Uptake and distribution of ultrafine nanoparticles and ...

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Uptake and distribution of ultrafinenanoparticles and microemulsions from thenasal mucosaBejgum, Bhanu Chanderhttps://iro.uiowa.edu/discovery/delivery/01IOWA_INST:ResearchRepository/12730576490002771?l#13730717490002771

Bejgum. (2019). Uptake and distribution of ultrafine nanoparticles and microemulsions from the nasalmucosa [University of Iowa]. https://doi.org/10.17077/etd.h0z4bv8z

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Uptake and Distribution of Ultrafine Nanoparticles and Microemulsions from the Nasal

Mucosa

by

Bhanu Chander Bejgum

A thesis submitted in partial fulfillment of the requirements for the Doctor of

Philosophy degree in Pharmacy (Pharmaceutics) in the Graduate College of

The University of Iowa

August 2017

Thesis Supervisor: Professor Maureen D. Donovan

Copyright by


2017

All Rights Reserved

Graduate College The University of Iowa

Iowa City, Iowa

CERTIFICATE OF APPROVAL

_______________________

PH.D. THESIS

_______________

This is to certify that the Ph.D. thesis of


has been approved by the Examining Committee for the thesis requirement for the Doctor of Philosophy degree in Pharmacy (Pharmaceutics) at the August 2017 graduation.

Thesis Committee: ___________________________________ Maureen D. Donovan, Thesis Supervisor

___________________________________ Douglas R. Flanagan

___________________________________ Aliasger K. Salem

___________________________________ Lewis L. Stevens

___________________________________ Laura B. Ponto

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To my family

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ACKNOWLEDGMENTS

I would like to express my heartfelt thanks to all the people who have been

involved either directly or indirectly, at various stages of my journey at The University of

Iowa (UIowa). I may not be able to list all the names here, but I sincerely appreciate all

the unforgettable help, support, advice, and encouragement I have received at UIowa.

First and foremost, I would like to express my deepest gratitude and appreciation

to my advisor, Professor Maureen Donovan for her continuous support, guidance, and

mentoring throughout my graduate career. Her expertise, enthusiasm and feedback

constantly assisted me in the right direction and led me towards the completion of my

dissertation research. Along with the academic and research skills, she has constantly

encouraged and supported me to take up the leadership roles and all other necessary skills

that are useful for my career. It has been an honor to work with her.

Secondly, I would like to acknowledge and appreciate my dissertation committee

members, Dr. Douglas Flanagan, Dr. Aliasger Salem, Dr. Lewis Stevens and Dr. Laura

Ponto for their valuable suggestions and guidance in my research, and for their time on

my dissertation. I would also like to thank Dr. Jennifer Fiegel for her help and input on

my comprehensive exam.

I would like to thank Central Microscopy Research Facility (CMRF) for assisting

me with the confocal and electron microscopy. I would also like to acknowledge the help

from staff of Small Animal Imaging Facility, especially Susan Walsh for her assistance in

fluorescence imaging and micro-CT studies. I am extremely grateful to Dr. Sarah Larsen

for allowing me to use inductively coupled plasma optical emission spectroscopy and Dr.

Aliasger Salem for allowing me to use Malvern Zetasizer. Furthermore, I am extremely

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thankful to all the faculty and staff members of the Division of Pharmaceutics and

Translational Therapeutics, College of Pharmacy, for their valuable support during my

graduate studies.

I thank Dr. Keith Guillory and pharmaceutics faculty for offering my financial

support during my graduate career in the form of Keith Guillory Fellowship, research and

teaching assistantships. I would like to thank graduate school of UIowa for awarding the

Summer Dissertation Fellowship.

I would like to express my gratitude to all former and current lab members in Dr.

Donovan’s lab including Nan Chen, Ana Ferreira, Wisam Al-bakri, Varsha Dhamankar,

Rakesh Awasthi, Maya George, Krupal Maity, Namita Sawant, and Shanti Chade. Also, I

thank all my friends at UIowa who made my graduate study a memorable one. They all

have been a valuable source of friendship and advice over the past years.

Last but not least, this work would not have been completed without the love and

support from my family and friends. I would like to thank my dad (Ramesh), mom

(Vanaja), brother (Bharath), and sister (Priyanka) for their unconditional love and

encouragement. Finally, I would like to thank my wife (Manasa) for her love and support.

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ABSTRACT

Various colloidal delivery systems, including polymeric nanoparticles, metal

colloids, liposomes, and microemulsions have been reported to enhance the delivery of

therapeutic agents following intranasal administration. However, the mechanisms

involved in the uptake of these nanomaterials, especially those in the ultrafine size ranges

(diameter < 20 nm) through nasal mucosa and their subsequent biodistribution in the

body are not well characterized. The objectives of this study address the knowledge gap

regarding ultrafine nanoparticle transfer in the nasal mucosa by quantifying nanoparticle

uptake and biodistibution patterns in the presence and absence of known inhibitors of

endocytic processes.

The uptake of ~ 10 nm fluorescent quantum dots (QDs) was investigated by

measuring the concentration of QDs following exposure to bovine respiratory and

olfactory mucosal explants. An inductively coupled optical emission spectroscopy

method was developed to measure the amount of QDs within the tissues. The results

demonstrated that carboxylate-modified QDs (COOH-QDs) show ~2.5 fold greater

accumulation in the epithelial and submucosal regions of the olfactory tissues compared

to the respiratory tissues. Endocytic inhibitory studies showed that in respiratory tissues

clathrin-dependent, macropinocytosis and caveolae-dependent endocytosis process were

all involved in the uptake of COOH-QDs. Whereas in olfactory tissues, clathrin-

dependent endocytosis was the major endocytic pathway involved in uptake of COOH-

QDs. Additional energy-independent pathways appeared to also be active in the transfer

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of COOH-QDs into the olfactory mucosa. Interestingly, PEGylated quantum dots (PEG-

QDs) of similar size ~15 nm were not internalized into the bovine nasal tissues.

In vivo fluorescence imaging was used to study the biodistribution of quantum

dots following nasal instillation in mice. These studies showed that majority of COOH-

QDs remain in the nasal tissues for relatively long periods of time (up to 24 h) whereas

PEG-QDs showed no such accumulation. Biodistribution studies of gold nanoparticles

(~15 nm) in mice using micro-CT showed that gold nanoparticles were transferred to the

posterior turbinate region and a fraction of the administered dose distributed to regions in

close proximity to the olfactory bulb. Both NIR imaging and micro-CT imaging were

useful tools for visualization of in vivo nanoparticle distribution.

A diazepam-containing microemulsion (dispersed phase ~40 nm) was formulated

to investigate the uptake mechanisms utilized for fluid-phase colloidal dispersions in the

nasal mucosa. The resulting diazepam-containing microemulsion showed enhanced

transfer of the drug into the bovine nasal respiratory and olfactory tissues. It is unclear if

endocytosis of the fluid-phase nanodispersions played a role in drug absorption from the

microemulsions in a manner similar to the uptake of solid-phase nanoparticles, however,

since there was significant loss of the epithelial cell layer following exposure to the

microemulsion formulation which likely altered the barrier properties of the epithelium.

These studies have increased the fundamental understanding of ultrafine

nanoparticle uptake in the nasal tissues and the resulting nanoparticle biodistribution

patterns. While ultrafine nanoparticles may have limited application in the development

of efficient drug delivery systems, an understanding of the size-dependent and tissue-

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dependent processes responsible for the uptake of particulates into mucosal tissues will

contribute to the rational development of nanoparticulate drug delivery strategies

investigating the nasal and other routes of administration.

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PUBLIC ABSTRACT

A variety of ultrafine nanomaterials including metals, engineered nanoparticles,

and viruses have been reported to be transported from the nasal tissues into either the

brain or the blood. The mechanisms involved in the uptake of these nanomaterials,

especially those in the ultrafine size range (diameter < 20 nm), through nasal mucosa and

their subsequent biodistribution in the body are not well characterized. The objective of

this study is to investigate the mechanisms involved in the uptake of quantum dots (a

model for ultrafine nanoparticles) into the nasal tissues and to characterize their

biodistribution patterns using mice as an animal model

The uptake of these ultrafine nanoparticles was observed to depend on their

surface characteristics; negatively charged, carboxylate quantum dots were shown to be

taken up by the nasal tissues to a greater extent (2-5 % in 120 min) than with a near-

neutral surface charge PEGylated QDs which showed negligible uptake into the tissues.

The ultrafine nanoparticles were internalized into the nasal tissues using multiple

pathways including micropinocytosis, clathrin-mediated and caveolae-mediated

endocytosis. Additional energy-independent pathways were involved in the uptake into

olfactory tissues. Following intranasal administration in mice, ultrafine gold

nanoparticles and carboxylate quantum dots were observed to migrate from the anterior

region of the nasal cavity to posterior regions, including near the olfactory regions, due to

mucociliary clearance and some particles were appeared to be retained in the posterior

regions for periods of time at least up to 24 h.

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While ultrafine nanoparticles may have limited application in the development of

efficient drug delivery systems, due to limitations of low drug loading efficiency and

potential toxic effects, these results contribute to the rational development of

nanoparticulate drug delivery strategies investigating the nasal and other routes of

administration.

