDIX E: NANOPARTICLE UPTAKE DATA ...... 74 APPENDIX F: NANOPARTICLE SIZE MEASUREMENT DATA ...... 76 REFERENCES ...... 97

vi LIST OF TABLES

Table 2. General full factorial study design for the evaluation of the effect of preparation parameters on size of nanoparticles prepared using nanoprecipitation method showing levels for three factors:1) polymer amount, 2) temperature and 3) surfactant type with two levels for each factor...... 35

Table 3. Parameters for the preparation of the nanoparticles and the resulting particle size using a surfactant-free nanoprecipitation method...... 39

Table A 1. Fluorescence intensity of Nile Red standard solutions in Cellosolve® Acetate (measurements were performed using 520-620 nm for the excitation and emission wavelengths, respectively)...... 61

Table A 2. Fluorescence intensity Lucifer Yellow VS in glucose 5 % (w/v) solutions (measurements were performed using 430-530 nm for the excitation and emission wavelengths, respectively)...... 64

Table C 1. Nanoparticle percent yield calculation for two groups, each group was prepared by the same technique but collected using different method. The starting amount represents the amount of PLGA polymer dissolved in DMF and the yield amount indicates the mass of PLGA nanoparticles collected after lyophilization...... 70

Table D 1. Chemicals used to prepare one liter PBS...... 71

Table D 2. Concentration of chemicals in one liter KRB (pH adjusted to 7.4)...... 72

Table E 1. Nanoparticle uptake in the olfactory mucosa...... 74

Table E 2. Nanoparticle uptake in the respiratory mucosa...... 75

vii

LIST OF FIGURES

Figure 1. Chemical structure for PLGA polymer; n= number of lactic acid, m= number of glycolic acid monomers (used with modification from PubChem)14...... 3

Figure 2. Anatomy of the nasal cavity, ST: superior turbinate, MT: middle turbinate, IT: inferior turbinate, NV: nasal vestibule (adapted with permission)26...... 6

Figure 3. Bovine nasal respiratory epithelium observed by bright field microscopy of a hematoxylin and eosin stained section (10 μm thickness), A: epithelial layer, B: sub-mucosal layer lamina propria C: goblet cell, D: epithelial cells (provided with courtesy by Ana C. Ferreira)...... 7

Figure 4. Schematic representation for the olfactory epithelium and the olfactory bulb (adapted with permission)17...... 8

Figure 5. Schematic representation of the endocytosis mechanisms with potential activity in the nasal mucosal (adapted with permission)31...... 11

Figure 6. Schematic representation for the preparation of PLGA nanoparticles by nanoprecipitation method (figures produced using ChemDoodle drawing software)...... 16

Figure 7. Chemical structure of Lucifer yellow VS di-lithium salt (molecular weight= 550.4 g/mole, adapted from PubChem)14...... 26

Figure 8. Chemical structure of Nile Red dye (adapted from PubChem)52...... 28

Figure 9. Navicyte® transport system showing the diffusion chambers in a controlled environment of temperature and aeration...... 29

Figure 10. Residual values particle size values showing randomly scattered residuals across the zero line...... 36

Figure 11. The main effect plot for the study factors showing the particle size increasing with increasing polymer load, decreasing with increasing temperature of the aqueous phase and increasing when PVA surfactant was used instead of F-127...... 37

Figure 12. Effect of aqueous phase temperature on the particle size prepared using a surfactant-free nanoprecipitation method (p value from ANOVA with Tukey’s comparison) (n=3 for each temperature level)...... 40

viii

Figure 13. Effect of internal needle diameter on the mean size of nanoparticles prepared using surfactant-free nanoprecipitation method (p value from unpaired t-test with n=3 and the error bars represent standard deviation)...... 41

Figure 14. Example of intensity-weighted nanoparticles size distribution for nanoparticle batch 3B 12 (Table F 1) using a Malvern Zetasizer Nano- ZS (the average size is 62 nm)...... 43

Figure 15. Example of intensity-weighted nanoparticles size distribution for nanoparticle batch 1B 1 (Table F 1) using Nicomp380 ZLS Zetasizer (mean particle size =94.01 nm)...... 44

Figure 16. Zeta potential measurement of nanoparticle batch 8B 18 (Table F 1) showing a value of -26.3 mV (measured with Malvern Zetasizer Nano ZS)...... 45

Figure 17. SEM images of Nile Red-loaded PLGA nanoparticles (Batch 8B 22) prepared by the surfactant-free nanoprecipitation method. A & B are the nanoparticles before dispersion in 5 % (w/v) glucose solution. C & D are nanoparticles after dispersion in 5 % glucose...... 46

Figure 18. Effect of 5 % glucose medium on nanoparticle size (no significant difference was observed by unpaired t-test, n=3 for each group, error bars represent standard deviation)...... 47

Figure 19. Nanoparticle size measurement after two hour incubation with Cellosolve® Acetate. The small peak at ~70 nm indicates some ~ 70 nm nanoparticles remain intact...... 49

Figure 20. Nanoparticle size measurements after 6 hours of incubation with Cellosolve® Acetate indicates the complete dissolution of the nanoparticles with no peak observed near the prepared nanoparticle size range...... 49

Figure 21. Cumulative amount of Lucifer Yellow VS transported across olfactory and respiratory mucosae. The amount is reported as percent of Lucifer Yellow VS in the receiver side compared to the amount of Lucifer Yellow in the donor side. (n=3 for each data point, error bars represent standard deviation)...... 50

Figure 22. Percent of Lucifer Yellow VS transported across olfactory and respiratory mucosae after 90 min incubation time (n=3 for each tissue type, error bars represent standard deviation)...... 51

Figure 23. Number of PLGA nanoparticles (~ 70 nm) recovered from the respiratory mucosa after exposure to a nanoparticle dispersion for 30 min (p value from the unpaired t-test (n=3))...... 53

Figure 24. Number of PLGA nanoparticles (~ 70 nm) recovered from the respiratory mucosa after exposure to a nanoparticle dispersion for 60 minutes (p value from the unpaired t-test (n=3))...... 53

Figure 25. Number of PLGA nanoparticles (~ 70 nm) recovered from the full thickness respiratory mucosa after 30 and 60 min incubation periods (p value from unpaired t-test (n=3))...... 54

Figure 26. Number of PLGA nanoparticles (~ 70 nm) recovered from the olfactory mucosa after exposure to a nanoparticle dispersion for 30 min (n=3). There was no detectable concentration of nanoparticles in the receiver...... 55

Figure 27. Number of PLGA nanoparticles (~ 70 nm) recovered from the olfactory mucosa after exposure to a nanoparticle suspension for 60 minutes (n=3). There was no detectable concentration of nanoparticles in the receiver...... 55

Figure 28. Number of nanoparticles recovered from full thickness olfactory mucosa (p value from unpaired t-test (n=3))...... 56

Figure 29. Comparison of PLGA nanoparticle (~ 70 nm) uptake in olfactory and respiratory nasal mucosal tissues normalized to tissue mass (p value from unpaired t-test (n=3))...... 57

Figure A 1. Minitab output for Tukey’s pairwise comparison Analysis of the means (means that do not share letter are significantly different)...... 62

Figure A 2. Example of standard curve for the Nile Red solutions in Cellosolve® Acetate...... 62

Figure A 3. Minitab output for the Tukey's pairwise comparison of the mean intensity (means that do not share a letter are significantly different)...... 65

Figure A 4. Standard curve for the Lucifer Yellow VS in glucose 5 % (w/v) solution...... 66

x 1

CHAPTER 1

INTRODUCTION

1.1 Nanoparticles as Drug Carrier

Nanoparticles are sub-micron particles which are affording great alternatives in a number of fields especially in engineering, medicine and pharmaceutical drug delivery.

Gregory et al. fabricated the first “nanoparticulate” drug delivery system known as the liposome in 19741. Currently, there are 1400+ nanoparticle-containing products already in the market for different uses2. Theoretically, nanoparticles can be formed in different shapes and sizes from various types of materials, and nanoparticles can be made from a variety of substances such as silicon, iron, gold and biodegradable polymers3-5.

Nanoparticles have physical and chemical properties that differ from the bulk material properties due to the large surface area of the nanoparticles and changes in material properties which occur at nanoscale6,7.

The ability to modify the properties of various types of nanoparticles makes them valuable in improving drug targeting and potentially minimize drug side effects and treatment costs8. Drugs could be loaded inside the nanoparticles which could make them suitable drug carriers in the treatment of many diseases9. Furthermore, the small size of the nanoparticles increases the probability for greater tissue uptake. Nanoparticulate drug delivery systems help in drug targeting; for instance, the FDA-approved pharmaceutical,

Marqibo® (vincristine sulfate liposome injection with particle size ~100 nm), which was developed by Spectrum Pharmaceuticals, improves the pharmacokinetic characteristics of

2 vincristine. This nanotechnology helped to decrease the volume of distribution and to concentrate the drug dose at tumor site when compared to the high volume of distribution and long elimination half-life for the parent drug10.

1.2 Biodegradable Polymers in Pharmaceutical Drug Delivery System

Biodegradable polymers are of great importance in the engineering of drug delivery devices due to their biological compatibility and the ability to control their physicochemical properties and degradation rates. Biodegradable polymers can be classified depending on their source including natural (biologically derived) polymers and synthetic polymers. Many of these polymers are considered relatively safe due to their safe biodegradation which can occur either non-enzymatically, for example as by hydrolysis or via chemical degradation by microorganisms11. Natural polymers such as chitosan experienced less attraction in nanoparticle synthesis as compared to synthetic biodegradable polymers due to the need for long and relatively complicated chemical modifications to prepare a polymer will suited for the preparation of nanoparticles12.

Biodegradable polymers of synthetic origins have been used in the engineering of nanoparticles for many years. Various types of these polymers are available, including polymers with hydrolyzable groups such as polyamides, polyanhydrides and polyesters or polymers with non-hydrolyzable linkages which require an oxidation process prior to their hydrolysis. Aliphatic polyester polymers are the most frequently used polymers in synthesis of the polymeric nanoparticles in drug delivery due to their safety and the ability to control their physicochemical properties via modification to their molecular

3 weight or properties of monomeric species. Examples of these aliphatic polyester polymers include polyglycolide (PGA), D,L polylactide (PLA), and poly D,L lactic-co- glycolic acid (PLGA). The later type is the current polymer of choice in drug delivery due to its high biocompatibility and physical stability11.

PLGA (Figure 1) is aliphatic polyester of lactic acid and glycolic acid monomers connected via ester linkages. Lactic acid has two enantiomers, D and L, and the PLGA polymers usually contains an equal percentage of each enantiomer11. The physicochemical properties of a PLGA polymer depend on its molecular weight and the lactic acid to glycolic acid ratio. PLGA degrades by the hydrolysis of the ester linkages to eventually yield its monomeric forms, which are subsequently converted to carbon dioxide and water in the Krebs cycle13. In this research we used a PLGA (50:50) polymer with inherent viscosity of 0.32-0.44 dl/g (0.1 % chloroform, 25 °∁) which corresponds to a molecular weight range of 24000-38000 Da.