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TABLE OF CONTENTS

LIST OF TABLES ........................................................................................................... xiii

LIST OF FIGURES ...........................................................................................................xv

CHAPTER I .........................................................................................................................1

INTRODUCTION ...............................................................................................................1 Nasal Anatomy and Physiology........................................................................3 The Respiratory Region ....................................................................................4 The Olfactory Region .......................................................................................7 Intranasal Delivery of Drugs and Vaccines ......................................................8 Fate of Inhaled Nanomaterials Deposited on the Nasal Mucosa ......................9 Nanomaterial Internalization Pathways Into and Across the Nasal Tissues ............................................................................................................10

Mechanisms of Intercellular Transfer of Nanomaterial ..........................11 Mechanisms of Intracellular Transfer of Nanomaterial ..........................12

In Vitro and In Vivo Intranasal Uptake Models .............................................16 Solid-phase Nanodispersions in Intranasal Uptake ........................................16

Polymeric Nanoparticles .........................................................................17 Nonbiodegradable Polymeric Nanoparticles ...........................................17 Biodegradable Polymeric Nanoparticles .................................................20 Metallic Nanoparticles .............................................................................23 Solid Lipid Nanoparticles ........................................................................28

Liquid-phase Nanopdispersions in Intranasal Drug Delivery ........................29 Liposomes ................................................................................................31 Microemulsions .......................................................................................33

Summary .........................................................................................................37

CHAPTER II ......................................................................................................................39

OBJECTIVES ....................................................................................................................39

CHAPTER III ....................................................................................................................41

UPTAKE AND TRANSPORT PATHWAYS FOR ULTRAFINE NANOPARTICLES (QUANTUM DOTS) IN THE NASAL MUCOSA .....41 Introduction .....................................................................................................41 Materials and Instrumentation ........................................................................44 Experimental Procedures ................................................................................45

Preparation of Quantum Dot Dispersions ................................................45 Determination of Particle Size and Zeta Potential ..................................45 Preparation of Bovine Respiratory and Olfactory Mucosal Tissues .......45 Quantum Dot Uptake Studies ..................................................................46 Quantification of Quantum Dots .............................................................47 Extraction of Cadmium from QDs ..........................................................50 Extraction of Cadmium from QDs in Bovine Respiratory and Olfactory Tissues .....................................................................................51 Visualization of QDs in Tissues Using Confocal Microscopy ................52

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Visualization of QDs in Tissues Using Transmission Electron Microscopy (TEM) ..................................................................................53 Investigation of the Endocytic Pathways Involved in the Uptake of Quantum Dots ..........................................................................................54 Statistical Analysis ..................................................................................55

Results.............................................................................................................55 Particle Size Analysis ..............................................................................55 Quantification of Quantum Dots .............................................................56 QD Translocation into Nasal Respiratory and Olfactory Mucosa ...........60 Visualization of QDs in Tissues Using Confocal and Electron Microscopy ..............................................................................................65 Transmission Electron Microscopy (TEM) .............................................68 Identification of Endocytic Pathways ......................................................71

Discussion .......................................................................................................74 Conclusion ......................................................................................................79

CHAPTER IV ....................................................................................................................80

DISTRIBUTION OF QUANTUM DOTS AFTER INTRANASAL ADMINISTRATION IN MICE IN VIVO LIVE ANIMAL IMAGING .......80 Introduction .....................................................................................................80

In Vivo Fluorescence Imaging ................................................................82 Quantum Dots in Small Animal Fluorescence Imaging ..........................83 Micro-Computed Tomography (Micro-CT) Small Animal Imaging ......84

Materials and Instrumentation ........................................................................85 Animals ....................................................................................................85 Administration of Quantum Dots ............................................................86 In Vivo Fluorescence Imaging ................................................................87 Image Analysis ........................................................................................89 Distribution of Gold Nanoparticles: Micro-CT Imaging .........................89

Results and Discussion ...................................................................................91 IRDye® Distribution Following Intranasal Administration .....................91 Distribution of COOH-QDs Following Intranasal Administration .........94 Effect of Particle Surface Modifications on Intranasal Uptake .............104 Mechanistic Evaluation of COOH-QD Uptake from Nasal Tissues Using Whole Animal Imaging ...............................................................110 Translocation of Gold Nanoparticles Measured Using Micro-CT ........115

Conclusions...................................................................................................121

CHAPTER V ...................................................................................................................123

UPTAKE OF MICROEMULSIONS FROM NASAL TISSUES ...................................123 Introduction ...................................................................................................123 Materials and Methods .................................................................................126

Solubility Studies ...................................................................................126 Microemulsion Formulation ..................................................................127 Characterization of Microemulsions .....................................................130 Preparation of Bovine Nasal Tissues .....................................................130 Uptake Studies of Diazepam-containing Microemulsions ....................130 Histological Evaluation of Bovine Nasal Tissues Exposed to Microemulsions .....................................................................................133

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Transport Inhibitor Studies ....................................................................134 HPLC Analysis ......................................................................................134 Statistical Analysis ................................................................................135

Results and Discussion .................................................................................136 Solubility Studies ...................................................................................136 Characterization of Microemulsions .....................................................136 Microemulsion Uptake Studies .............................................................139 Histological Evaluation .........................................................................143 Mechanism of DZME Uptake into Nasal Tissues .................................144

Conclusions...................................................................................................147

CHAPTER VI ..................................................................................................................148

CONCLUSIONS..............................................................................................................148

REFERENCES ................................................................................................................151

APPENDIX-A..................................................................................................................169 Technical Specifications and Optical Spectrum of COOH-QDs ..................169

APPENDIX-B ..................................................................................................................170 Particle Size Distribution of 0.05 mg/mL COOH-QD and PEG-QD Dispersion in KRB ........................................................................................170

APPENDIX-C ..................................................................................................................172 Data Showing Efficiency of ICP-OES Measurements in Presence of Blank Tissues ................................................................................................172

APPENDIX-D..................................................................................................................174 Sample Calculation of Mass of QD from Measured Cd Concentration .......174

APPENDIX-E ..................................................................................................................175 TEER Measurements Representing Respiratory and Olfactory Tissue Integrity .........................................................................................................175

APPENDIX-F ..................................................................................................................176 Sequential Fluorescence Images from Individual Mice after Intranasal Administration of COOH-QDs .....................................................................176 Sequential Fluorescence Images from Individual Mice after Intravenous Administration of COOH-QDs .....................................................................178 Sequential Fluorescence Images of Individual Mice after Intranasal Administration of PEG-QDs .........................................................................180 Sequential Fluorescence Images of Individual Mice after Intranasal Administration of COOH-QDs in the Presence of an Endocytic Inhibitor Cocktail ..........................................................................................182

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LIST OF TABLES

Table 1-1 Summary of studies showing intranasal uptake of polystyrene nanoparticles. ............................................................................................................18

Table 1-2 Summary of selected studies showing potential use of biodegradable PLGA and PLA particles in delivering a variety of drugs and vaccines via the intranasal route. .........................................................................................................22

Table 1-3 Summary of studies showing translocation of metallic nanoparticles following intranasal administration. .........................................................................25

Table 1-4 Summary of selected studies showing toxicity effects of metallic nanoparticles following intranasal instillation. .........................................................27

Table 1-5 Summary of studies investigating intranasal uptake of drugs/antigens using solid lipid nanoparticle systems. .....................................................................30

Table 1-6 Summary of studies showing intranasal uptake of drugs/vaccines using liposomal vesicular systems......................................................................................32

Table 1-7 Summary of studies showing intranasal uptake of drugs/vaccines using microemulsion systems. ............................................................................................36

Table 3-1 Quantum dot (~ 7nm) particle size distribution (n=3, mean ± standard deviation) and surface charge for 0.05 mg/mL samples in KRB..............................56

Table 3-2 Operating conditions and measurement parameters of Varian ICP-OES 720 ES. ......................................................................................................................57

Table 3-3 Measurement of the percent transport (relative to the donor QD loading) of quantum dots across bovine respiratory and olfactory mucosal explants. Experiments were initiated by placing 1 mL of QD dispersion containing approximately 44.5 ± 3.9 µg of QD in the chamber facing the mucosal surface of the tissue. Incubations of 30, 60 and 120 min were conducted and 3 tissues were evaluated at every time period. Recovery of QDs in donor chamber, receiver chamber, mucosal tissue, and from tissue washings following the transport studies are provided as percentage of the initial dose. The values are given as mean (n=3) ± (standard deviation). ....................................63

Table 3-4 Measurement of the transport of quantum dots across bovine respiratory and olfactory mucosal explants. Experiments were initiated by placing 1 mL of QD dispersion containing approximately 44.5 ± 3.9 µg of QD in the chamber facing the mucosal surface of the tissue. Incubations of 30, 60 and 120 min were conducted and 3 tissues were evaluated at every time period. Recovery of QDs in donor chamber, receiver chamber, mucosal tissue, and from tissue washings following the transport studies are provided in the table. The values are given as mean (n=3) ± (standard deviation). ....................................64

Table 5-1 Composition of microemulsions with and without drug and 2,4-DNP. ..........128

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Table 5-2 Size and zeta potential of microemulsions with and without drug and 2,4-DNP (mean ± std dev). ...........................................................................................138

Table A-1 Specifications of COOH-QDs provided by NN-Labs, LLC (Fayetteville, AR). Catalog # CZW-R. .........................................................................................169

Table C-1 Correlation between theoretical mass of Cd as added QDs to blank olfactory tissues and the Cd concentration measured from the digested samples of those tissues. QD dispersion (0.2 mL) spiked into known olfactory tissue weight and digested in 1 mL of nitric acid followed by dilution to 10 mL with DI water. ............................................................................172

Table C-2 Correlation between theoretical mass of Cd as added QDs to blank respiratory tissues and the Cd concentration measured from the digested samples of those tissues. QD dispersion (0.2 mL) spiked into known respiratory tissue weight and digested in 1 mL of nitric acid followed by dilution to 10 mL with DI water. ............................................................................173

Table E-1 TEER values across respiratory and olfactory tissues exposed to COOH-QD dispersion measured at the beginning and end of the transport study for each time point. Values represent maintenance of tissue integrity before and after the transport. ...................................................................................................175

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LIST OF FIGURES

Figure 1.1 Schematic representation of nasal anatomy showing the location of turbinate regions, olfactory mucosa and olfactory bulb. Reproduced with permission23. ...............................................................................................................4

Figure 1.2 Schematic representation of the composition of the respiratory epithelium and submucosal region27. The epithelial barrier of respiratory mucosa typically consists of ciliated pseudostratified columnar epithelial cells, goblet cells and basal cells. The underlying lamina propria consists of connective tissue (CT), fibroblasts, blood vessels (BV), and serous glands. .............6

Figure 1.3 Schematic representations of the location of the olfactory region, the olfactory epithelium and olfactory neuronal pathways from nose to olfactory bulb28. Reproduced with permission. ..........................................................................7

Figure 1.4 Possible pathways of nanomaterial transfer from nose to brain and systemic circulation. Inhaled nanoparticles have a chance to be taken up by either respiratory or olfactory mucosa and subsequently enter the brain and/or the systemic circulation. Reproduced with permission38. .........................................10

Figure 1.5 Schematic representations of internalization pathways of nanomaterial (left). Schematic representations macropinocytosis, clathrin-mediated endocytosis and caveolae-mediated endocytosis pathways (right). Reproduced with permission39. .....................................................................................................11

Figure 3.1 Schematic showing the working principle of ICP-OES135. Metal-containing samples are nebulized using a peristaltic pump and an auxiliary argon flow directly into the plasma. The metal-containing droplets are atomized, ionized and finally exited to higher energy levels. The characteristic emissions from metal ions are separated using a high precision prism and a photomultiplier tube detector captures the intensity of each emission. ...................................................................................................................49

Figure 3.2 Sample calibration curve for elemental Cd using ICP-OES (n=3). Calibration equation was Intensity (a.u) = 3.4984 * Cd Conc. (ng/mL), r2 = 0.9999. ......................................................................................................................57

Figure 3.3 Correlation between theoretical QD concentrations versus ICP-OES measured QD concentration. A correlation equation of Intensity (a.u) = 0.9697 * Cd. Conc (ng/mL) was observed (r2 =0.9992). ..........................................58

Figure 3.4 Correlation of the theoretical mass of Cd as added QD dispersion to blank respiratory tissues and the Cd concentration measured from digested samples of these tissues. A good correlation between added mass of QDs and measured mass of QDs was observed (y=0.997x, r2=0.999). ...................................59

Figure 3.5 Correlation of the theoretical mass of Cd as added QDs to blank olfactory tissues and the Cd concentration measured from digested samples of

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these tissues. A good correlation between added mass of QDs and measured mass of QDs was observed (y=996x, r2=0.999). ......................................................60

Figure 3.6 Comparison of QD uptake into full thickness olfactory and respiratory tissues after a 120 min incubation period. A) Column graph showing the mean and standard deviation of the groups Uncorrected Fischer’s LSD test showed significant difference (p<0.05) in uptake of QD between respiratory and olfactory tissues after 120 min incubation B) Box Whisker plots of the same data showing the median and range of the data. (n=3). ...................................61

Figure 3.7 Confocal laser scanning microscopic images of respiratory tissues showing the transport of QDs. Column I shows the nuclear stain (DAPI) channel. The epithelial region is indicated by a solid line and the submucosal region by a double arrowed line. Column II shows the QD channel, and column III shows merged images from both channels. Each row of images is labeled with the exposure time of the respiratory tissues to QDs or to a control samples with no QD exposure. White arrows highlight the QD localization in the merged image of the respiratory tissue. (Scale bar = 20 µm). ...........................................................................................................................66

Figure 3.8 Confocal laser scanning microscopic images of olfactory tissues showing the transport of QDs. Column I shows the nuclear stain (DAPI) channel. The epithelial region is indicated by a solid line and the submucosal region by a double arrowed line. Column II shows the QD channel, and column III shows merged images from both channels. Each row of images is labeled with the exposure time of the olfactory tissues to QDs or to a control samples with no QD exposure. White arrows highlight the QD localization in the merged image of the olfactory tissue. (Scale bar = 20 µm). ...............................67

Figure 3.9 Transmission electron micrographs of bovine respiratory epithelial cells exposed to COOH-QDs for 120 min. Distinct nucleus (N), mitochondria (M), cellular junction (CJ), golgi apparatus (G) can be observed. a: magnification x7000. b: Enlarged mucosal region (orange circle) showing dispersed, electron-dense particles in cytoplasm (green circles) and in vesicle structures (red circles) of the epithelial region (magnification x21000). ..................................69

Figure 3.10 Transmission electron micrographs of bovine olfactory epithelial cells exposed toCOOH-QDs for 120 min. Distinct nucleus (N), mitochondria (M), cellular junction (CJ) can be observed. a: Image with magnification of x7000. b: Enlarged mucosal region (orange box) showing dispersed, electron-dense particles in cytoplasm (green circles) and vesicular structures (red circles) in the epithelial region(magnification: x24000)...........................................70

Figure 3.11 Uptake of QDs in the nasal respiratory tissue in the presence of inhibitors: 2,4-dinitrophenol (DNP), amiloride, methyl-β-cyclodextrin (MBC) and chlorpromazine (CPZ). A) Bar graph showing mean and standard deviation, * indicates significant difference between the control and inhibited uptake compared used Student’s t-test, n=3 or 6, p<0.05. B) Box Whisker plot showing the median and range of the same data. ................................72

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Figure 3.12 Uptake of QDs in the nasal olfactory tissue in the presence of inhibitors: 2,4-dinitrophenol (DNP), amiloride, methyl-β-cyclodextrin (MBC) and chlorpromazine (CPZ).A) Bar graph showing mean and standard deviation, * indicates significant difference between the control and inhibited uptake compared used Student’s t-test, n=3 or 6, p<0.05. B) Box Whisker plot showing the median and range of the same data. ................................73

Figure 4.1 Anatomical planes of mouse placed in prone position, showing transverse, sagittal and frontal planes169. Reproduced with permission. ..................91

Figure 4.2 Structure of IRDye 800CW, chemical formulation: C46H50N2Na4O15S4, molecular weight of 1091.1 g/mol173. .......................................................................92

Figure 4.3 Co-registered fluorescence and x-ray whole animal images of a representative mouse showing biodistribution of IRDye after intranasal administration. Green color represents the pseudo-colored NIR emission signal from the IRDye. Presence of IRDye in the nasal region can be observed in both dorsal and side views up to 4 h and only in the dorsal view at 24 h. In the 2 h side view image, a strong presence of dye in the throat region can be observed. The majority of the dye seemed to reside in the abdominal region and was likely associated with the digestive and urinary systems. .....................................................................................................................93

Figure 4.4 Composite fluorescence images co-registered with x-ray images of mice showing the distribution of COOH-QDs after intranasal (top row) and intravenous (middle row) administration. The control group received normal saline is shown in the bottom row. The red color represents the fluorescence signal from COOH-QDs. A gradual decrease of fluorescence intensity in the nasal region (yellow circle in top row image) from 5 min to 24 h can be observed after intranasal dosing, whereas intravenous dosing resulted in high fluorescence intensities in the abdominal region within 2 h. Gradual decreasing intensities can be seen up to 24 h. Sequential images for an individual mouse are provided in Appendix-F. ........................................................95

Figure 4.5 Mean fluorescence intensity from nasal regions (55 mm2 ROI; circled region shown in insert) of animals administered COOH-QDs via intranasal (i.n.), or intravenous (i.v.) routes. * represents statistical significance between i.n. and i.v. fluorescence intensities when tested with Student’s t-test at p<0.05 (n=3). ............................................................................................................96

Figure 4.6 Fluorescence images co-registered with x-ray images of mice 24 h after COOH-QD administration. Upper panel shows images of anesthetized, live mice with intact nasal cavity and lower panel shows images of euthanized mice with exposed nasal cavity. Opening of the nasal cavity enabled visualization of the strong fluorescence signal from COOH-QD accumulation in the deeper nasal tissues following intranasal administration that was not visible in the mice with intact nasal cavities. Intravenous administration did not show nasal tissue accumulation, even in the exposed nasal cavity images. Images of all mice after opening nasal cavity are provided in Appendix-F. ............98

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Figure 4.7 Mean fluorescence intensities of COOH-QDs from the nasal regions of mice with intact nasal cavities and exposed nasal cavities after 5 min and 24 h following intranasal (30 ug/animal) and intravenous (60 ug/animal) administration. * represents statistical significance between i.n. and i.v. fluorescence intensities when tested using Student’s t-test at p < 0.05 (n=3). .........99

Figure 4.8 Fluorescence images co-registered with X-ray images from various harvested organs of mice 24 h following intranasal administration (A) and intravenous administration (B). ..............................................................................100

Figure 4.9 Mean fluorescence intensities from various organs of mice 24 h after intranasal (30 ug/animal) and intravenous administration (60 ug/animal) of QDs.* represents statistical significance between i.n. and i.v. fluorescence intensities when tested using Student’s t-test at p < 0.05 (n=3). ............................101

Figure 4.10 Composite fluorescence images of mice co-registered with corresponding x-ray images. These images compare the distribution of (upper panel) PEG-QDs (30 μg/animal in 5 μL volume) and (lower panel) COOH-QDs(30 μg/animal in 5 μL volume) at various time points after intranasal administration. Sequential images for all individual mice are provided in Appendix-F. ............................................................................................................105

Figure 4.11 Comparison of mean fluorescence intensities from the intact nasal region of mice (from a 55 mm2 area ROI as shown in Figure 4.4) following intranasal administration of COOH-QDs and PEG-QDs at same dose of 30 μg/animal in 5 μL volume. (n=3). ...........................................................................106

Figure 4.12 Fluorescence images of mice co-registered with corresponding x-ray images 24 h after intranasal administration of PEG-QDS and COOH-QDs. The upper panels are images of mice with intact nasal cavities. Lower panels are images of mice following the opening of the nasal cavity along the septum prior to imaging. Images of all mice after opening nasal cavity are provided in Appendix-F. ............................................................................................................107

Figure 4.13 Mean fluorescence intensities from the deeper nasal tissues of mice with intact and exposed nasal cavities 24 h after intranasal administration of PEG-QDs and COOH-QDs, at same dose of 30 μg/animal * represents statistical significance between COOH-QDs and PEG-QDs fluorescence intensities tested using Student’s t-test at p<0.05 (n=3). ........................................108

Figure 4.14 Whole animal fluorescence images of mice co-registered with corresponding x-ray images comparing the distribution of COOH-QDs (top row) in the presence of a cocktail of endocytic inhibitors (bottom row) at various time points after intranasal administration. The red fluorescence in the nose region represents signal from QDs. Inclusion of inhibitors resulted in no difference in QD fluorescence signal when compared to QDs without inhibitors. Sequential images for all individual mice are provided in Appendix-F. ............................................................................................................111

Figure 4.15 Mean fluorescence intensities from nasal regions (from a 55 mm2 area ROI as shown in Figure 4.4) of live animals in presence and absence of

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inhibitor cocktail. No difference in fluorescence signal from QDs in presence of inhibitor cocktail is observed. (n=3). ..................................................................112

Figure 4.16 Images of mice showing the fluorescence from the deeper nasal regions 24 h after intranasal administration of COOH-QDs in the presence and absence of an endocytic inhibitor cocktail. The upper panels are images of mice with intact nasal cavities. Lower panels are images of mice following the opening of the nasal cavity along the septum prior to imaging. Images of all mice after opening nasal cavity are provided in Appendix-F. ...........................113

Figure 4.17 Mean fluorescence intensities from the nasal tissues of mice with intact and exposed nasal cavities 24 h after intranasal administration of COOH-QDs in presence and absence of an endocytic inhibitor cocktail (n=3). .........................114

Figure 4.18 In vivo micro-CT imaging of a mouse head region 24 h after intranasal (A) and intravenous (B) administration of 15 nm gold nanoparticles. Axial (left column), dorsal (middle column) and lateral views (right column) are depicted. Yellow colored arrows show the accumulation of gold particles in the nasal conche following intranasal administration, whereas no such accumulation was observed in the same regions after intravenous administration. ........................................................................................................116

Figure 4.19 In vivo micro-CT 3D lateral view of the head regions of a mouse showing the accumulation of AuNPs in anterior and posterior regions of the nasal cavity (yellow arrows) 24 h after intranasal administration. .........................117

Figure 4.20 In vivo micro-CT images of mouse in axial (left), dorsal (middle) and lateral (right) views 24 h after intravenous administration of AuNPs. High contrast CT signal in heart (yellow arrow), liver (purple arrow) and spleen (blue arrow) can be observed. .................................................................................118

Figure 4.21 In vivo micro-CT images of the head region of a mouse 6 h (A) and 24 (B) after multiple dosing of AuNPs via the intranasal route. Accumulation of AuNPs (colored in red) in the nasal mucosa can be observed. ...............................120