Figure 1. Chemical structure for PLGA polymer; n= number of lactic acid, m= number of glycolic acid monomers (used with modification from PubChem)14.

1.3 Intranasal Drug Delivery

The intranasal route is used for local drug administration for the treatment of nasal congestion and allergies. In addition, it has been recently used as an alternative route for systemic delivery of drugs that have difficulties in administration through other routes such as peptides and proteins. Intranasal delivery provides a rapid, noninvasive route for drug administration due to its highly perfused tissues, a relatively large, permeable epithelial surface, and the potential for rapid absorption resulting in fast onset of action and, in some cases, fewer side effects15,16.

Another proposed pathway for drug absorption after intranasal administration is the direct transport of drugs to the CNS known as “nose-to-brain delivery” via the olfactory and trigeminal nerves17. It was observed early in the last century that viruses like poliomyelitis and stomatitis can enter to the brain via the nose16,18. Later, this pathway was reported in a study by Reiss et al. who showed that stomatitis virus can enter to the brain via the olfactory nerves19. In addition, Chou and Donovan suggested that there is a direct pathway for the transport of hydroxyzine (antihistamine) to the CNS after intranasal administration demonstrated by the higher concentration of the drug in the cerebrospinal fluid (CSF) after intranasal delivery compared to that after intra-arterial administration in rats20. Theoretically many drugs, like Alzheimer’s, Parkinson’s medications could be transported via nose-to-brain pathway and enter to the brain and the spinal cord directly without passing through the blood brain barrier (BBB) and greater transport of drug to the brain might increase treatment efficiency.

Intranasal drug delivery, however, is not without limitations that affect the efficiency and extent of drug uptake. For instance, mucociliary clearance activity can limit the absorption of drugs and nanoparticles and, instead, clears them away from the mucosal surface prior to absorption21. In addition, the volume of the nasal cavity limits the volume of the dose that can be administered to ~ 100-150 μl. While absorption from the nasal cavity does bypass hepatic first pass metabolism, the activity of the metabolizing enzymes found in the nasal mucosa can also decrease the amount of drug available for uptake16.

In order to understand the advantages of the nasal cavity or the upper respiratory tract as an efficient route for drug delivery, a succinct description of the anatomy and histology of the upper respiratory tract is important.

1.4 Nasal Anatomy and Histology

The main functions of the human nose include olfaction and conditioning of the inspired air. The human upper respiratory tract consists of extended vestibules (nostrils) and the internal nasal cavity; these two regions are separated by the nasal valve. The vestibules are covered with keratinized squamous epithelium and contain sweat glands, sebaceous glands and nasal hairs22. This region is of limited value in drug absorption due to the small surface area and the low permeability of the keratinized tissue23. The internal nasal cavity (Figure 2) lies posterior to the nasal valve (NV) region and continues to the nasopharyngeal region; it is divided by the nasal septum into two separate airways. The length of the main nasal passage in an adult is about 5-8 cm with a surface area of

130-180 cm2 on each side of the nasal septum6,24,25. There are three turbinates protruding from the lateral surfaces of each side; the superior (ST), middle (MT) and inferior (IT) turbinates are covered by a columnar epithelium.

Figure 2. Anatomy of the nasal cavity, ST: superior turbinate, MT: middle turbinate, IT: inferior turbinate, NV: nasal vestibule (adapted with permission)26.

The mucosal epithelium is supported by a thin layer (2-10 µm) of collagen fibers called the basement membrane which separates the epithelial layer from the underlying lamina propria25. In the respiratory mucosa, the lamina propria layer is highly vascularized and contains numerous serous and mucous glands6. The major functions of the lamina propria include roles in air humidification and as a defense mechanism from external toxins due to the high vascularization and the presence of the lymphoid tissues; the nasal associated lymphoid tissue (NALT). These lymphoid tissues are spread throughout the sub-mucosal layer and contain antigen presenting T and B cells 27,28.

Brooking et al. suggested that the NALT is an important pathway for nanoparticle uptake, especially for nanoparticles with a positive surface charge29.

The middle and inferior turbinates are covered with a pseudo-stratified columnar epithelium (respiratory epithelium) as shown in Figure 3. This type of epithelium consists of four types of cells: ciliated (columnar), non-ciliated (columnar), goblet cells (mucus secreting cells) and basal cells (progenitor cells)24. The respiratory epithelium covers the majority of the surface of the nasal cavity occupies the posterior two thirds of the nasal cavity. The majority of the epithelium in this region is populated by ciliated epithelial cells (~ 80% of cell population). The ciliated cells are responsible for the mucociliary clearance activity that clears chemicals and particles, including nanoparticles, away from the epithelial surface.

The superior turbinate and the adjacent septum are covered with a specialized pseudo-stratified epithelium (olfactory mucosa) (Figure 4), which consists of olfactory neuronal cells, supporting cells (microvilli containing columnar cells), basal cells and

Bowman’s glands (olfactory glands). The olfactory neurons are bipolar ciliated neuronal cells that act as peripheral receptors. The axons of the olfactory neurons are bundled together in the lamina propria forming small nerves that pass through the cribriform plate and synapse with the neurons in the olfactory bulb. The role of the supporting cells is not fully understood, but they contain ion channels, which are important for olfactory function and for olfactory neurons survival22. Bowman’s glands are large serous glands that extend from the lamina propria to the outer surface and they produce fluids that help in making the olfactory surface ready for the capture of new odorants. The basal cells, which lie on the basal membrane, act as stem cells for olfactory and supporting cells.

Figure 4. Schematic representation for the olfactory epithelium and the olfactory bulb (adapted with permission)17.

1.5 Nanoparticle Internalization in the Nasal Mucosa

Substances are transported across the epithelial layer either by transcellular or paracellular pathways. The paracellular pathway involves the transport of materials through the tight junctions between the epithelial cells and is mainly controlled by the size and hydrophilicity of the substance. Only small, hydrophilic substances may be transported paracellularly29. The transcellular pathways include the transport of material through the cell membrane through diffusion, facilitated and active and also includes endocytotic mechanisms. Nanoparticles were reported by Huang and Donovan to translocate into the nasal mucosa via paracellular and endocytotic (transcellular) pathways30. Generally, nanoparticles with size < 20 nm are sufficiently small to translocate across the mucosal surfaces via paracellular pathways6.

Endocytosis is the main pathway for nanoparticle uptake by epithelial cells. Petros et al. summarized the endocytosis mechanisms (Figure 5) which include two major processes: 1) phagocytosis (cell eating) and 2) pinocytosis (cell drinking) 31. Phagocytosis occurs by forming large (> 250 nm) vesicles called phagosomes and includes the uptake of the solid particles and bacteria32,33. This endocytic pathway is a receptor mediated pathway activated by the interaction of receptors on the cell surface or by the interaction of the ligands adsorbed on the surface of the nanoparticles with those surface receptors.

Phagocytosis occurs only in specific cell types including monocytes, macrophages, and neutrophils and in the nasal cavity likely only occurs in the NALT (nasal associated lymphoid tissues)6.

The other endocytosis pathway is called the pinocytosis and occurs in all cell types using at least four different mechanisms: macropinocytosis, clathrin-dependent endocytosis, clathrin/ caveolae-independent endocytosis and caveolae dependent endocytosis. In these processes, considerable amounts of extracellular fluids are engulfed along with the transferred particles. Macropinocytosis occurs via actin mediated vesicle

(0.5-10 m diameter) formation and involves the uptake of cell debris, bacteria, viruses and particles >1 m. The clathrin (internal protein) mediated pathway (CME) is a receptor-mediated pathway involving the transport of nanoparticles through vesicles

~ 120 nm in diameter. The caveolar (internal protein) mediated pathway is another pathway for nanoparticle uptake which involves the formation of small (60-80 nm) flask- shaped invaginations of the plasma membrane. Moreover, clathrin and caveolar- independent pathways occur by forming endocytic vesicles with diameters of 90-120 nm.

Figure 5. Schematic representation of the endocytosis mechanisms with potential activity in the nasal mucosal (adapted with permission)31.

A study by Chen on the uptake of the carboxylate-modified polystyrene nanoparticles across the nasal mucosa showed that the 20 nm size particles are transported mainly by CME and paracellular pathways in the respiratory mucosa while there were no endocytosis mechanisms involved in the transport of the nanoparticles in the olfactory mucosa. On the other hand, 100 nm nanoparticles are transported using macropinocytosis and caveolar-mediated endocytosis in the respiratory and olfactory mucosae6.

1.6 Effect of Nanoparticle Size on the Uptake in the Nasal Mucosa

Size and surface charge are the main factors that are reported to affect uptake of non-targeted polymeric particles across epithelial surfaces. There are also reports showed that the geometry of the nanoparticles also affects their uptake across mucosal surfaces34,35. However, the production of polymeric nanoparticles with a constant shape other than spherical with similar dimensions to current is still under nanoparticles investigation.

Surface charge could affect nanoparticle uptake in a variety of ways. For example, positively charged particles have been reported to have higher uptake in epithelial cells due to the activation of certain uptake pathways, mainly CME36,37. In addition, nanoparticles with a positive surface charge are believed to interact with the slightly negative charge on the cell surface, and this might increase the interaction of the nanoparticle with the surface receptors. On the other hand, the positive charge on nanoparticles could increase interactions with negatively charged mucin (main component of the mucous layer) which could reduce nanoparticle uptake8,21,38.

The size of nanoparticles was previously reported to influence the uptake of nanoparticles into epithelial cells. A study by Desi et al. on the uptake of PLGA nanoparticles by Caco-2 cells showed that the uptake of 100 nm particles was significantly greater than particles with diameters of 1 μm and 10 μm. Furthermore, uptake of the 100 nm particles was higher even in the presence of the mucus layer in the in situ rat intestinal loop model. It is also observed that particles with 100 nm diameters go deeper in the sub-mucosal layer and accumulate in the Peyer’s patches (aggregation of

13 lymphoid nodules) as compared to particles with diameters of 500 nm and 1 µm which was found to remain in the epithelial cells39,40. Another study by Chen on the uptake of carboxylate-modified polystyrene nanoparticles into the nasal mucosa showed that the uptake of 20 nm polystyrene nanoparticles was greater than 100 nm particles in both respiratory and olfactory mucosae6. The higher uptake of the small size nanoparticles by epithelial cells may be due to the ability of these particles to penetrate the mucus layer effectively and across multiple endocytosis mechanisms active in the epithelial cells.

CHAPTER 2

OBJECTIVES

Nanoparticles made from PLGA, a biodegradable polymer, are suitable for intranasal drug administration, which could result in drug delivery to the systemic circulation and potentially to the CNS. The main objective of this study was to investigate the uptake of PLGA nanoparticles within the size range of 50-70 nm across the nasal respiratory and olfactory mucosae. This objective was investigated through the following specific aims:

1- Optimization of nanoparticle preparation parameters in order to produce PLGA

nanoparticles within the size range of 50-70 nm.