Figure 5.1 Chemical structure of diazepam203. ................................................................124

Figure 5.2 Schematic showing the preparation of diazepam-containing microemulsion. Microemulsions were prepared by adding the required amount of a sorbitol solution of known concentration to a mixture containing IPM, Tween 80, and diazepam in a stirred beaker at 55 °C followed by cooling to room temperature. ..................................................................................129

Figure 5.3 Sample calibration curve for analyzing diazepam using HPLC method (n=3). Good linearity of the method was observed from 1 μg/mL to 1000 μg/mL with AUC=35.913* Concentration (μg/mL) and r2=0.9999. Insert is presented for better visualization of the linearity in low concentrations (0 to 100 μg/mL). ............................................................................................................135

xx

Figure 5.4 Solubility of diazepam in various components used in microemulsions. Column bars are labeled with the mean solubility value (mg/mL). Results are expressed as mean ± standard deviation of three replicates. ..................................136

Figure 5.5 Appearance of diazepam-containing microemulsion. ....................................138

Figure 5.6 Comparison of the cumulative percent of diazepam (relative to the donor chamber initial concentration) appearing in the receiver chamber as a function of time across bovine respiratory and olfactory tissue explants following exposure to DZS (diazepam IPM solution, 4 mg/mL) and DZME (diazepam-containing microemulsion, 4 mg/mL). A) Data shown are mean ± standard deviation (n=6 per tissue type). B) Mean percent diazepam transferred to receiver results shown without error bars. ..............................................................140

Figure 5.7 Comparison of diazepam percent remaining in the donor chamber after 120 min exposure of diazepam IPM solution (DZS) and diazepam-containing microemulsion (DZME) to the respiratory, olfactory and artificial membrane. Dashed line represents 100%. Data shown are mean ± standard deviation (n=6 per tissue/membrane type). ...........................................................................141

Figure 5.8 Comparison of diazepam percent (relative to the initial diazepam concentration in the donor chamber) accumulated in the respiratory and olfactory tissues after 120 min exposure to diazepam-IPM solution (DZS) and diazepam-containing microemulsion (DZME). Data shown are mean ± standard deviation (n=6 for each tissue type). * Indicates a statistically significant difference when compared using an unpaired, two-tailed Student’s t-test with p<0.05. ...................................................................................................141

Figure 5.9 Comparison of diazepam accumulation in the thickness normalized olfactory and respiratory tissue explants after 120 min exposure to a diazepam IPM solution (DZS) and a diazepam-containing microemulsion (DZME). Data shown are mean ± standard deviation (n=6 for each tissue type). .................143

Figure 5.10 Brightfield microscopic images of control (left) and DZME exposed (right) respiratory tissue explant. Solid line arrow shows intact epithelium in control tissue and the dashed-line arrow shows the damaged epithelium in DZME-exposed respiratory tissue. .........................................................................144

Figure 5.11 Brightfield microscopic images of control (left) and DZME exposed (right) olfactory tissue explant. Solid line arrow shows intact epithelium in control tissue and the dashed-line arrow shows the damaged epithelium in DZME-exposed olfactory tissue. ............................................................................144

Figure 5.12 Comparison of diazepam accumulation in the respiratory and olfactory tissues after 120 min exposure to DZME in the presence and absence of the metabolic inhibitor 2,4 –dinitrophenol (2,4-DNP). Data shown are mean ± standard deviation (n=6 for each tissue type). * Indicates a statistically significant difference when compared using an unpaired two-tailed Student’s t-test with p<0.1. .....................................................................................................146

xxi

Figure A.1 Typical UV-Vis absorption (a) and emission (b) spectrum of COOH-QDs purchased from NN-Labs, Inc (Fayetteville, AR), Lot # LW074414A21104. ................................................................................................169

Figure B.1 Representative particle size distribution of 0.05 mg/mL COOH-QDs (Lot# LW074414A21104) in KRB using a volume-weighted distribution analysis (Nicomp Particle Sizer, Model 380 ZLS, Santa Barbara, CA). ................170

Figure B.2 Representative particle size distribution of 0.05 mg/mL PEG-QDs (Lot# LW134414A23108) in KRB using a volume-weighted distribution analysis (Nicomp Particle Sizer, Model 380 ZLS, Santa Barbara, CA). ..............................171

Figure F.1 Fluorescence images co-registered with x-ray images from individual mice showing the distribution of COOH-QDs after intranasal administration. The red color represents the fluorescence signal from COOH-QDs. .....................176

Figure F.2 Fluorescence images co-registered with x-ray images of euthanized individual mice with exposed nasal cavities observed 24 h after intranasal administration of COOH-QDs. The red color represents the fluorescence signal from COOH-QDs. ........................................................................................177

Figure F.3 Fluorescence images co-registered with x-ray images from individual mice showing the distribution of COOH-QDs after intravenous (retro-orbital injection) administration. The red color represents the fluorescence signal from COOH-QDs. Mouse 2 died after the 2 h time point. .....................................178

Figure F.4 Fluorescence images co-registered with x-ray images of euthanized individual mice with exposed nasal cavities observed 24 h after intravenous administration of COOH-QDs. ...............................................................................179

Figure F.5 Fluorescence images co-registered with x-ray images from individual mice showing the distribution of PEG-QDs after intranasal administration. The red color represents the fluorescence signal from PEG-QDs. .........................180

Figure F.6 Fluorescence images co-registered with x-ray images of euthanized individual mice with exposed nasal cavities observed 24 h after intranasal administration of PEG-QDs. ...................................................................................181

Figure F.7 Fluorescence images co-registered with x-ray images of individual mice showing the distribution of COOH-QDs in the presence of an endocytic inhibitor cocktail following intranasal administration. The red color represents the fluorescence signal from COOH-QDs. ............................................182

Figure F.8 Fluorescence images co-registered with x-ray images of euthanized individual mice with exposed nasal cavities observed 24 h after intranasal administration of COOH-QDs in the presence of an endocytic cocktail inhibitor. The red color represents the fluorescence signal from COOH-QDs. .....183

1

CHAPTER I

INTRODUCTION

Nanotechnology is the science and engineering of extremely small colloidal

materials less than 100 nm in size. Materials at such small sizes possess a wide variety of

physical and chemical properties that are different than those same materials at larger

sizes. Due to these unique properties, the applications of nanotechnology have expanded

rapidly into several fields including biomedical and pharmaceutical applications.

Nanotechnology in medicine shows immense potential for improving the lives of humans

by improving disease diagnosis, delivery of therapeutic agents and tissue engineering.

The term ‘colloidal dispersion’ is used to describe a homogenous system in which

particles of size between 1 nm to 1000 nm of any nature (e.g. solid, liquid or gas) are

dispersed in a continuous phase of a different composition (or state)1. Systems that

contain solid colloidal material dispersed in liquid media are termed “solid-phase

nanodispersions” and semi-solid/liquid colloidal materials dispersed in liquid media are

“liquid-phase nanodispersions”. Advancements in nanotechnology have enabled

researchers to explore several varieties of colloidal dispersions to deliver therapeutic

agents to target selected regions in the body. For example, liquid-phase nanodispersions

like liposomes2, niosomes3, micelles4, microemulsions5 and solid-phase nanodisperions

like polymeric nanoparticles6, metallic nanoparticles7, drug nanocrystals8, and polymer

drug conjugates9 have been reported to enhance delivery of therapeutic agents via

common routes of administration including oral, parenteral, transdermal, pulmonary,

ocular and intranasal routes.

2

Some of these colloidal systems have already been approved by the FDA and are

being used clinically in humans. For example, Doxil® injection is a liposomal formulation

of doxorubicin approved by the FDA in 1995 for treatment of ovarian cancer10. Even

though nanotechnology has advanced significantly, there are still only a few approved

drug products using these technologies due to the limited knowledge regarding the

cellular interactions of colloidal systems. A great amount of work is in progress aimed to

understand the interactions of nanomaterials with biological tissues and the mechanisms

of nanoparticle cellular internalization.

Delivery of nanoparticles via the intranasal route has gained the attention of

numerous investigators. Several nanoparticle drug delivery systems have been reported to

increase the transport of drugs/vaccines across the nasal mucosa compared to

drugs/vaccines delivered as solutions11-12. Studies have shown that both engineered

nanoparticles (carbon nanotubes, quantum dots) and natural nanoparticles (smoke, dust,

and viruses) can transfer to both the brain and systemic circulation following intranasal

exposure13-15. Understanding the role of the nasal mucosa in the internalization of the

nanoparticle systems is essential to develop advanced drug delivery systems and improve

delivery of drugs/vaccines via this route.

In addition to a useful route for the delivery of drugs, the nasal route is also a

potential route for exposure of unwanted airborne nanoparticles. As utilization of

nanomaterials has increased, humans are being continuously exposed to airborne

nanoparticles in workplaces, from consumer products and in public places16. Research

has repeatedly shown several harmful effects of some nanomaterials, including

cytotoxicity and oxidative stress17-18. Nanoparticles of various chemistries are highly

3

reactive and can interact with cellular systems to produce reactive oxygen species (ROS),

which, in turn, cause oxidative stress and alter signal transduction mechanisms,

eventually damaging the cells and tissues19.

Prior to understanding the specifics of nanomaterial transit through nasal tissue, it

is essential to understand the anatomy and physiology of human nose along with its

cellular composition and some key endogenous processes. A detailed description of the

anatomy, physiology and function of nasal cavity was published by Mygind and Dahl20,

and a brief brief overview of nasal anatomy and physiology is summarized below.

Nasal Anatomy and Physiology

Main functions of the nose include the sense of olfaction, regulation of

temperature and humidity of inhaled air, and filtering of particulate matter from the

inhaled air. The structure of the human nose is complex. Anatomically, the human nose is

divided into three distinct regions: the anterior region (nasal vestibule), the

middle/turbinate region (respiratory and olfactory region) and the posterior region

(nasopharynx) (Figure 1.1). The nasal septum differentiates the nasal cavity into two

halves. The surface of the nasal vestibule is composed of stratified, squamous and

keratinized epithelial cells along with sebaceous glands21. These cells are very resistant to

dehydration and can withstand noxious environmental substances while preventing their

permeation. The nasal valve is a narrowing of the airway passage; it lies between the

vestibule and the turbinate regions. The nasal valve is anatomically responsible for most

of the resistance to airflow in the respiratory tract. Beyond the nasal valve, the airway

opens into the large main nasal passage or the turbinate region. Turbinates are

cartilaginous protrusions from the lateral surfaces covered by mucosal tissue and are

4

responsible for the airway dynamics due to their baffle like behavior and for the air

conditioning functions of the nasal cavity due to their relatively large surface areas which

allow for rapid heat and water exchange22. Anteriorly the lower two turbinates and the

associated mucosal tissues in the area are referred to as the respiratory region and the

superior turbinate region called the olfactory region.

Figure 1.1 Schematic representation of nasal anatomy showing the location of turbinate regions, olfactory mucosa and olfactory bulb. Reproduced with permission23.

The Respiratory Region

The respiratory region contains the largest surface area of the nasal cavity; the

total surface area of the nasal mucosa is estimated to be 180 cm2 of which respiratory

mucosa occupies ~170 cm2. Morphologically, the respiratory mucosa is composed of two

distinct layers: the respiratory epithelium and the lamina propria (submucosal region).

The respiratory epithelium is composed of ciliated pseudostratified columnar epithelial

cells, goblet cells and the basal cells (Figure 1.2). The epithelial cells are approximately

5

25 μm in height, 7 μm wide at the tip and 3 μm wide at the base. These cells are

connected via tight junctions and possess a barrier function, inhibiting the permeation of

many substances. Each epithelial cell in the respiratory region contains 300 to 400

microvilli that are 0.1 μm in diameter and 2 μm in height on their apical surfaces24.

Goblet cells are also present within the respiratory epithelium and, along with the mucus

glands present in the submucosal region secrete and maintain the mucus layer. Mucin, a

glycoprotein is the main constituent of mucus, and the mucus layer forms a continuous,

10 -20 μm thick blanket covering the epithelial cells. Cilia present on the epithelial cells

continuously beat at a rate of 10 to 20 times per second and are responsible for the

clearance of mucus and trapped particulate matter into the nasopharynx24. Basal cells are

present on the basement membrane separating the epithelium from the submucosal

region; these stem cells can mature into ciliated epithelial cells or goblet cells. The

submucosal region is composed of connective collagen fibrils and is highly vascularized

with an abundant capillary network and contains serous and mucous glands. Materials

reaching the submucosal region have access to the systemic circulation through the

vascular network.

The submucosal region also contains the “ Nasal Associated Lymphoid Tissue

(NALT)” which is responsible for the local mucosal defense system and involves both

the production of antibodies and serves as the source of circulating immune cells25.

NALT is an aggregation of organized lymphoid tissue consisting of aggregates of paired

lymphocyte-like follicles situated in the region closer to the nasopharyngeal duct. These

follicles are reported to contain T- and B-cells, and a layer of specialized membranous

cells called M-cells covers the surfaces of the follicles. Reports have shown that these M-

6

cells function similarly to the M-cells in the Peyer’s Patches present in the

gastrointestinal tract and it is reported that these cells are responsible for nanoparticle

uptake in the nasal mucosa26. Several vaccine delivery systems actively target antigens to

the lymphoid NALT via intranasal route to trigger mucosal immunity12.

Figure 1.2 Schematic representation of the composition of the respiratory epithelium and submucosal region27. The epithelial barrier of respiratory mucosa typically consists of ciliated pseudostratified columnar epithelial cells, goblet cells and basal cells. The underlying lamina propria consists of connective tissue (CT), fibroblasts, blood vessels (BV), and serous glands.

7

Figure 1.3 Schematic representations of the location of the olfactory region, the olfactory epithelium and olfactory neuronal pathways from nose to olfactory bulb28. Reproduced with permission.

The Olfactory Region

Figures 1.3 illustrates the location and cellular composition of the olfactory region

in humans. The olfactory region constitutes only about 5% of the total area of the nasal

cavity, and it is in close proximity to the olfactory bulb in the brain29. Similar to the

respiratory region, the olfactory mucosa can be divided into a mucosal (olfactory

epithelium) and submucosal region. The olfactory epithelium is composed of three types

of cells: olfactory neural receptor cells (olfactory axons), supporting cells (sustentacular

cells) and basal cells. Neural receptor cells are un-myelinated axons that originate at the

olfactory bulb and pass through the cribriform plate and terminate at the apical surface of

8

the olfactory epithelium. Supporting cells are columnar cells that provide mechanical

support to the neuronal cells. The basal cells are capable of differentiating into olfactory

neuronal cells or into sustancular cells. The neuronal cells and sustentacular cells are

connected by tight junctions, which form a physical barrier to material transfer. The

submucosal region is located beneath the epithelium and contains the vascular supply,

Bowman’s glands which secrete mucus, and nasal lymphatic vessels. The lamina propria

of the olfactory region is innervated by a neuronal supply that consists of olfactory axon

bundles, autonomic nerve fibers and the maxillary branch of the trigeminal nerve system.

The olfactory region makes a direct connection to the olfactory bulb of the brain.

Intranasal Delivery of Drugs and Vaccines

Nasal delivery of therapeutics has been conventionally limited to the treatment of

local/topical nasal problems like allergies and rhinitis. However, recently nasal delivery

is being used as a substitute for parenteral and oral delivery or in attempts to target

therapeutic agents to the brain30. The highly vascularized and immunologically

competent nasal mucosa offers several advantages including improved bioavailability,

quicker onset of action and excellent mucosal immune response for the delivery of drugs

and vaccines. Nose-to-brain pathways, potentially bypassing the blood-brain barrier,

present an added advantage for the delivery of therapeutics to the brain via the nasal

route. Despite of these advantages, intranasal delivery is restricted to the delivery of

highly-permeable, potent compounds because the nasal cavity can hold only a limited

volume in dose and mucociliary clearance rapidly removes material from the absorption

site31. Advanced delivery systems may be able to overcome some of these limitations

with innovative formulations such as nanoparticles and microemulsions.

9

Fate of Inhaled Nanomaterials Deposited on the Nasal

Mucosa

Deposition of nanomaterials within the nasal cavity is not very clearly

understood; several reports have described computational fluid dynamic models

developed to understand the deposition of nanomaterials in the nasal cavity32-33.

Deposition was found to depend on several factors including the particle size, density and

shape of the nanoparticles34.

Unlike larger, micron-sized particles, nanoparticles are much smaller in size and

mass, and nanoparticles can enter the main nasal cavity via the inspired airstream,

randomly collide with gas molecules in the airstream and may deposit in the posterior

nasal regions after passing through the nasal valve35-36.

Following nasal deposition, nanoparticles can have access to distant regions in the

body by using mucosal uptake and transfer pathways outlined in Figure 1.4. Previous

reports showed that nanoparticles deposited in the nasal cavity have the potential to

transfer to deeper brain regions by bypassing the blood-brain barrier using the olfactory

neuronal and trigeminal nerve pathways35. Nanoparticles reaching the submucosal

regions of the nasal mucosa may also transfer to the systemic circulation and distribute to

distant tissues in body37. Some nanoparticles can be internalized via the NALT and may

transfer to the lymphatic system. Although there is some evidence showing these

pathways exist, the total quantity of nanomaterials transferred to these sites is limited,

and the factors governing the uptake of nanoparticles into various regions of the nasal

mucosa are still unclear.

10

Figure 1.4 Possible pathways of nanomaterial transfer from nose to brain and systemic circulation. Inhaled nanoparticles have a chance to be taken up by either respiratory or olfactory mucosa and subsequently enter the brain and/or the systemic circulation. Reproduced with permission38.

Nanomaterial Internalization Pathways Into and Across the

Nasal Tissues

Nanomaterials need to traverse the mucus layer and the epithelial barrier to have

access to the blood and/or lymph vessels present in the submucosal layer of the nasal

tissues. The possible transport pathways for nanomaterial internalization across and into

the nasal epithelia are shown in Figure 1.5. Material deposited on the nasal mucosal

surface can be internalized into deeper tissue regions by intracellular/transcellular and

paracellular/intercellular routes.

11

Figure 1.5 Schematic representations of internalization pathways of nanomaterial (left). Schematic representations macropinocytosis, clathrin-mediated endocytosis and caveolae-mediated endocytosis pathways (right). Reproduced with permission39.

Mechanisms of Intercellular Transfer of Nanomaterial

Transfer via the paracellular pathway is a convective process, which occurs

between adjacent epithelial cells through the intercellular spaces and tight junctions.

Hydrophilic materials are preferentially transferred through this route. The diameter of

the tight junctions between the nasal epithelial cells is reported to be 3.9 to 8.4 Å40. It is

unlikely that any nanomaterial greater than this diameter will be able to pass through

these junctions in their closed state. However, there is evidence showing the presence of

nanoparticles of diameter 10 – 200 nm in the paracellular spaces. Huang and Donovan

were able to visually show 10 nm carboxylate polystyrene nanoparticles in the

paracellular spaces of rabbit nasal mucosa41. More recently, Chen reported that 20 nm

carboxylate polystyrene nanoparticles were internalized into bovine nasal tissues through

the paracellular route38. These authors have postulated the possible opening of the tight

Endocytic pathways for transcellular

uptake

12

junctions to enable nanoparticle passage. Involvement of paracellular transport pathways

for nanomaterials is still controversial and needs further investigation. The integrity of

tight junctions can be compromised under some pathological conditions or due to usage

of absorption enhancers, which might potentially open up the intercellular spaces and

lead to passage of extremely small nanoparticles through paracellular routes42.

Mechanisms of Intracellular Transfer of Nanomaterial

Since the efficient transfer of nanomaterials through paracellular routes is

unlikely, the particles must use the transcellular routes to reach the deeper tissue regions.

Nanoparticles can interact with the cell membrane and be internalized into the cell

through endocytic processes. Endocytosis of particulate matter can occur by multiple

mechanisms, broadly divided into two categories: phagocytosis (cell eating) and

pinocytosis (cell drinking). Both of these endogenous processes are essential for the

transfer of biomaterials into and out of cells. Drugs and nanocarriers can also access these

endogenous processes or it may be possible for the nanoparticles to trigger these

endocytic pathways based on their physiochemical and surface properties.

Phagocytosis is the process by which large diameter (>1 μm) particulate matter,

including several types of bacteria, is internalized into cells. Phagocytosis primarily

occurs in specialized cells like macrophages, monocytes, neutrophils and dendritic cells.

The primary result of this process is to present unwanted foreign material to the immune

system. Particulate matter deposited on the nasal mucosa has shown to be internalized by

dendritic cells and subsequently to be presented to the NALT43. As these large materials

enter the body, the receptors present on phagocytic cells recognize foreign particles and

engulf them by wrapping the cell membrane around the particle. The wrapped membrane

13

detaches from the cell membrane and forms vesicles that eventually convert to

phagosomes. The phagosomes are rich in enzymes and have a low pH, which assists in

the degradation of the particulate matter into soluble cellular debris. These digested

materials are released into the cytoplasm and are eventually eliminated from the cell by

either diffusion or exocytosis44. Several particulate vaccine delivery systems target these

phagocytosis mechanisms by using specific ligands/antigens to efficiently deliver vaccine

into the immune cells12.