2- Perform in vitro investigations of the uptake of these nanoparticles across the

bovine olfactory and respiratory nasal mucosae.

3- Quantify the number of the nanoparticles that translocate into the epithelial and

sub-mucosal layers of the nasal mucosa using dye extraction method and a

lipophilic fluorescence dye, Nile Red, incorporated into the nanoparticles.

CHAPTER 3

MATERIALS AND METHODS

3.1 Materials

PLGA polymer [Resmor® 503, 50:50 with inherent viscosity 0.32 - 0.44 dl/g

(0.1 % in chloroform, 25 °C) and molecular weight range of (24-38 kDa)] was obtained from Evonik Industries AG (Darmstadt, Germany). Acetone ACS grade, was purchased from Avantor Performance Materials, Inc. (Center Vally, Pennsylvania, USA). Nile Red dye technical grade; Pluronic® (F-127) surfactant; Polyvinyl alcohol (PVA) 80 % hydrolyzed with molecular weight 9000-10000 Da; Trypsin-EDTA solution (ready to use solution) contains porcine trypsin (2.5 g) EDTA (0.2 g) per liter of Hanks’ Balanced Salt

Solution (HBSS Modified, with phenol red, without calcium, without magnesium);

Lucifer Yellow VS Dilithium salt, were obtained from Sigma-Aldrich (St. Louis,

Missouri, USA). N, N dimethylformamide (DMF) ACS grade, was obtained from Fisher

Scientific (New Jersey, USA). SnakSkin® pleated dialysis tubing with 7000 Da molecular weight cutoff was purchased from ThermoFisher Scientific (Rockford, Illinois, USA).

Silastic® silicon membrane, medical grade, with 0.5 mm, was purchased from Dow

Croning Corporation (Midland, Michigan, USA). Amicon® Ultra-15 centrifugal tube with

100 kDa molecular weight cutoff, was obtained from Merck Mellipore Ltd. (Cork,

Ireland). Cellosolve® Acetate (2-Ethoxyethyl acetate) AlfaAesar (Ward Hill, Maryland,

USA). D-Glucose Research Products International Corp. (Mt. Prospect, Illinois, USA).

3.2 Fabrication of PLGA Nanoparticles Using Nanoprecipitation Method

PLGA nanoparticles were prepared using the nanoprecipitation method (Figure 6) originally described by Fessi et al 41,42. Briefly, 50 or 100 mg of PLGA were dissolved in

10 ml acetone in a 20 ml glass test tube and vortexed until dissolved completely. Then,

200 l (equivalent to 15 μg) of Nile Red solution in acetone (75 μg/ml) were added to the polymer solution while vortexing until mixed completely. The resulting solution was then added using a 20 ml syringe with a 23 G needle into an 80 ml beaker containing 50 ml of

0.5 % (w/v) or 5 % (w/v) solution of F-127 and PVA, respectively, under moderate magnetic stirring (~ 300 rpm). The temperature of the aqueous phase was maintained at room temperature or higher up to (50 ±2℃) depending on the desired experimental conditions.

Figure 6. Schematic representation for the preparation of PLGA nanoparticles by nanoprecipitation method (figures produced using ChemDoodle drawing software).

After complete mixing of the organic phase and the aqueous phase, the stirring process was continued for two hours, and, if high temperature was used, the temperature was decreased gradually to return to room temperature. The solvent was removed either under reduced pressure using a Buchi 011 RE 121 Rotavapor (Flawil, Switzerland) with a water bath using a temperature of 45 ℃ or by stirring overnight in a fume hood.

Thereafter, nanoparticles were collected using a two-step centrifugation procedure: 1) suspension was centrifuged at low speed (3000 x g) for 30 minutes at 10℃ using an Eppendorf 5810R centrifuge (Hamburg, Germany) in order to remove large particles. 2) the supernatant that contained small particles in suspension was transferred to a high speed centrifuge to collect the nanoparticles43. Nanoparticles were collected following centerfugation at 21,000 x g using Sorvall Legend XTR centrifuge,

ThermoFisher Scientific (Am Kalkberg, Germany) for 30 minutes at 10℃. In this step, nanoparticles were washed three times with Nanopure® water in order to remove traces of the surfactant, and each time the supernatant was removed, the pellet was resuspended in

30 ml Nanopure® water. After the third washing the pellet was resuspended in 3 – 5 ml

Nanopure® water and filtered with 0.22 µm syringe filter (Merck Millipore Ltd., Cork,

Ireland) into a 15 ml polypropylene centrifugal tube to remove any aggregates.

Nanoparticle size was measured using a NICOMP® 380 ZLS, particle sizing system (Santa Barbara, California, USA) or a Zetasizer Nano-ZS (Malvern Instruments

Limited, Worcestershire, UK). For the nanoparticles prepared using the nanoprecipitation method, no further characterizations other than particle size measurement using dynamic light scattering were performed. In this study, the effect of varying three preparation parameters on particle size was investigated using a general full factorial experimental

18 design: 1) the polymer amount, 2) the temperature of the aqueous phase and 3) the type of surfactant. The experiments were performed using two levels for each factor and each condition was measured in duplicates (Table 1).

Table 1. Experimental conditions included in the preparation of nanoparticles using the nanoprecipitation method. Nanoparticles were prepared by varying three preparation parameters with two levels for each factor: 1) the polymer amount, 2) the temperature of the aqueous phase and 3) the type of surfactant. Standard Polymer Temperature Surfactant Order (mg) (℃) Type 1 50 ~ 25 F-127 (0.5 % w/v) 2 50 ~ 25 PVA (5 % w/v) 3 50 ~ 50 F-127 (0.5 % w/v) 4 50 ~ 50 PVA (5 % w/v) 5 100 ~ 25 F-127 (0.5 % w/v) 6 100 ~ 25 PVA (5 % w/v) 7 100 ~ 50 F-127 (0.5 % w/v) 8 100 ~ 50 PVA (5 % w/v)

3.3 Preparation of Nanoparticle Using Surfactant-free Nanoprecipitation Method

Nanoparticles were prepared using a modified nanoprecipitation method in which the aqueous phase was Nanopure® water only without the addition of an additional surfactant41,42,44. Briefly, PLGA was dissolved in DMF (5:1 w/v) in a 20 ml glass test tube. Nile Red dye solution in acetone (100 µg/ml) was added to the polymer solution while vortexing (equivalent to 10 g of Nile Red for each 5 mg of polymer) until completely mixed. This solution was added to a 40 ml glass beaker containing Nanopure® water using a 20 ml syringe with 20 G or 27 G needle using moderate magnetic stirring

(~300 rpm). Stirring was continued after the addition of the organic phase for an additional two hours. The solution was transferred to a dialysis bag (Snake skin dialysis tube with 7000 Da molecular weight cutoff (ThermoFisher Scientific, IL, USA) and dialyzed against ~ 3 liters of Nanopure® water. The dialysis medium was changed after

~ 4 hours and then every 7-9 hours for a total time of ~ 40 hours.

In order to optimize the preparation conditions, we investigated two preparation parameters for their effects of the nanoparticle size: 1) the effect of the temperature of the aqueous phase, and 2) the internal diameter of the needle. In this study we used two sizes of syringe needles with diameters of 20 G or 27 G.

Three temperature levels were also investigated: 1) room temperature (25 ℃),

2) 40 ℃ and 3) 75 ℃. The later temperature was selected based on the boiling point of the solvent (DMF) which is 153 ℃ and provided the opportunity to increase the temperature even further without concerns regarding a solvent phase change.

3.3.1 Characterization of Nanoparticles

Particle size and zeta potential were analyzed using a NICOMP® 380 ZLS, particle sizing system (Santa Barbara, California, USA) or a Zetasizer Nano-ZS (Malvern

Instruments Limited, Worcestershire, UK). Nanoparticles prepared from optimized preparation conditions were further evaluated using scanning electron microscopy (SEM) with a Hitachi S-4800 (Hitachi High Technologies Corporation, Japan) for size and shape analyses. In order to prepare the samples for SEM, a drop of nanoparticle dispersion was placed on a semi-conductor silicon chip, air dried and coated with 5 nm of a gold and palladium mixture by ion beam evaporation. The coated samples were examined with the

SEM operated at 3 kV accelerating voltage.

3.3.2 Nanoparticle Collection

After removal of DMF using dialysis, the nanoparticle suspension remaining in the dialysis bag was collected using two techniques:

I: The volume of the nanoparticle suspension in water was reduced to 5-7 ml using an Amicon® Ultra-15 (100 kDa molecular weight cutoff) centrifugal tube (Merck

Millipore Ltd., Cork, Ireland) prior to lyophilization in order to decrease the time required for freeze drying. The Amicon® tube enables the concentration of the nanoparticle suspension with minimal mechanical action in order to reduce nanoparticles aggregation. The suspension was concentrated in the Amicon® tube using centrifugation at 650 x g for 10 minutes at 10 ℃ using Eppendorf 5810R centrifuge (Hamburg,

Germany). The resulting suspension (5-7 ml) was filtered through a 0.22 µm Millipore

21 syringe filter (Merck Millipore Ltd., Cork, Ireland) into a 15 ml polypropylene centrifugal tube to remove any large aggregates.

II: In order to minimize the variability in yield and to prevent any aggregation of nanoparticles during centrifugation and lyophilization, nanoparticles were prepared and the resulting suspension was used immediately without any further processing.

3.3.3 Calculation of Nanoparticle Yield

To estimate the percent yield of the prepared nanoparticles, three batches were prepared using identical preparation conditions. After that, nanoparticle suspensions

(three batches for each collection technique described in Section 3.3.2) were freeze-dried in a pre-weighed 15 or 50 ml polypropylene centrifugal tube using a Free Zone 4.5 Liter

Bench-top Freeze Dry System (Labconco, Kansas City, MO, USA) at -52 C and

0.08 mbar. After lyophilization, the test tube containing the remaining nanoparticle powder was weighed and the mass corresponding to the nanoparticles was calculated by subtracting the mass of the empty test tube. The percent yield was calculated using the following equation:

mass of PLGA nanoparticles after lyophylization % yeild = Equation (1) original mass of PLGA used in the preperation

In this study, the mass of PLGA used in the preparation of nanoparticles using the surfactant-free nanoprecipitation method was either10 or 36 mg depending on the preparation conditions.

In order to investigate the efficiency of using the Amicon® Ultra-15 centrifugal tube for collecting the nanoparticles, the percent yield was calculated for three batches of nanoparticles collected with the Amicon tube and three batches of nanoparticles collected with standard 50 ml polypropylene tube. The average percent yield was compared between the two different techniques. In addition, the percent yield was compared among the batches prepared using the same method to compare the reproducibility between triplicates in each collection technique.