Pinocytosis is the process of internalization of solutes and colloidal substances

(1 nm to 1000 nm) into the cells39. Pinocytosis is active in many types of epithelial cells,

including nasal epithelial cells45. There are different mechanisms that result in

pinocytosis. Broadly pinocytosis can be subdivided into three main morphologically

distinct processes based on the proteins involved: 1) clathrin-dependent endocytosis, 2)

caveolae-dependent endocytosis and 3) macropinocytosis. More detailed descriptions of

these pathways can be accessed elsewhere in a series of publications45-47. Figure 1.5

shows a schematic of events that take place in these three endocytic processes and a brief

description of these processes is presented below.

Macropinocytosis is a non-specific endocytic process which involves the

formation of actin-driven plasma membrane protrusions that collapse onto and fuse with

the plasma membrane and generates large endocytic vesicles (1-5 μm diameter) called

macropinosomes39. Owing to the large size of macropinosomes, this process provides

cells with a way to non-selectively internalize large quantities of external cell milieu.

Unlike other endocytic vesicles, macropinosomes that are larger in size, do not contain

any specific coating, and are believed to be relatively more fluid. Escape of the

14

internalized cargo from macropinosomes is also reported due to its more fluid nature

compared to other endocytic vesicles. Once formed, macropinosomes undergo maturation

and fuse with a lysosome which possess an enzyme-rich acidic environment or recycle

their contents to the surface of the cell. Pharmacologically active drugs, including

amiloride and its derivatives have been shown to inhibit the Na+/H+ exchange pump in

the plasma membrane, which leads to low pH in the cytosol48. Formation of membrane

ruffling was reported to be inhibited at low pH and thus at appropriate concentrations of

amiloride or its analogs inhibit macropinocytosis.

Clathrin-mediated endocytosis is an endogenous process involved in all

mammalian cells for the uptake of essential nutrients, including cholesterol-laden low-

density lipoprotein particles that bind to the LDL receptor and iron-laden transferrin that

binds to transferrin receptors39. This process typically occurs in a membrane region

enriched in clathrin, a main cytosolic coat protein. Clathrin polymerization is responsible

for the formation of the ‘coated pits’ on the cell membrane upon activation of specific

receptors. After formation, coated pits pinch off from the cell membrane with the help of

dynamin and form clathrin-coated vesicles. The resulting endocytic vesicles are relatively

small in diameter at 100-150 nm39. As these vesicles mature to form early endosomes,

they lose their clathrin coat and any associated ligands into the cytosol that may recycle

back these substances to the cell membrane. The early endosomes travel further into the

cytosol and form late endosomes which finally fuse to lysosomes, an acidic and enzyme-

rich environment able to degrade entrapped material. In some cases, the inert entrapped

material, resistant to degradation, may release into the cytosol and return to the

membrane from which it was internalized, or it could traverse the cell and be delivered to

15

an opposite/adjacent cell in a process called transcytosis39. Several compounds have been

reported to dissociate the clathrin lattice and thereby inhibit clathrin-mediated

endocytosis. Chlorpromazine is a commonly used inhibitor for clathrin-mediated

endocytosis; it is believed to inhibit clathrin-coated pit formation49.

Caveolae-mediated endocytosis is also a pinocytosis process involved in

internalization of biomolecules including cholesterol into cells. Caveolae are flask-shaped

invaginations of the plasma membrane rich in cholesterol and sphingolipids, abundantly

present on endothelial cells46. The sizes of caveolae are reported to be 50-60 nm and the

shape and structural organization of caveolae are conferred by caveolin, a dimeric protein

that binds cholesterol50. Similar to clathrin-mediated endocytosis, caveolae-mediated

processes is also highly regulated process involving complex signaling, which may be

driven by the entrapped material itself. Materials on the cell surface move along the

plasma membrane to reach caveolae invaginations, where they may be tethered through

receptor-ligand interactions. Caveosomes are then sorted to cell organelles including the

endoplasmic reticulum. This process bypasses lysosomal formation and is believed to be

a useful pathway for internalization of material that are sensitive to degradation in acidic,

enzyme-rich lysosomes39. Compounds that inhibit the formation of caveolae either by

inhibiting cholesterol binding or interrupting the actin cytoskeleton have been shown to

inhibit caveolae-mediated endocytosis. Methyl-β-cyclodextrin binds to cholesterol in the

caveolae and thus depletes the availability of cholesterol for formation of caveolae, which

results in the inhibition of this endocytic process50.

Materials present on the cell surface, including nanoparticles are believed to be

internalized into the cells through the above described pinocytosis pathways. One

16

common feature among these endocytic processes is formation of vesicles that are finally

presented to the cytosol. Nanoparticles internalized into the cells through these pathways

are either degraded in lysosomes or presented to cell organelles like the Golgi apparatus,

endoplasmic reticulum, or mitochondria39.

In Vitro and In Vivo Intranasal Uptake Models

Studying the intranasal uptake of nanoparticles requires a reliable nasal model. In

vivo models including mouse51, rat37, rabbit41, and sheep52 are being used to study the

biodistribution and bioavailability of therapeutic agents administered in nanoparticle

systems. While these in vivo models are helpful in mimicking the human nasal

conditions, mechanistic studies of nanoparticle transfer into the nasal tissues is difficult

using in vivo models. In vitro studies using excised nasal mucosa from either bovine53,

ovine and porcine40 species have been demonstrated to be efficient and easier models for

understanding the underlying mechanisms of intranasal transit of nanoparticles. In

addition, investigators have used cell culture systems of nasal epithelial airways

developed from a variety of species including bovine, hamster, humans as in vitro models

for studying nanoparticle uptake54.

Solid-phase Nanodispersions in Intranasal Uptake

The term “solid-phase nanodispersions” is used to describe colloidal dispersions

of solid-state nanoparticulate materials of diameter between 1-1000 nm dispersed in a

continuous liquid medium. Nanoparticles are made from a wide range of materials and

can be used as drug carriers or as vaccines, in which therapeutic agents are entrapped,

dissolved, encapsulated, adsorbed or chemically attached. The extent of uptake and

cellular internalization pathways of nanoparticles from the nasal mucosa is dependent on

17

several factors, including size, shape, surface chemistry, and composition of the

nanoparticles. A brief review of different solid nanoparticle systems used in intranasal

delivery is provided below.

Polymeric Nanoparticles

Nonbiodegradable Polymeric Nanoparticles

Initial use of polymeric nanoparticles for drug delivery was limited to

nonbiodegradable and/or non-biocompatible systems like polystyrene, poly (methyl

methacrylate) (PMMA), polyacrylamide and polyacrylates12, 55. While most of these

polymer systems serve as excellent adjuvants for vaccines or carriers for small and large

drug molecules, the accumulation of these particulate systems in tissues raises

biocompatibility concerns and lead to a major concern requiring clearance of these

particulate systems from the body and potential safety concerns. Nonbiodegradable

polyermic systems act as good models for the investigation of the processes used for

nanoparticle uptake into the nasal epithelium. Among the nonbiodegradable

nanoparticles, polystyrene nanoparticles are widely used model nanoparticles for

intranasal studies because they are readily available in different sizes with different

surface chemistries and contain fluorescent dyes enabling easy detection. Table 1-1

summarizes some of polystyrene nanoparticulate systems previously used in the

investigation of intranasal nanoparticle transfer.

18

Table 1-1 Summary of studies showing intranasal uptake of polystyrene nanoparticles.

Surface Modification

Particle size (nm)

Experimental model

Findings Reference

Carboxylate-modified

20 and 100

Excised bovine nasal

mucosa

Smaller nanoparticles (20 nm) showed greater uptake compared to larger nanoparticles (100 nm)

Clathrin-mediated endocytosis in 20 nm nanoparticle uptake in respiratory tissue and possible paracellular transit in olfactory tissues

100 nm particles were internalized only by macropinocytosis in olfactory tissue whereas macropinocytosis and caveolae-mediated endocytosis were involved in uptake by respiratory tissue

Chen et al.38


40 Excised

bovine nasal mucosa

Olfactory tissues showed ~2-fold higher uptake compared to respiratory tissues

Macropinocytosis, clathrin-mediated and caveolae-mediated endocytosis pathways were involved in uptake into both respiratory and olfactory tissue

Al Khafaji et

al.56

125I radiolabeled sulphate-modified

20, 100, 500 and

1000

In vivo rat model

20 nm (3.25% of dose) particles entered systemic circulation to a greater extent than 100 (0.3% of dose), 500 (0.15% of dose), or 1000 nm (0.15% of dose) particles

Brooking et al.37


10

Excised rabbit nasal

mucosa

Cumulative amounts of nanoparticles, irrespective of size and surface characteristics were similar

10 nm carboxylated particles were transported mainly via paracellular routes whereas other larger carboxyl particles (100 nm and 500 nm) crossed membrane via transcellular routes

200 nm amine particles were observed to cross via paracellular and transcellular routes

Huang et al.41

Carboxyl-modified 100 and

500

Amine-modified 200

Polysorbate, and chitosan coating

100 and 200

In vivo mice model

Exclusive transcellular transport reported and no evidence of particles in olfactory bulb or olfactory axons was observed.

Mistry et al.57

19

Mistry et al. investigated the possibility of nose-to-brain transfer of chitosan-

coated polystyrene and polysorbate-coated polystyrene nanoparticles (100 – 200 nm in

diameter) using fluorescence microscopy and stereology techniques following repeated

intranasal administration every 24 h for two days in mice57. The authors found no

evidence of either of the nanoparticle types in the olfactory axons or the olfactory bulbs

of mice after the 4th day of first administered dose; instead, the majority of particles

remained in the olfactory epithelial cells. The use of nonbiodegradable polystyrene

polymeric particles allowed the authors to visualize the nanoparticles using fluorescent

microscopy to observe nasal epithelial cross-sectional images and authors were able to

conclude that nanoparticles should have a diameter of < 100 nm for transcellular

transport of nanoparticles through the olfactory axons beyond the basement membrane.

Huang et al. reported the transcellular and paracellular transfer of 10 nm carboxylate

surface modified polystyrene nanoparticles following exposure to excised rabbit nasal

mucosa using fluorescence microscopy41. In another study, Chen et al. demonstrated

greater uptake of 20 nm sized carboxylate polystyrene nanoparticles than 100 nm sized

nanoparticles into excised bovine nasal respiratory and olfactory tissue using

spectrophotometric and fluorescence detection of the incorporated dye38. Chen also

showed that intact nanoparticles could be taken up into nasal tissues using multiple

endocytic mechanisms, and additional paracellular pathways for smaller, 20 nm

nanoparticles were also reported38. Although these nonbiodegradable nanoparticles are

unlikely to be effective drug delivery systems, they serve as excellent models for

systematically studying the size and surface properties of nanoparticles that influence

cellular interactions and biodistribution patterns.

20

Biodegradable Polymeric Nanoparticles

Natural polymers like chitosan, gelatin, alginate and synthetic polymers like

polylactic acid (PLA), poly (lactic-co-glycolic acid) (PLGA), and poly-caprolactones, are

some of the biocompatible and biodegradable polymers used in nanoparticle

preparations58. Nanoparticles made of biodegradable materials may provide controlled

drug delivery, lower toxicity, and improved biocompatibility, with minimal immunogenic

and inflammatory responses59. Table 1-2 summarizes some of the selected biodegradable

nanoparticulate systems evaluated as nasal drug delivery systems. PLGA and PLA are

most frequently used in nanoparticulate delivery systems because these polymers undergo

hydrolysis and produce the biodegradable monomers, lactic acid and glycolic acid.

PLGA particles are being studied widely for their ability to deliver large molecules like

vaccines and DNA or RNA for gene delivery strategies. Surface-modified PLGA

particles have been repeatedly shown to improve the induction of systemic and mucosal

immune responses. For example, Pawar et al. reported augmented systemic and mucosal

immune responses after intranasal administration of Heptitis B Surface Antigen (HBsAg)

in PLGA particles of diameter 160-180 nm in mice60. They also showed that modifying

the surface of PLGA particles using chitosan and trimethyl chitosan ligands increased the

zeta potential of the HBsAG PLGA particles, which further potentiated the mucosal

immune response. Along with nasal vaccinations, PLGA particles have also been

reported to deliver small drug molecules to the brain via the intranasal route. Seju et al.

developed olanzapine-containing PLGA nanoparticles of ~90 nm diameter for intranasal

delivery to the brain and reported a ~10 times increase in olanzapine concentrations in

brain when compared to olanzapine solution when administered through the intranasal

21

route in rats61. Most formulations in the nasal cavity are cleared rapidly through

mucociliary clearance, and such rapid clearance from the nasal cavity limits the

internalization of particulate therapeutic agents. To overcome these limitations, several

varieties of surface modifications using ligands like chitosan, trimethyl-chitosan, lecithin

and polyethylene glycol (PEG) have been examined62-63. Surface modification with

anionic or neutral polymers increases the otherwise negative zeta potential of PLGA

particles and may improve mucoadhesion, which may enhance the residence time in the

nasal mucosa. For example, surface modification of PLGA particles with chitosan or its

derivative, glycol chitosan has shown to improve the retention time of Hepatitis B surface

Antigen in the nasal mucosa compared to the uncoated PLGA particles64. Also, the glycol

chitosan coating of PLGA particles was shown to induce significantly higher systemic

and mucosal immune response compared to uncoated PLGA particles64. Although

biodegradable nanoparticles offer several advantages, researchers need to be careful to

understand the particle aggregation properties of these systems and their mechanisms of

particle transit in the nasal mucosa when investigating them as nanoparticulate delivery

systems. Nanoparticles deposited on the nasal mucosa can potentially release drug into

the mucosal secretions and the free drug has the potential to transfer into the nasal

mucosa or the drug can also be released within the nasal mucosa from internalized

nanoparticles without the particles being further translocated. Hence, it is important to

study the drug release behavior from these nanoparticles to better understand how

nanoparticles can potentially improve the drug delivery across the nasal mucosa.

22

Table 1-2 Summary of selected studies showing potential use of biodegradable PLGA and PLA particles in delivering a variety of drugs and vaccines via the intranasal route.

Type of particulate system

Particle size (nm)

Antigen/Drug Experimental

model Findings Reference

Unmodified PLGA, chitosan

coated and trimethyl chitosan

(TMC) coated

420 – 490

Heptitis B surface antigen

BALB/c mice

TMC coated PLGA particles induced substantially higher antibody titers (anti HBsAg) in local and distal mucosal secretions

as compared to chitosan coated, uncoated PLGA and soluble or alum adsorbed HBsAg.

Pawar et al.60

PLGA ~180 Diazepam Sprague

Dawley Rats

Approximately 2 -3-fold increase in brain/blood diazepam concentration was observed with PLGA particles compared to

drug in solution

Sharma et al.65

Wheat germ agglutinin

conjugated PEG-PLA

~110 6-coumarin Sprague

Dawley Rats

Nanoparticles were transferred into the olfactory submucosa through transcellular pathways followed by subsequent transfer to

olfactory bulb through nerve bundles or their surrounding connective tissue

Liu et al.66

PLA, PEG-PLA 200 Tetanus toxoid BALB/c mice PEG-PLA particles showed ~2-fold increase in anti-tetanus IgG antibodies after 24 weeks compared to PLA particles or antigen

solution Vila et al.67

PLGA 91 Olanzapine Albino Rat Approximately 6 - 8 times higher concentration of drug in brain

with PLGA particles compared to drug in solution Seju et al.61

PLGA ~70 Nile red Excised Bovine nasal Mucosa

Around 4% of exposed nanoparticles internalized into nasal mucosa within 1 h

Greater uptake into olfactory tissue compared to respiratory tissue

Majority of particles were observed in submucosal regions of nasal tissues

Albarki et al.68

23

Metallic Nanoparticles

Nanoparticles made of metals, metal alloys, metal oxides and other metallic salts

can be generally described as “metallic nanoparticles”. These include nanoparticles of

gold, silver, copper, iron oxide, manganese oxide, and indium oxide. Also included in

this category semi-conductor-based quantum dots and non-metal, carbon-based

nanoparticles like fullerenes, graphenes and carbon nanotubes. Several metallic

nanoparticles have been shown to transfer from nose to brain, lungs and other distant

regions14, 69-72. In 1960, De Lorenzo reported nanomaterials, less than the diameter of the

olfactory axons, can be transferred through these axons, and the author showed evidence

of colloidal gold in the olfactory axons beyond the basement membrane of the olfactory

mucosa of squirrel monkeys following intranasal instillation73-74. Olfactory axons in

humans 100 – 700 nm in diameter and the colloidal gold used by DeLorenzo was

~50 nm75. Following nasal exposure, the extremely small size of these metallic

nanoparticles may result in entrapment in olfactory neuronal cells in the nasal mucosa

with subsequent transfer to the olfactory bulb and other regions of the brain. In another

study, Edler and coworkers reported a 3-4-fold increased concentration of manganese in

the olfactory bulb compared to control after a 12-day inhalation exposure of 30 nm

manganese oxide nanoparticles in rats72. In a more recent study, Hopkins et al. used

fluorescent microscopy and showed that short-term inhalation exposure of 15 – 20 nm

diameter solid, cadmium-selenide quantum dot nanoparticles in mice resulted in axonal

transport to the olfactory bulb76. A study by Garzotto and Marchis investigated the

localization of cadmium selenide quantum dots in the olfactory epithelium following

intranasal administration in CD1 mice71. These investigations imaged cross-sections of

24

the nasal tissues using confocal microscopy and reported that quantum dots were able to

cross the olfactory epithelium and reach the underlying lamina propria. An absence of

quantum dots in the olfactory neurons caused the authors to conclude that quantum dots

cross the olfactory epithelium through extracellular spaces around the olfactory neuronal

cells and supporting cells. The investigations did not report any quantum dots being

transferred to the brain, however, but they did show small amounts of quantum dots

present around the olfactory nerve bundles in the submucosal 24 h after intranasal

administration. Quantum dots in the lamina propria have the potential to reach the brain

by transfer along the perineuronal spaces, and small quantities of particles in nerve

bundles could transfer through the neuronal axons and reach the brain. A summary of the

few studies showing the transfer of metallic nanoparticles from the nose to the brain

following either intranasal instillation or inhalation exposure is shown in Table 1-3.

These investigations have led researchers to engineer advanced metallic

nanoparticles for disease diagnosis and also for studying the efficiency of targeted drug

delivery to the brain77-80. For example, Joshi and coworkers administered insulin attached

to colloidal gold (~4 nm diameter) via per-oral and intranasal routes in diabetic Wister

rats and compared the resulting reduction in blood glucose levels81. They reported that

within 2 hours of dosing an approximately 50% reduction in blood glucose levels

following intranasal administration but only a 19% reduction following per-oral

administration. Although gold nanoparticles served as excellent carriers for insulin, the

clearance, accumulation and toxicity of gold following intranasal administration was not

discussed in the report; gold accumulation is likely a safety concern regarding the

sustained and chronic use of these nanoparticles to treat type I diabetes.

25

Table 1-3 Summary of studies showing translocation of metallic nanoparticles following intranasal administration.

Nanoparticle type Size (nm) Experimental

model Findings Reference

CdSe/ZnS QDs encapsulated in

polyethylene glycol, phosphatidyl,

ethanolamine micelle

15-20 Mice

Three hours after inhalation exposure, quantum dots were visualized in olfactory axons and olfactory bulb

Authors concluded transfer for QDs from nose to brain via olfactory axonal transport

Hopkins et al.76

Carboxylate-modified CdSe/ZnS QDs

15-20 Mice

Transfer of QDs was visualized from olfactory epithelium to lamina propria within 4 h post instillation

No evidence of QDs in olfactory neuronal components, instead QDs were present as clusters in extracellular spaces in lamina

propria

Garzotto et al.71

Manganese oxide particles

~ 30 Rats 12 days of exposure resulted in 3.5-fold increase of Mn in olfactory bulb and ~1 fold increase in Mn content in lungs

Elder et al.72

Radiolabeled carbon particles (13C )

36 Rats Gradual increase in 13C carbon in the olfactory bulb from day 1 to

day 7 following 6 h inhalation exposure Oberdorster

et al.70

Iron oxide particles 280 Mice

Higher iron levels in olfactory bulb and brain stem 2 week after instillation

Possibility of both olfactory neuronal and trigeminal pathways for brain uptake

Wang et al.82

Silver coated gold particles

~50 Squirrel

Monkeys

Within 30 - 60 min of instillation, gold particles were detected in olfactory neuronal axons and olfactory bulb which showed the

involvement of olfactory neuronal pathways

De Lorenzo et al.74

26

Recent advances in nanoparticle engineering have led to the synthesis of a variety

of surface functionalized metallic nanoparticles with high stability in biological

environments. The inherent metallic properties of nanoparticles, including the magnetic

properties of iron oxide particles, optical properties of quantum dots and high density of

gold particles, make these particles excellent choices for magnetic resonance imaging

(MRI), multimodal optical imaging and computerized tomography (CT), respectively83-85.

Intranasal delivery of metallic nanoparticles for diagnostic purposes is still limited to cell

based and small animal studies due to the potential for accumulation of a heavy metal

load in the body, as the clearance properties of these of nanoparticles is uncharacterized.

Nevertheless, improved diagnostics used in cells and small animals can still have a huge

impact on life science research which could assist considerably in translational research.