3.3.4 Calculation of Nanoparticle Loading

Nile Red loading in the nanoparticles was calculated by measuring the amount of

Nile Red recovered from a sample of prepared nanoparticles and comparing the results to the original amount of Nile Red added to the PLGA solution prior to nanoprecipitation

(in this study the amount of Nile Red was 10 μg of Nile Red per 5 mg of PLGA) according to the following equation:

amount of Nile Red recovered following nanoparticles preparation Equation (2) loading % = amount of Nile Red used in initial PLGA ∗ 100% solution

In order to calculate the amount of the Nile Red remaining in the nanoparticles after preparation, 1 ml of the nanoparticle suspension was partially dried with a gentle stream of nitrogen gas (TurboVap® LV, Caliper Life Science, MI, USA) in a 37 °C water bath to remove the most of the water in order to increase the quantification sensitivity.

Then the particles were incubated for ~8 hours with 1 ml of Cellosolve® Acetate to

23 completely dissolve the PLGA nanoparticles and release the entrapped Nile Red dye.

A particle size measurement was performed to confirm complete dissolution of the nanoparticles. The fluorescent intensity of the Nile Red in the Cellosolve® Acetate solution was measured and the dye concentration was calculated using the appropriate standard curve (Appendix A). The total amount of Nile Red recovered after nanoparticle preparation was calculated by multiplying the amount in 1 ml by the total volume of nanoparticles suspension remaining after dialysis (in this study, the total volume of nanoparticles suspension after dialysis was ~ 30 – 32 ml).

® Cellosolve Acetate (2-ethoxyethyl acetate, C6H12O3) was chosen as a solvent to extract the Nile Red entrapped in the nanoparticles due to its water immiscibility and its ability to dissolve both the Nile Red dye and the PLGA polymer. Cellosolve® Acetate was used previously in our laboratory to dissolve polystyrene nanoparticles, and it was observed to dissolve PLGA nanoparticles. The water immiscibility of Cellosolve®

Acetate allows for rapid separation of the nanoparticle and the dye from the aqueous medium to recover the maximum amount of the dye.

3.4 Preparation of Nanoparticle Dispersion for Tissue Uptake

In order to deliver nanoparticles to the nasal mucosal tissues, nanoparticles need to be suspended in a medium that maintains tissue viability and does not cause any nanoparticle aggregation6. Solutions including buffers (with pH ~ 7) or alternative solutions such as glucose 5 % (w/v) solution were used in this study as a suspension medium to maintain tissue viability during the incubation of nasal mucosal tissues with

24 the nanoparticle suspensions. In this study we used three buffer solutions (pH ~ 7):

1) KRB (Krebs-Ringer Bicarbonate Buffer) with added glucose, 2) PBS (Phosphate

Buffered Saline) and 3) TBS (Tris Buffered Saline). These buffers were prepared as shown in Appendix D. In addition to buffer solutions, glucose 5 % (w/v) solution was also investigated in this study as an alternative incubation medium.

In these studies, nanoparticles prepared with the surfactant-free nanoprecipitation method were lyophilized to provide a nanoparticle powder or nanoparticles suspension was used immediately after preparation without lyophilization to decrease any particle aggregation resulting from freeze drying process. In cases where lyophilization was used, the nanoparticle powder was resuspended in the buffer solution by vortexing. In the case of the nanoparticles suspension without lyophilization, a small amount of glucose equivalent to 5 % (w/v) of the nanoparticle suspension volume was added slowly to the suspension under medium speed vortexing until the glucose powder was dissolved completely.

In order to test the effect of the suspension medium on nanoparticles characteristics, nanoparticle suspension was investigated for any visual aggregation and then it was investigated using dynamic light scattering for any changes in the nanoparticles’ average size range. The effect of the dispersion medium on the viability of the nasal mucosa was investigated as described in Section 3.4.1.

3.4.1 Test of Tissue Integrity in Incubation Medium

The integrity of the nasal mucosal tissues exposed to KRB buffer solution with added glucose as an incubation medium was previously tested in our laboratory and results showed the tissues were viable for at least 3-4 h after harvest16,45.

In order to study the time period over which the excised tissues remained viable when using glucose 5 % (w/v) as the incubation medium, the transport of Lucifer Yellow

VS Dilithium salt was measured during an experiment where the donor and receiver solutions contained 5 % glucose in water and were oxygenated using Carbogen.

Lucifer Yellow VS, (4-Amino-N-[3-(vinylsulfonyl) phenyl] naphthalimide-3,6- disulfonate) Dilithium salt), is a widely used permeability marker to evaluate paracellular permeability through cell monolayers and mucosal tissues 46-50. Lucifer yellow VS

(Figure 7) is a non-permeable, auto-fluorescent dye. It can only be transported between cells, so intact, viable tissues exclude the transport of drug quantities of dye. The detection of greater than 1 % of the donor concentration of Lucifer Yellow in the receiver chamber is an indication of damage to the tissue or of increased paracellular permeability due to the loss of tight junctional adhesion48.

The concentration of Lucifer Yellow was quantified by measuring the fluorescence intensity using a SpectraMax® M5 Multi-Mode Microplate Reader

(Molecular Devices, Sunnyvale CA, UAS) at 430 – 530 nm for excitation and emission wavelengths, respectively. A Lucifer Yellow VS standard curve was prepared by measuring the fluorescence intensity of Lucifer Yellow VS solutions of various concentrations. The fluorescence measurements were performed in triplicate and in three

26 different wells for each concentration (in order to account for instrument and well to well variability) in a 96-well polypropylene plate, and the average intensity of the three measurements was calculated. A linear regression of the fluorescence intensity versus concentration was performed and the detection limit of Lucifer Yellow VS was performed using Minitab 17 software (licensed by the University of Iowa). The detailed calculations and the linear regression equation are explained in detail in Appendix A.

Figure 7. Chemical structure of Lucifer yellow VS di-lithium salt (molecular weight= 550.4 g/mole, adapted from PubChem)14.

Three vertical Navicyte® diffusion chambers (Harvard Apparatus, Holliston, MA,

USA) were prepared for each type of mucosal tissues (respiratory, olfactory). Tissue collection and diffusion chamber setting are described in Section 3.6. The donor side was incubated with a pre-warmed (1 ml) solution containing 55 µg/ml (100 µM) of Lucifer yellow VS in glucose 5 % (w/v) solution and the receiver side was incubated with 1 ml glucose 5 % (w/v). Uptake studies were performed on the same day using two techniques:

I: Measuring the cumulative amount of Lucifer Yellow VS transported at various time points during the 1.5 h incubation.

Aliquots (300 μl) were taken from the receiver side at 15 min time intervals for up to 90 min and replaced by the same volume of glucose 5 % (w/v) solution. The amount of Lucifer Yellow VS in the receiver side was calculated by comparing the fluorescent intensity with the appropriate standard curve. The percent transported was calculated by comparing the receiver side concentration at each sampling time point with the starting Lucifer Yellow VS concentration in the donor side (55 µg/ml in this study).

The cumulative amount was calculated by measuring the dye concentration at each time point and adjusting for the amount withdrawn for fluorescence measurement (amount of

Lucifer Yellow in the 300 µl aliquot used for measurements).

II: Three transport chambers for each tissue type were incubated with a pre- warmed 1 ml solution containing 55 μg/ml (100 μM) Lucifer Yellow VS in glucose

5 % (w/v) solution in the donor side and 5 % glucose in the receiver. After 1.5 h receiver sides were collected and the fluorescence intensities were measured with a SpectraMax®

M5 Multi-Mode Micro Plate Reader. The Lucifer Yellow concentration was calculated using the appropriate standard curve.

3.5 Nile Red Fluorescent Dye

Nile Red (Figure 8), 9-(diethylamino)-5H-benzo[a]phenoxazin-5-one; 9-

(diethylamino)benzo[a]phenoxazin-5-one, is a lipophilic fluorescent dye which is soluble in typical organic solvents. Nile Red gives an excellent fluorescence in many organic solvents, and it is easily quantified with excellent accuracy51.

Figure 8. Chemical structure of Nile Red dye (adapted from PubChem)52.

3.6 Nanoparticle Uptake Studies

3.6.1 Preparation of Bovine Nasal Tissues

Bovine nasal tissues were used for the uptake studies. due to their reported effectiveness to evaluate drug transport across the nasal mucosa6,16,50. Nasal tissues were obtained from Bud’s Custom Meats Co. Riverside, IA, from a freshly killed cow. A flat cut was made to remove the nasal bone and to expose the turbinate bones, which are covered by the respiratory and the olfactory mucosae. These tissues were removed

29 carefully with a sharp knife and transferred immediately to glucose 5% solution and stored over ice. Tissues were transferred to the laboratory within 30-40 minutes.

3.6.2 Quantification of Nanoparticle Uptake

Respiratory and olfactory tissues were separated from the underlying cartilage using tweezers, and 3-4 cm2 sections were mounted in vertical Navicyte® diffusion chambers (Harvard Apparatus, Holliston, MA, USA) with the mucosal surface facing the donor side. The receiver and the donor sides were filled with glucose 5 % (w/v) solution and equilibrated for 10 min at 37 ℃ and aerated with Carbogen gas (95% oxygen and

5 % carbon dioxide) at a rate of 3-4 bubbles per second (Figure 9).

Figure 9. Navicyte® transport system showing the diffusion chambers in a controlled environment of temperature and aeration.

After 10 minutes of equilibration, the solutions in the donor and receiver sides were replaced with a previously warmed, Nile Red-loaded PLGA nanoparticle dispersion

(1 ml) and 1 ml of glucose 5 % (w/v) solution in the donor and receiver cells, respectively. After 30 or 60 minutes (depending on the study design), the receiver and the donor side volumes were collected and kept at room temperature for further analysis. The nasal tissues were removed from the Navicyte® chambers, and the area exposed to the donor chamber was excised, weighed and washed with glucose 5 % (w/v) solution to remove any adsorbed nanoparticles from the surface of the tissue. The tissues were transferred to a 10 ml glass test tube containing 2 ml trypsin-EDTA solution, component was described in Section 3.1 (Sigma-Aldrich, St. Louis, MO, USA) and incubated in

37 ℃ for two hours to remove the epithelial cells. The remaining sub-mucosal layer was transferred to a 10 ml glass tube containing 1 ml of Cellosolve® Acetate in order to dissolve the nanoparticles entrapped in the sub-mucosal layer and extract the Nile Red content.

The trypsin-EDTA + epithelial cells and receiver side solutions were partially dried with a gentle stream of nitrogen gas (TurboVap® LV, Caliper Life Science, MA,

USA) in a 37 °C water bath to remove the most of the water in order to increase the quantification sensitivity. Cellosolve® Acetate (1 ml) was added and mixed thoroughly to dissolve the nanoparticles and to extract the Nile Red dye content. Samples were incubated overnight to extract all of the Nile Red. The amount of the Nile Red released from the dissolved nanoparticles was quantified by measuring the fluorescence intensity using a SpectraMax® M5 Multi-Mode Microplate Reader (Molecular Devices, Sunnyvale

CA, UAS), and the amount of Nile Red was calculated from the appropriate standard

31 curve. The fluorescence intensity for Nile Red was measured at 520 – 620 nm for the excitation and emission wavelengths, respectively.

A Nile Red standard curve was prepared by measuring the fluorescence intensity of Nile Red solutions at various concentrations. The fluorescence measurements were performed in triplicate and in three different wells for each concentration (in order to account for instrument and well to well variability) in a 96-well quartz plate. One concentration was studied at a time, and the average intensity three measurements was calculated. The linear regression of the fluorescence intensity versus concentration and the determination of the detection limit of Nile Red were performed using Minitab 17 software (licensed by the University of Iowa). The detailed calculations and the linear regression for the Nile Red standard curve are provided in Appendix A. The number of nanoparticles corresponding to the amount of extracted dye was calculated based on the determined correlation between particle mass and entrapped dye amount which is described in Section 3.9.

In order to compare the nanoparticle uptake in the nasal mucosa between the respiratory and the olfactory tissues, the number of nanoparticles in the full thickness tissues (epithelial cells and sub-mucosal layer) was calculated and normalized by the mass of the tissue region exposed to the donor solution. The exposed area of the tissues was weighed and the total number of nanoparticles per gram of tissue was calculated.

3.7 Calculation of the Number of PLGA Nanoparticles

The total number of nanoparticles in the suspension after preparation using the surfactant-free nanoprecipitation method was calculated by measuring the mass of individual nanoparticles and determining the total yield of PLGA nanoparticles which resulted after preparation (nanoparticles yield section 3.3.3)53. The mass of single nanoparticle was calculated from the density of PLGA and the volume of sphere according to the following equation:

Mass of individual particle = ρ *(4/3 π푟3) Equation (3)

where ρ is the density of PLGA polymer (50:50 with inherent viscosity of 0.32 - 0.44 dl/g

(0.1 % in chloroform, 25 C) and (r) is the average radius of the nanoparticle measured using dynamic light scattering.

Total number of the nanoparticles was calculated by dividing the total mass of nanoparticles (in this study was 27.47 mg) by the calculated mass of an individual particle calculated in Equation 3.

The density of the PLGA polymer used in this study was calculated by using

Archimedes method by weighing a small amount of polymer in air and determining its volume in water. The density of the PLGA polymer was determined to be (1.16 g/cm3).

The mathematical calculations for the density and the number of particles are shown in

Appendix B and C, respectively.

3.8 Quantification of Nile Red Mass Per Single Nanoparticle

The amount of Nile Red per single nanoparticle was calculated according to the following equation:

total mass of Nile Red (after preperation) Equation (4) Mass of dye per particle = total number of nanoparticles

where the total mass of Nile Red recovered after preparation was calculated in section (3.3.4) and the total number of nanoparticles was calculated in section 3.3.3.

Using the amount of Nile Red per single nanoparticle, the number of nanoparticles translocated into the nasal mucosa could be calculated by comparing the recovered amount of Nile Red from the mucosal tissues with the amount per single nanoparticle.

3.9 Data Analysis

Minitab 17 statistical software, Minitab Inc., USA (licensed by the University of

Iowa) was used to analyze the data for the effect of the preparation parameters on the resulting size of nanoparticles that were prepared using the nanoprecipitation method. A general, full-factorial design was used to find the effect of each factor on the particle size with a significance level of p < 0.05. Furthermore, this design of experiment enabled testing of the effect of the interactions between different factors and their levels when they were combined to prepare the nanoparticles. The experiments were done in duplicate and the average nanoparticle diameter was used to test the effect of each preparation parameters. The remaining data were analyzed using Graph Pad Prism 6 (licensed by the

University of Iowa). Graph Pad Prism 6 was also used to plot the uptake/transport data and to perform basic statistical tests in order to compare different groups of results.

CHAPTER 4

RESULTS AND DISCUSSION

4.1 Nanoparticle Preparation Using Nanoprecipitation Method

4.1.1 Analysis of the General Full Factorial Design

Results for the preparation of nanoparticles using the nanoprecipitation method are summarized in Table 2 including experimental conditions incorporated in the general full factorial design.

Code Standard Run Polymer Temperature Surfactant Type Particle size (nm) name Order (mg) Order (ºC) Replicate Replicate 1 2

1A, A9 1 1 50 ~ 25 F-127 (0.5 % w/v) 107 100

2A, 10A 2 2 50 ~ 25 PVA (5 % w/v) 161 161

3A, 11A 3 3 50 ~ 50 F-127 (0.5 % w/v) 96 98

4A, 12A 4 4 50 ~ 50 PVA (5 % w/v) 140 146

5A, 13A 5 5 100 ~ 25 F-127 (0.5 % w/v) 163 165

6A, 14A 6 6 100 ~ 25 PVA (5 % w/v) 178 173

7A, 15A 7 7 100 ~ 50 F-127 (0.5 % w/v) 137 157

8A, 16A 8 8 100 ~ 50 PVA (5 % w/v) 162 162

Data analysis was performed using the general factorial regression function in

Minitab 17 software and the overall p-value for the model was significant (p < 0.0001) with a coefficient of determination (r2) value of 0.97. Analysis showed that all factors were significant p < 0.005 and the interaction terms were non-significant p > 0.05 except for the interaction between the polymer amount and the F-127 surfactant which was significant p < 0.01. The residuals were normally distributed and randomly scattered across the zero line as shown in Figure10 for the residual versus particle size.

Figure 10. Residual values particle size values showing randomly scattered residuals across the zero line.

In order to predict the effect of the three factors particle size, the factorial plot function was used to analyze the main effects of these terms. The main effect plot

(Figure 11) showed that increasing the amount of polymer used in the preparation of nanoparticles resulted in a nanoparticle with a larger size as compared to using lower amount of polymer during nanoparticle preparation. In addition, the model showed a moderate decrease in the particle size with increasing temperature of the aqueous phase during nanoparticle preparation. Furthermore, the main effect plot predicted a decrease in

37 particle size when F-127 (0.5 % w/v) was used in the fabrication of nanoparticles instead of PVA (5 % w/v).

The observed changes in the size of nanoparticles prepared by the nanoprecipitation method using different amounts of polymer and surfactant types at different temperature of the aqueous phase, suggested that the parameters tested have a significant effect on the particle size within size range tested (96-190 nm). Effect of polymer amount and temperature of the aqueous phase on nanoparticle size could be explained by their effect on the viscosity of the organic phase. Increases in the viscosity

38 decreased the shear forces of water and reduced the rate of diffusion of the organic phase

(polymer solution) into the continuous phase (aqueous phase) and resulted in larger particle sizes. Decreasing the polymer amount or increasing the temperature of the aqueous phase decreased viscosity and resulted in decreasing of particle size. These results were in agreement with previously reported studies about the effect of these parameters on particle size42,44,54,55.

The ability of F-127 surfactant to decrease the particle size is related to the ability of the surfactant to reduce the surface tension of water, which results in a better distribution of the organic phase54. However, no significant difference in particle size was observed by Menon et al. when they used similar concentration (5 % w/v) for each surfactant56. Hence, the observed difference in the particle size between F-127 and PVA in this study may be explained by higher concentration of PVA (5% w/v) compared to

F-127 (0.5 % w/v) which, in turn, resulted in a higher viscosity for the PVA solution and resulting in a larger particle size.

4.2 Nanoparticle Preparation with the Surfactant-Free Nanoprecipitation Method

Nanoparticles were prepared using a surfactant-free nanoprecipitation method according to the conditions listed in Table 3. The size of the nanoparticles was reported as the average size of 3-8 replicates ± standard deviation.

Table 3. Parameters for the preparation of the nanoparticles and the resulting particle size using a surfactant-free nanoprecipitation method.

Code PLGA Amount Aqueous Heat Needle Number of Particle size name (mg ± 0.5) of DMF phase (℃) Size replicates (nm ± SD) (ml) (ml) (n)

1B 10 2 20 25 27 3 88 ± 8

2B 10 2 20 40 27 6 68 ± 4

3B 10 2 40 75 27 3 74 ± 10

7B 36 5 30 40 27 3 62 ± 5

8B 36 5 30 40 20 8 55 ± 4

The use of DMF (N,N dimethyl formamide) as an organic solvent resulted in a significant reduction in particle size to diameters < 100 nm for all batches prepared using this method. The effect of the DMF solvent in reduction of the particle size may be explained by good solvent water miscibility which enhances the distribution of the organic phase into the aqueous phase resulted in particle size reduction. This result was in agreement with the findings observed by Cheng et al. in which the particle size was reduced by increasing solvent water miscibility44.

Increasing the temperature of the aqueous phase resulted in decreases to the particle size. However, increasing the temperature to 75 ℃ resulted in highly variable particle sizes which were not statistically different from the sizes obtained at 25 ℃ or

40 ℃ (Figure 12).

1 1 0

p < 0 .0 1

)

(

e t

e 9 0

i t

r 7 0

e M 5 0

5 0 5 2 4 7

T e m p e r a tu re  C

In addition, an increase in the internal diameter of the needle from 27 G to 20 G resulted in a significant decrease in the mean particle diameter (Figure 13). Although the effect of the needle size was not highly significant (p = 0.048), the use of the larger diameter needle decreased the time required for the addition of the organic phase to the

41 aqueous phase from an average of 40 min to an average of 2 min during nanoparticle preparation. The effect of increasing needle diameter on nanoparticles size reduction could be explained by the better distribution of the organic solvent into the aqueous phase associated with using a larger needle diameter. Molpeceres et al. reported that the higher flow rate of the organic solvent into aqueous phase resulting from the larger diameter needle produced smaller size particle. However, variation of needle diameter and flow rate of organic solvent did not explain formation of nanoparticles < 100 nm57.

) 70

(

r p =0.048

a i

d 60

a n

e M

G G 7 0 2 2 le le d d e e e e N N Figure 13. Effect of internal needle diameter on the mean size of nanoparticles prepared using surfactant-free nanoprecipitation method (p value from unpaired t-test with n=3 and the error bars represent standard deviation).

As an additional observation, the use of the Amicon® centrifugation tube in the collection of nanoparticles resulted in the appearance of visually detectable aggregates that needed to be removed by filtration before measuring the final particle size. This additional filtration step resulted in a highly variable nanoparticle yield and low nanoparticle recovery. On the other hand, the collection of nanoparticles without centrifugation was highly effective and reproducible with a yield percent of ~ 75 % relative to the amount of polymer used in the preparation.

Hence, the conditions listed under group (8B) in Table 3 were used as optimal conditions to prepare Nile Red loaded PLGA nanoparticles for further experiments in these studies. These conditions include dissolving 36 mg of PLGA in 5 ml DMF containing 700 µl (equivalent to 70 g) of Nile Red solution (100 g/ ml) in acetone.

That solution was added to 30 ml Nanopure® water (preheated to 40 ℃) using a 20 ml syringe with a 20 G needle under medium magnetic stirring (~300 rpm). The solvent was removed by dialysis against water for ~40 h and the nanoparticle dispersion was collected from the dialysis bag and transfered to a 50 ml polypropylene tube and sealed for further analyses.

4.2.1 Particle Characterization

The intensity-weighted particle size measurements were performed using a

NICOMP 380 ZLS, Nicomp particle sizing system (Santa Barbara, California, USA) or a Zetasizer Nano-ZS, (Malvern Instruments Ltd. Worcestershire, UK). Figure 14 & 15 show the measurements of the mean particle size for two nanoparticle batches using both instruments, respectively. The detailed measurements for particle size are described in

Appendix F.

Figure 14. Example of intensity-weighted nanoparticles size distribution for nanoparticle batch 3B 12 (Table F 1) using a Malvern Zetasizer Nano-ZS (the average size is 62 nm).

Figure 15. Example of intensity-weighted nanoparticles size distribution for nanoparticle batch 1B 1 (Table F 1) using Nicomp380 ZLS Zetasizer (mean particle size =94.01 nm).

The polydispersity index (PDI), a measurement of the heterogeneity of the distribution, for the nanoparticles prepared using the surfactant-free nanoprecipitation method (Table F 1) was 0.112  0.050 (mean  SD).

The zeta potential, the measurement of the particles’ electrophoretic mobility, was measured with the Malvern Nano-ZS. The zeta potential for the nanoparticles prepared using the optimized conditions was -27.9  5.4 mV (mean  SD) as shown in Figure 16.

Zeta Potential Distribution

160000 140000 120000 100000 80000 60000 40000 20000 0 -200 -100 0 100 200 Zeta Potential (mV)

Figure 16. Zeta potential measurement of nanoparticle batch 8B 18 (Table F 1) showing a value of -26.3 mV (measured with Malvern Zetasizer Nano ZS).

SEM was used to analyze the shape and size of the nanoparticles prepared using the optimized conditions. The selected group (8B 22 Table F1) had an average hydrodynamic diameter (measured with dynamic light scattering) of 62 nm and 69 nm before and after the addition of the glucose 5 % (w/v) solution, respectively. The images in Figure 17 show spherical nanoparticles with an average size of 50-70 nm, similar to the size range obtained from the dynamic light scattering measurements.

A B

C D

4.2.2 Effect of Dispersion Medium on Particle Size

Nanoparticles were unstable in the presence of buffer solutions as an incubation medium, and visual flocculation was observed after the addition of all three buffer solutions used in this study (KRB with added glucose, PBS and TBS). This is probably due to the ions in the buffer solutions bridging small size nanoparticles together and resulted in flocculation58,59.

In comparison, no flocculation was observed using glucose 5 % (w/v) solution as a dispersion and incubation medium. In order to investigate the effect of the glucose medium on the particle size, size was measured before adding the glucose and after a two hour incubation in glucose 5 % (w/v) solution at 37 ℃ for three nanoparticles batches

prepared using the same optimized surfactant-free method. The change in size was non-

) m

significant (p > 0.05) as shownn in Figure 18.

(

e 7 0

p o

n 6 0

n n n i i

a d d

e e e d d

M 5 0 n n n e e o p p i s s t u u u l s r s o e s s t s e a e l e l % c r w c i  i 5 t u t r p r e a a s o p p o n c o a o u n n l a N a g N N

Figure 18. Effect of 5 % glucose medium on nanoparticle size (no significant difference was observed by unpaired t-test, n=3 for each group, error bars represent standard deviation).

4.2.3 Calculation of Nanoparticle Yield

The percent yield was ~12 % for nanoparticle batches collected using the

Amicon® centrifuge tube followed by filtration using a 0.22 µm syringe filter. The yield varied by more than 5 % between triplicate batches. In comparison, the yield was ~75 % for the nanoparticles prepared and collected without any further centrifugation and filtration processes and the variation in yield between replicates was less than 3 %. The detailed calculations for nanoparticle yield are shown in Appendix C.

4.2.4 Calculation of Nanoparticle Loading

Nile Red loading/ recovery was 26 ± 0.4 % of the original amount (70 g added to the polymer solution) used in the preparation of surfactant-free nanoparticles. The particle size measurements of the nanoparticle dispersion after the addition of the

Cellosolve® Acetate indicated the nanoparticles required at least 6 hours to completely dissolution. Figure 19 shows the particle size measurement 2 hours after adding

Cellosolve® Acetate, and a small peak at ~ 70 nm indicates that the nanoparticles were not completely dissolved. On the other hand, Figure 20 shows the particle size measurement 6 hours after solvent addition and there was no peak at 50 nm, or at any other size, indicated the nanoparticles were dissolved. The observed peak at ~ 750 nm was with a very high poly-dispersity index (value = 0.95), which is out of the range of the

Malvern Zetasizer instrument (upper limit = 0.7 and typical value 0.05). The same peak was observed when Cellosolve® Acetate solvent and pure water were evaluated for particle size.

Size Distribution by Intensity 40

0 0.1 1 10 100 1000 10000 Size (d.nm)

Figure 19. Nanoparticle size measurement after two hour incubation with Cellosolve® Acetate. The small peak at ~70 nm indicates some ~ 70 nm nanoparticles remain intact.

30 20

0 0.1 1 10 100 1000 10000 Size (d.nm)

4.3 Lucifer Yellow VS Transport

Results from the cumulative transport of Lucifer Yellow VS (Figure 21) and the

Lucifer Yellow VS transport (%) at 1.5 h (Figure 22) demonstrate ~ 0.6 % Lucifer

Yellow VS transport across the olfactory mucosa and ~ 0.1 % transport across the respiratory mucosa. This confirmed the tissues remain intact and biologically active for at least 2 hours after tissues retrieval.

0 .6

p s

n 0 .4 R e s p ira to ry

a r

t O lfa c to ry

f 0 .2

u o

0 .0

m a

2 0 4 0 6 0 8 0 1 0 0

% T im e (m in ) -0 .2 Figure 21. Cumulative amount of Lucifer Yellow VS transported across olfactory and respiratory mucosae. The amount is reported as percent of Lucifer Yellow VS in the receiver side compared to the amount of Lucifer Yellow VS in the donor side. (n=3 for each data point, error bars represent standard deviation).

t r

o 0 .8

p R e s p ira to ry

s n

a O lfa c to ry

r 0 .6

o l

l 0 .4

r e

f 0 .2

u L 0 .0

% y r y o r t o a t r c i a p lf s e O R T is s u e ty p e

Figure 22. Percent of Lucifer Yellow VS transported across olfactory and respiratory mucosae after 90 min incubation time (n=3 for each tissue type, error bars represent standard deviation).

4.4 Quantification of Nanoparticle Uptake

The number of nanoparticles entrapped in the nasal mucosal tissues, measured using the corresponding amount of recovered Nile Red dye was calculated based on the previously determined correlation between particle mass and entrapped dye amount

(Section 3.8 and Appendix C). The Nile Red concentration was calculated after measuring the fluorescence intensity (Em.: 520, Ex.: 620 nm) after dissolving the nanoparticles in Cellosolve® Acetate.

In the respiratory tissue, the total number of nanoparticles recovered in the tissues were 2.3 % and 5.0 %, after 30 and 60 minutes, respectively, of the number of nanoparticles placed in the donor side. Of these ~ 30 % and ~ 70 % of the nanoparticles were collected from the epithelial cells and the sub-mucosal layer, respectively. Figures

23 & 24 show the results for the nanoparticle uptake in the nasal respiratory tissues after

30 and 60 minutes, respectively. The nanoparticle uptake in full thickness tissues

(epithelial cells + sub-mucosal layer) was time dependent with a greater number of nanoparticles recovered after 60 min as compared to 30 min (Figure 25). Nanoparticles recovered from the receiver only represented 0.19 % and 0.15 % of the total nanoparticle load in the donor side after incubation for 30 and 60 minutes, respectively

0 5 5 0

)

(

4 5 0

f p < 0 .0 0 1

r 1 0

e b

m 5

u N n r e 0 io s ll y s r a r e l e c l e v p i l a s e ia s i c l o d e e c P R h u it N m p - r E b o u n o S D

Figure 23. Number of PLGA nanoparticles (~ 70 nm) recovered from the respiratory mucosa after exposure to a nanoparticle dispersion for 30 min (p value from the unpaired t-test (n=3)).

0 5 5 0

)

(

4 5 0

P N

p < 0 .0 5

r 2 5

e b

1 5

m u

N 5 n r e io s ll y s r a r e l e c l e v p i l a s e ia s i c l o d e e c P R h u it N m p - r E b o u n o S D

Figure 24. Number of PLGA nanoparticles (~ 70 nm) recovered from the respiratory mucosa after exposure to a nanoparticle dispersion for 60 minutes (p value from the unpaired t-test (n=3)).

40 p < 0.05

)

1 0

1 30

(

P N

r e

b 10

u N

0 in in m m 0 0 3 6

Figure 25. Number of PLGA nanoparticles (~ 70 nm) recovered from the full thickness respiratory mucosa after 30 and 60 min incubation periods (p value from unpaired t-test (n=3)).

In the olfactory mucosa, the total nanoparticle uptake (Figures 26 & 27) was 1.8 % and 3.8 % of the total nanoparticle load in the donor side after incubation for 30 and

60 minutes, respectively. Approximately 30 % and 70 % of the particles were recovered from the epithelial cells and sub-mucosal layer, respectively, in both experiments. The total number of nanoparticles recovered from the full thickness olfactory mucosa

(Figure 28) was time dependent, where a greater number of nanoparticles was recovered after 60 minutes as compared to 30 minutes. There was no detectable number of nanoparticles recovered from receiver after 30 and 60 minutes.

6 0 0

)

(

P 4 0 0

f o

1 5

b m

u 5 N

n r o s e i r ll y s e e a r l e iv c l p e l a s c ia s i e l o d e R c h u P it m N p - r E b o u n S o D

6 0 0

)

(

P 4 0 0

f o

1 5

b m

u 5 N

n r o s e i r ll y s e e a r l e iv c l p e l a a s is c i e l o d e c R h u P it m N p - r E b o u n S o D

25 p < 0.05

)

1 1

0 20

(

P 15

f o

e b

m 5

u N 0 in in m m 0 0 3 6

Figure 28. Number of nanoparticles recovered from full thickness olfactory mucosa (p value from unpaired t-test (n=3)).

Since the respiratory and olfactory tissues have significantly different thickness, the nanoparticle uptake was normalized per gram of tissues. After 60 min, the number of nanoparticles recovered per gram of full thickness (epithelial cells and sub-mucosal layer) nasal mucosa (Figure 29) was higher in the olfactory mucosa as compared to the respiratory mucosa. This suggests that the unmodified PLGA nanoparticles in the size range of 50-70 nm have greater ability to translocate into the olfactory mucosa than the respiratory mucosa. This higher uptake by the olfactory tissue may be due the anatomical differences of the epithelial layer such as differences in cell types between respiratory and olfactory epithelium. In particular, the junctions between the neurons and supporting cells may have different reduced barrier properties compared to the epithelial cell- epithelial cell junctions in the nasal respiratory mucosa.

60 p < 0.05

g Olfactory

/ 40

) 2

1 Respiratory

(

f 20

m u N 0

in in m m 0 0 3 6 Figure 29. Comparison of PLGA nanoparticle (~ 70 nm) uptake in olfactory and respiratory nasal mucosal tissues normalized to tissue mass (p value from unpaired t-test (n=3)).

4.5 Conclusion

This study has demonstrated the effect of preparation parameters, polymer amount, surfactant type, temperature of the aqueous phase and the internal needle diameter, on the resulting size of nanoparticles prepared from the well-characterized, biodegradable, biocompatible polymer, PLGA, using nanoprecipitation methods. The use of a surfactant- free nanoprecipitation method for the preparation of Nile Red-loaded

PLGA nanoparticles with size < 100 nm was found to be an efficient and reproducible method. The surfactant-free nanoprecipitation method was an appropriate method to prepare small, relatively stable Nile Red-loaded PLGA nanoparticles. Use of DMF as an organic solvent and selecting a 40 ℃ aqueous phase temperature were shown to provide optimal conditions for preparation of polymeric nanoparticles in the size range of 50-70 nm using the surfactant-free nanoprecipitation method.

The number of nanoparticles were quantified using a dye extraction method involving Cellosolve® Acetate as a solvent for PLGA and the released-dye intensity was measured and correlated with the amount of dye per particle. Using this released dye quantification method, in vitro studies of the uptake of Nile Red-loaded PLGA nanoparticle into bovine nasal mucosa showed that these small nanoparticles can permeate into the nasal mucosa within 30 min of exposure, yet only in a relatively small number (< 5 % of nanoparticles presented to the mucosal surface). A greater number of particles appears to locate within the submucosa compared to the epithelial cells.

Normalization of the quantified nanoparticle number with respect to the mass of tissue revealed the greater uptake of nanoparticles in the olfactory mucosa after 60 min as

59 compared to the respiratory mucosa. This greater uptake suggested that the tissue type plays a significant role in nanoparticle uptake which may be useful for further targeted delivery of antigens. The rapid and quantitative transfer of PLGA nanoparticles into the nasal mucosal tissues indicates that they may be used as a delivery vehicle for drugs for either local or systemic activities, yet the low transfer efficiency may need further evaluation of the clinical utility of these delivery system.

Appendix A: Standard Curves

Nile Red Standard Curve in Cellosolve® Acetate

A Nile Red standard curve in Cellosolve® Acetate was prepared by measuring fluorescence intensity at various concentrations at (Ex.:520 – Em.:620 nm) using a

SpectraMax M5 Multi-Mode Plate Reader (Molecular Devices, Sunnyvale, California,

USA). Measurements (Table A 1) were performed in triplicate for each concentration in three different wells in a 96-well quartz plate. One concentration was studied at a time and the average intensity of three measurements was calculated for each concentration.

Before measuring fluorescence intensity for the Nile Red solutions, measurements were performed first with the blank wells and then with wells containing solvent only to avoid any interference from the background with our measurements. Minitab17 software was used to determine detection limit and to create simple linear regression Model.

Table A 1. Fluorescence intensity of Nile Red standard solutions in Cellosolve® Acetate (measurements were performed using 520-620 nm for the excitation and emission wavelengths, respectively).

Intensity Nile Red Solution concentration name Measurement Measurement (ng/ml) Measurement 3 1 2

Blank 0 0.3 0.4 0.4

Solvent 1.4 1.4 1.6 only 0

Dilution 11 0.20 3.0 2.5 2.8

Dilution 10 0.39 4.1 5.9 5.2

Dilution 9 0.78 6.9 6.7 7.3

Dilution 8 1.56 12.4 11.8 13.4

Dilution 7 3.13 21.6 23.4 23.4

Dilution 6 6.25 43.3 44.8 45.3

Dilution 5 12.5 88.1 88.4 90.6

Dilution 4 25 176.0 178.4 179.7

Dilution 3 50 353.3 355.1 349.1

Dilution 2 100 697.2 720.6 749.1

Dilution 1 200 1396.6 1301.4 1392.1

Stock 2557.0 2608.1 2621.8 solution 400

In order to test the lower detection limit of Nile Red, mean fluoresce intensities were compared for blank wells, solvent only wells and the lowest four concentrations using ANOVA with Tukey’s pairwise comparison as shown in (Figure A 1). The lowest detection limit was determined to be 0.39 ng/ml. The regression line (Figure A 2) was performed with Minitab 17.

Tukey’s Pairwise Comparisons Grouping Information Using the Tukey’s Method and 95% Confidence

Solution name N Mean Grouping Dilution 12 (1.563 ng/ml) 3 12.5 A Dilution 13 (0.781 ng/ml) 3 6.9 B Dilution 14 (0.391 ng/ml) 3 5.1 C Dilution 15 (0.195 ng/ml) 3 2.8 D Solvent only (0.0 ng/ml) 3 1.5 D E Blank (0.0 ng/ml) 3 0.4 E

Figure A 1. Minitab output for Tukey’s pairwise comparison Analysis of the means (means that do not share letter are significantly different).

Figure A 2. Example of standard curve for the Nile Red solutions in Cellosolve® Acetate.

Standard Curve for Lucifer Yellow VS in 5 % Glucose Solution

A Lucifer Yellow VS Standard curve in glucose 5 % solution was prepared by measuring fluorescence intensity at various concentrations at (Ex.: 430- Em.:530 nm) using a SpectraMax M5 Multi-Mode Plate Reader (Molecular Devices, Sunnyvale,

California, USA). Measurements (Table A 2) were performed in triplicate for each concentration in three different wells in a 96-well polypropylene plate and the average intensity was calculated for each concentration. Before measuring fluorescence intensity for the Lucifer Yellow solutions, measurements were performed first with blank wells and then with wells containing solvent only to make sure that there is no interference from the background with our measurements. Minitab17 software was used to determine the detection limit of Lucifer Yellow VS and to create the simple linear regression model.

Table A 2. Fluorescence intensity Lucifer Yellow VS in glucose 5 % (w/v) solutions (measurements were performed using 430-530 nm for the excitation and emission wavelengths, respectively).

Name (concentration) Measurement 1 Measurement 1 Measurement 1

Blank 6.1 6.7 8.8

Glucose 5 % (w/v) solution 6.9 6.3 6.6

Dilution 14 (0.38 ng/ml) 6.2 6.5 6.6

Dilution 13 (0.76 ng/ml) 6.6 7.0 7.1

Dilution 12 (1.53 ng/ml) 7.2 7.2 7.6

Dilution 11 (3.05 ng/ml) 8.9 8.8 10.8

Dilution 10 (6.1 ng/ml) 12.1 12.1 12.6

Dilution 9 (12.21 ng/ml) 18.1 17.9 19.0

Dilution 8 (24.41 ng/ml) 37.0 38.9 38.3

Dilution 7 (48.83 ng/ml) 39.9 41.3 39.7

Dilution 6 (97.66 ng/ml) 75.9 78.0 78.6

Dilution 5 (195.31 ng/ml) 147.4 150.5 144.3

Dilution 4 (390.63 ng/ml) 297.6 298.9 300.9

Dilution 3 (781.25 ng/ml) 577.9 582.4 571.8

Dilution 2 (1562.5 ng/ml) 1198.3 1174.9 1143.3

Dilution 1 (3125 ng/ml) 2201.1 2278.4 2293.0

Stock solution (6250 ng/ml) 4559.9 4512.2 4333.6

In order to calculate the lowest detection limit of Lucifer Yellow VS, fluoresce emission intensities were compared for blank wells, solvent only wells and the lowest seven concentrations from the standard curve dilutions and were tested using one-way

ANOVA and Tukey’s pairwise analysis as shown in (Figure A 3). The Lower detection limit was 3.1 ng/ml. The regression line (Figure A 4) was performed with Minitab 17.

Tukey’s Pairwise Comparisons

Grouping Information Using the Tukey’s Method and 95% Confidence

Name Concentration N Mean Grouping Dilution 8 (24.41 ng/ml) 3 38.1 A Dilution 9 (12.21 ng/ml) 3 18.3 B Dilution 10 (6.10 ng/ml) 3 12.3 C Dilution 11 (3.05 ng/ml) 3 9.5 D Dilution 12 (1.53 ng/ml) 3 7.4 E Blank (0.0 ng/ml) 3 7.2 E Dilution 13 (0.76 ng/ml) 3 6.9 E Glucose 5 % solution 3 6.6 E Dilution 14 (0.38 ng/ml) 3 6.4 E

Figure A 3. Minitab output for the Tukey's pairwise comparison of the mean intensity (means that do not share a letter are significantly different).

Figure A 4. Standard curve for the Lucifer Yellow VS in glucose 5 % (w/v) solution.

Appendix B: Calculation of the PLGA Polymer Density

1- The density of the water was chosen to be 0.998 g/cm3 for liquid state and 20 C.

mass ρ = 2- volume

13.275 g 3- Volume of water without adding PLGA polymer ( = 13.302 cm3) 0.998 g/cm3

13.287 g 4- Volume of water after adding the PLGA polymer ( = 13.314 cm3) 0.998 g/cm3

5- Volume of PLGA used (13.3136 - 13.30124) = 0.012 cm3

6- Mass of the PLGA in air = 0.014 g

0.014 g 7- ρ (PLGA) = = 1.16 g/cm3 0.012 cm3

Appendix C: Calculation of PLGA Nanoparticles Number, Yield and Loading

Calculation of the Number of Nanoparticles

1- Starting mass of PLGA used to prepare nanoparticles = 36 mg

2- The mean mass of the PLGA remaining after preparation = 27.47 mg

(calculated in Table C 1 below)

3- Mean diameter (for nanoparticles batch under investigation) = 55 nm

 Radius= 27.5 nm

4- Mass of an individual (55 nm diameter) PLGA nanoparticle = ρ *(4/3 π풓ퟑ)

= (1.16 g/cm3) *(4/3 π (2.75 * 10-6 cm)3

= 1.02 * 10-17 g

mass of PLGA remaining after preparation 5- Number of PLGA nanoparticles = mass of single nanoparticle

= (0.0275 g/ 1.02* 10-17 g)

= 2.7* 1015 particles

this number represents the approximate total number of nanoparticles in a batch of nanoparticle prepared using surfactant-free nanoprecipitation method that had a diameter of 55 nm.

Quantification of the Amount of Dye per Particle

1- The total amount of Nile Red after preparation was calculated as follow: 1 ml aliquot

of nanoparticles suspension was incubated with Cellosolve® Acetate for ~ 8 h in order

to dissolve PLGA nanoparticles and extract Nile Red content. The amount of Nile

Red was quantified by measuring the fluorescence intensity and the amount of Nile

Red was calculated using appropriate standard curve. The amount of Nile Red in 1 ml

of nanoparticle suspension and total volume of nanoparticle suspension after

preparation were used to calculate the total amount of Nile Red as following: a- Amount of Nile Red in 1 ml of nanoparticle suspension (calculated from standard

curve) = 578.82 ng b- Volume of nanoparticle suspension after preparation = 32 ml c- Total amount of Nile Red after preparation (in 32 ml) = 578.82 ng * 32

= 18522.24 ng/ 32 m

2- Amount of dye per single nanoparticle (for nanoparticle batch that had diameter of 55

nm) was calculated from as follow:

total amount of Nile Red after preperation Amount of Nile Red per perticle = total number of nanoparticles total amount of Nile Red after preparation = 18522.24 ng total number of nanoparticles in the same nanoparticles batch = 2.7 * 1015 particles Amount of Nile Red per particle = 18522.24 / (2.7 * 1015) = 6.9 *10-12 ng of Nile Red / particle

Calculation of Nanoparticle Yield

The nanoparticle yields for the nanoparticles prepared using the surfactant free nanoprecipitation method was calculated by lyophilizing three batches prepared and collected using the same conditions in pre-weighed 15 or 50 ml polypropylene tubes. The percent yield was calculated by comparing the amount of PLGA remaining after lyophilization with the initial amount added to the preparation of (in this study the initial amount of PLGA was 10 or 36) as shown in Table C 1.

Batch Starting PLGA Group (2B*) Starting PLGA Group (8B**) order amount (mg) amount (mg) (yield amount) (yield amount)

1 10.23 1.8 36.57 27.01

2 10.33 0.6 37.24 27.49

3 10.31 1.1 36.59 27.911

Mean 10.29 1.16 36.8 27.47

Yield 11.3% 74.65% (± 1.4 %) percent

* Nanoparticles prepared by surfactant free nanoprecipitation method and collected with Amicon centrifugation tube (for batches 2B see page 85) ** Nanoparticles prepared by surfactant free nanoprecipitation method and collected from dialysis bag into a 50 ml polypropylene tube. (for batches 8B see page 85)

Appendix D: Preparation of Buffer Solutions

Phosphate Buffer Saline (PBS) (pH7.4)

PBS was prepared by mixing the materials in Table D 1 with 900 ml Nanopure® water. The pH was adjusted to 7.4 with 1N HCl or 0.1N NaOH and the volume was completed to one liter with adequate amount of Nanopure® water60.

Table D 1. Chemicals used to prepare one liter PBS.

Chemicals Amount Concentration (g) (mM)

1 Sodium chloride (NaCl) [Research Products 8 137 International Corp., Mt. Prospect, IL, USA]

2 Potassium chloride (KCl) [Research Products 0.2 2.7 International Corp., Mt. Prospect, IL, USA]

3 Sodium phosphate dibasic (Na2HPO4) 1.44 10 [Research Products International Corp., Mt. Prospect, IL, USA]

4 Potassium phosphate monobasic (KH2PO4) 0.24 1.76 [Research Products International Corp., Mt. Prospect, IL, USA]

KRB (Krebs-Ringer Bicarbonate Buffer) with Added Glucose

KRB was prepared by mixing materials 1 through 7 listed in Table D 2 in 900 ml of Nanopure® water. The mixture was bubbled with a Carbogen (95 % oxygen and 5 % carbon dioxide) for 10 minutes to equilibrate CO2/Carbonic acid in the system. Calcium chloride was added and the pH was adjusted with 0.1 N NaOH or 1 N HCl to 7.4. The volume was completed to one liter with Nanopure® water6.

Table D 2. Concentration of chemicals in one liter KRB (pH adjusted to 7.4). Chemicals Amount Concentration

(g) (mM)

D-Glucose [Research Products International Corp., Mt. 1.8 10.00 1 Prospect, IL, USA] Magnesium chloride (MgCl ) [Sigma-Aldrich, St. Louis, 0.16 1.67 2 2 MO, USA] Sodium chloride (NaCl) [Research Products International 6.95 119.78 3 Corp., Mt. Prospect, IL, USA] Potassium chloride (KCl) [Research Products 0.34 4.56 4 International Corp., Mt. Prospect, IL, USA] Sodium phosphate dibasic (Na HPO ) [ Research 0.12 0.83 5 2 4 Products International Corp., Mt. Prospect, IL, USA] Sodium phosphate monobasic (NaH PO [Research 0.18 1.5 6 2 4) Products International Corp., Mt. Prospect, IL, USA] Sodium bicarbonate (NaHCO ) [ Research Products 1.26 14.99 7 3 International Corp., Mt. Prospect, IL, USA] Calcium Chloride (CaCl ) [EM science, Gibbstown, NJ, 0.13 1.2 8 2 USA]

Tris Buffered Saline (TBS) pH 7.4

Tris base (Research Products International Corp., Mt. Prospect, IL, USA) with a concentration of 10 mM (1.21 g) and 150 mM (8.7 g) sodium chloride (NaCl) was dissolved in 900 ml Nanopure® water. The pH was adjusted to 7.4 with 1N HCl and the volume was completed to one liter with Nanopure® water60.

Appendix E: Nanoparticle Uptake Data

Olfactory Uptake Data

Table E 1. Nanoparticle uptake in the olfactory mucosa.

Time Tissue Mean *Nile Red **Number of Weight of (min) fluorescent (ng/ml) nanoparticles tissue (mg) intensity (x1011)

30 Epithelial 19.43 0.68 0.75 51.54

42.45 4.20 4.65 29.22

38.28 3.56 3.94 27.32

30 Sub-mucosal 28.46 2.06 2.28 51.54

75.45 9.25 10.23 29.22

56.26 6.31 6.98 27.32

60 Epithelial 47.41 4.95 5.48 44.9

73.91 9.01 9.97 37.06

37.23 3.40 3.76 42.62

60 Sub-mucosal 98.39 12.75 14.10 44.9

73.53 8.95 9.90 37.06

106.41 13.98 15.46 42.62

* Nile Red concentration was calculated from regression equation: [Intensity = 15 +6.538 * concentration] ** Number of nanoparticles was calculated by comparing Nile Red concentration with the previously determined amount of dye per single particle (value of 9.04 * 10-12 ng/particle)

Respiratory Uptake Data

Table E 2. Nanoparticle uptake in the respiratory mucosa.

Time Tissue Fluorescent *Nile Red **Number of Weight (min) intensity concentration nanoparticles of tissue (ng/ml) (x 1011) (mg)

30 Epithelial 51.53 4.73 4.30 48.29

44.67 3.92 3.55 53.93

46.58 4.14 3.76 44.25

30 Sub-mucosal 89.85 9.31 8.45 48.29

83.98 8.61 7.82 53.93

87.49 9.03 8.20 44.25

60 Epithelial 70.90 7.05 6.40 79.51

58.19 5.53 5.02 56.7

97.10 10.18 9.24 60.28

60 Sub-mucosal 239.02 27.14 24.62 69.26

163.91 18.16 16.48 56.7

130.50 14.17 12.86 60.28

* Nile Red concentration was calculated from regression equation: [Intensity = 11.899+8.37 * concentration] ** Number of nanoparticles was calculated by comparing Nile Red concentration with the previously determined amount of dye per single particle (value of 1.10 * 10-11 ng/particle)

Appendix F: Nanoparticle Size Measurement Data

Nanoparticles Prepared using the Nanoprecipitation Method

Note: Graphs are defined by the code name mentioned in table (1)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm)

Size measurement for experiment 1A (average size =107 nm, PDI=0.057)

Size measurement for experiment 2A (average size =161 nm)

Size measurement for experiment 3A (average size = 96 nm, PDI = 0.055)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 4A (average size =140 nm, PDI= 0.045)

Size measurement for experiment 5A (average size =163 nm)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (r.nm) Size measurement for experiment 6A (average size d =178 nm, PDI= 0.056)

Size measurement for experiment 7A (average size =137 nm, PDI= 0.038)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 8A (average size =162 nm, PDI= 0.018)

Size measurement for experiment 9A (average size = 100.5 nm)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 10A (average size =161 nm, PDI= 0.022)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 11A (average size = 98 nm, PDI= 0.033)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 12A (average size =146 nm, PDI= 0.064)

Size measurement for experiment 13A (average size =164.7 nm, PDI= 0.078)

Size measurement for experiment 14A (average size =173.4 nm, PDI= 0.015)

Size measurement for experiment 15A (average size =157 nm, PDI= 0.082)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 16A (average size =162 nm, PDI= 0.072)

Nanoparticles Prepared with the Surfactant-Free Nanoprecipitation Method Table F 1. Data for individual nanoparticle batches prepared by surfactant-free nanoprecipitation method Code Order PLGA DMF Aqueous Temp. Needle Particle size Name (mg±0.5) (ml) phase (ml) (℃) Gauge (nm) (G)

1B 1 10 2 20 25 27 94 2 10 2 20 25 27 92 3 10 2 20 25 27 79 2B 4 10 2 20 40 27 68 5 10 2 20 40 27 68 6 10 2 20 40 27 72 7 10 2 20 40 27 62 8 10 2 20 40 27 64 9 10 2 20 40 27 71 3B 10 10 2 40 75 27 82 11 10 2 40 75 27 77 12 10 2 40 75 27 62 7B 13 36 5 30 40 27 57 14 36 5 30 40 27 66 15 36 5 30 40 27 64 8B 16 36 5 30 40 20 50 17 36 5 30 40 20 57 18 36 5 30 40 20 53 19 36 5 30 40 20 51 20 36 5 30 40 20 57 21 36 5 30 40 20 54 22 36 5 30 40 20 61 23 36 5 30 40 20 61

Note: Graphs are coded by the group name and order number, for example (1B 1)

Size measurement for experiment 1B 1 (average size = 94.1 nm)

Size measurement for experiment 1B 2 (average size = 91.6 nm)

Size measurement for experiment 1B 3 (average size = 79 nm)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 2B 4 (average size = 67.64 nm, PDI= 0.153)

Size measurement for experiment 2B 5 (average size = 67.8 nm)

Size measurement for experiment 2B 6 (average size = 72.4 nm)

Size measurement for experiment 2B 7 (average size = 62.4 nm)

measurement for experiment 2B 8 (average size = 64.4 nm)

measurement for experiment 2B 9 (average size = 71.4 nm)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 3B 10 (average size = 82.44 nm, PDI= 0.038)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 3B 11 (average size = 77.01 nm, PDI= 0.074)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 3B 12 (average size = 62.44 nm, PDI= 0.09)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 7B 13 (average size = 56.96 nm, PDI= 0.213)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 7B 14 (average size = 65.69 nm, PDI= 0.142)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 7B 15 (average size = 64.25 nm, PDI= 0.072)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 8B 16 (average size = 50.34 nm, PDI= 0.119)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 8B 17 (average size = 56.95 nm, PDI= 0.193)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 8B 19 (average size = 53.23 nm, PDI= 0.078)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 8B 20 (average size = 50.50 nm, PDI= 0.096)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 8B 21 (average size = 56.72 nm, PDI= 0.17)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm) Size measurement for experiment 8B 22 (average size = 53.64 nm, PDI= 0.088)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm)

Size measurement for experiment 8B 23 (average size = 60.53 nm, PDI= 0.087)

Size Distribution by Intensity

0 0.1 1 10 100 1000 10000 Size (d.nm)

Size measurement for experiment 8B 24 (average size = 60.78 nm, PDI= 0.081)

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