A rapid increase in the utilization of metallic nanoparticles has been seen, not

only in the biomedical fields, but also in the automobile and consumer product

industries86-87. This increased utilization is accompanied by the potential risk of increased

levels of nanomaterials in the environment, which also cause increased human exposure

to nanoparticles.

One of the major routes of human exposure to these extremely small, elemental

nanoparticles is through inhalation. Bioaccumulation of unwanted harmful airborne metal

nanoparticles along the olfactory neurons and in the brain could potentially result in

several CNS-related diseases like Alzheimers88 and Parkinsons diseases89. As multiple

investigations have shown the potential transfer of nanomaterial from the nose to the

brain, the potential neurotoxicity associated with accumulation of metal particles has also

been widely studied (Table 1-4).

27

Table 1-4 Summary of selected studies showing toxicity effects of metallic nanoparticles following intranasal instillation.

Nanoparticle type Size (nm) Animal Model

Toxicity Findings References

Ferric oxide 21 and 280 Mice

Induction of oxidative stress and elevation of glutathione levels in olfactory bulb and hippocampus

Alterations in nerve cells, neurodendron degeneration, membrane disruption, swollen mitochondria and dilation in

rough endoplasmic reticulum in olfactory bulb

Wang et al.90

Silica 15 Rats

Induction of oxidative stress and increased inflammatory response in striatum.

Depletion of dopamine and down regulation of tyrosine hydroxylase

Wu et al.91

Titanium dioxide 80 and 155 Mice Increased tumor necrosis factor alpha and interleukin

Increased oxidative damage due to lipid peroxidation Wang et al.14

Silver 25 Mice

Elevated tissue glutathione levels in nasal epithelia

Increase oxidative stress in nose and in blood

Mild enhancement of erythrocyte destruction

Genter et al.92

28

Solid Lipid Nanoparticles

Solid lipid nanoparticles (SLNs) constitute attractive colloidal drug delivery

systems consisting of lipid particles dispersed in aqueous surfactant solutions. Generally,

SLNs contain a solid hydrophobic core made of lipids stabilized using surfactants. Lipids

that are well tolerated by the body like phospholipids (e.g. phosphatidyl choline,

phosphatidylehanolamine), triglycerides (e.g. tristearin), fatty acids (e.g. stearic acid) and

waxes (e.g. cetyl palmitate) can also be used for the production of the solid lipid cores93.

Unlike most polymeric nanoparticles, lipids used in SLNs are known to be biocompatible

and biodegradable, hence they may have minimal cytotoxicity. Furthermore, production

of polymeric nanoparticles involves the use of organic solvents and there is always a

concern for the residual solvents in the finished product, whereas production of SLNs

does not need to employ potentially toxic organic solvents94. SLNs can be prepared using

simple techniques like high pressure homogenization, and can be easily scaled to large-

scale production compared to more sophisticated production methods required for

polymeric nanoparticles95. SLNs are similar to microemulsions and liposomes, differing

primarily in the lipid physical state. The solid lipid in SLNs replaces liquid/semi-solid

phase lipids used in vesicular systems. Because of the solid-state lipid in the core, SLNs

show better stability than lipid vesicular systems. Concern regarding the safety of

surfactants used as stabilizers in SLNs has not been established, however, and this is a

significant limitation in developing safer SLN systems.

Lipophilic drugs can be dissolved or dispersed in the lipid portion of the

phospholipids and hydrophilic drugs can be dissolved or dispersed in the hydrophilic

portion of the phospholipid matrix, thus SLNs have the potential to act as carriers for a

29

variety of hydrophilic and lipophilic drugs/vaccines 94. The hydrophobic lipid cores of

SLNs can be tuned to a variety of sizes and surface characteristics and enable controlled

and targeted delivery of drugs by various routes of administration. One of the widely

studied applications of SLNs is targeting drugs to the brain following intravenous and

intranasal administration. Intravenous administration of surface engineered SLNs with

appropriate surfactants like polysorbate 80 and/or appropriate ligands like polyethylene

glycols have been shown to escape clearance by the reticuloendothelial system and allow

enhanced uptake of SLNs via receptor-mediated endocytosis across the blood-brain

barrier96-97. Several reports in the literature also describe the superiority of the intranasal

route over the intravenous route for delivery of drugs in SLNs. Table 1-5 summarizes

some of those investigations. For example, Patel et al. developed risperidone SLNs (150

nm diameter) using glyceryl behenate as the lipid and Pluronic F-127 as the surfactant.

The authors reported rat brain/blood ratio ~ 5 fold higher 1 h after administration when

delivered via intranasal route compared to the intravenous route98.

Liquid-phase Nanopdispersions in Intranasal Drug Delivery

The term “liquid-phase nanodispersions” is used to describe colloidal dispersions

of liquid/semisolid-state nanomaterials of diameters 1-1000 nm dispersed in continuous

liquid medium. Liquid-phase nanodispersions have been employed as carriers for drugs

and vaccines for several decades. Alternative terminologies for liquid-phase dispersions

are used in literature including vesicular systems, which include microemulsions,

liposomes, niosomes, and micelles. Liposomes and microemulsions are widely studied as

liquid-phase, nasally-administrated nanodispersions, and a brief review of these systems

is summarized in the following sections.

30

Table 1-5 Summary of studies investigating intranasal uptake of drugs/antigens using solid lipid nanoparticle systems.

SLNs composition Particle

size (nm) Drug

Experimental model

Findings Reference

Compritol 888 ATO and Pluronic F-127

148 Risperidone Mice 5-fold higher brain/blood ratio of drug was

observed1 h post administration via intranasal route compared to intravenous route

Patel et al.98

Compritol 888 ATO, Tween 80, Soy Lecithin

140 Streptomycin

sulfate Mice

3.5-fold higher bioavailability of drug in brain when delivered in SLNs compared to free drug in solution

Kumar et al.99

Glycerol monostearate, Tween 80, Soy lecithin

150 Rosmarinic

acid Rats

Intranasal delivery of SLNs produced significant therapeutic action compared to intravenous delivery

of SLNs

Bhatt et al.100

Glycerol monostearate, Lecithin, Poloxomer 188

320 – 500 Ondansetron

HCl Rabbits

Accumulation of radiolabeled drug in the brain and distant tissues was observed from 1 h to 6 h after

intranasal administration

Joshi et al.101

31

Liposomes

Liposomes are spherical vesicles composed of lipid bilayers made of natural,

biodegradable, nontoxic constituents like phospholipids surrounding a central aqueous

core. They may contain cholesterol as a membrane stabilizer that helps maintain their

fluid-like physical state. Unique bilayer structures mimic natural cell membranes and

incorporating charged agents can provide the opportunity to alter the liposomal surface

chemistry. Liposomes can encapsulate a wide variety of lipophilic drugs in the lipid

bilayers and hydrophilic drugs can be retained in the aqueous cores. Wide size ranges of

liposomes have been reported (from 150 nm to > 5000 nm) with positive (cationic),

negative (anionic) and neutral surface charges. In general, encapsulation of vaccines or

drugs in the inner core of liposomes is preferred because it protects the cargo from

enzymatic and chemical degradation inside the body102.

In nasal delivery, liposomes are most widely investigated as carriers for vaccines.

A variety of viral (e.g., influenza103, hepatitis B virus104) and bacterial (e.g.,

mycobacterium tuberculosis105, pseudomonas aeruginosa106) pathogens have been the

target for nasal vaccines. Some of these studies are summarized in Table 1-6.

Production of immunoglobulin A (IgA) at mucosal surfaces is a critical first line

of defense against pathogens that infect the host. Studies in animal models investigating

the efficacy of liposomal vaccine systems delivered through the intranasal route have

demonstrated local production of specific IgA in nasal, bronchoalveolar, vaginal and

rectal secretions and there are also reports showing increased production of IgA in saliva

and bile102.

32

Table 1-6 Summary of studies showing intranasal uptake of drugs/vaccines using liposomal vesicular systems.

Liposomes composition Vesicle

size (nm) Drug/antigen

Experimental model

Findings Reference

Distearoylphosphatidyl choline, cholesterol, N-

glutarylphophatidyl ethanolamine, hemaglutinin

712 Heptitis B

surface antigen (HBsAg)

Mice

Liposome formulation showed high levels of anti-HBsAg antibody titers in plasma, nasal, salivary,

vaginal and intestinal secretions compared to antigen alum formulation.

Both mucosal and systemic immune responses were observed with liposomes

Tiwari et al.104

Phosphatidylcholine, cholesterol

2300 Tetanus toxoid Rabbits After 10th week post-vaccination, liposomal formulation showed highest IgA titers in nasal lavage compared to

solution or alum formulation

Tafaghodi et al.107

Egg phosphatidylcholine, dioleylphosphatidylethanala

mine, cholesterol, glycol chitosan coating

1000 Plasmid

pRc/CMV-HBs (DNA)

Mice High humoral mucosal immune response (high sIgA

antibody levels) with liposomal formulations compared to naked DNA solution

Khatri et al.108

Egg phosphatidylcholine, cholesterol

166 Rivastigmine Rats Higher levels of drug in olfactory bulb and other brain regions were observed within 15 min after intranasal

administration Yang et al.109

33

Nasal immunization has been shown to trigger both humoral and celluar immune

responses102. There are also reports showing liposomal formulations of small molecules

increasing the distribution of small drugs across the nasal mucosa to reach the systemic

circulation and CNS. Yang and coworkers studied the pharmacokinetics and

pharmacodynamics of 179 nm rivastgmine-containing liposomes following intranasal

delivery in rats. Intranasal administration of a rivastgimne liposomal formulation showed

greater distribution of rivastigmine to the CNS, plasma, liver and kidney compared to the

intravenous injection of rivastigmine solution109. In spite of these advantages, intranasal

liposomal formulations are still not commercially available. Chemical instability of

liposomal formulations due to degradation of the liposome components and physical

instability including size changes and loss of entrapped drug upon storage are major

problems in liposomal formulations110. Overcoming these limitations could make

liposomal formulations promising nanoparticulate drug delivery systems for intranasal

administration.

Microemulsions

Microemulsions are thermodynamically stable, isotropic translucent systems

consisting of oil and water with surfactants and stabilizers. The dispersed phase of a

microemulsion is typically 10-100 nm in diameter. Microemulsions differ from

macroemulsions in two significant ways. Microemulsions are thermodynamically stable

systems whereas macroemulsions pose problems like creaming, coalescence and phase

separation. The dispersed phase in microemulsions is typically in the low nanometer size

range and thus microemulsions are transparent systems whereas the dispersed phase in

macroemulsions is frequently several microns in diameter and the systems are turbid or

34

milky white111. Microemulsions are pseudo-ternary systems which spontaneously form

upon mixing specific proportions of oil, water and surfactants. Stabilizers and/or

cosurfactants are typically used in these systems to impart thermodynamic stability to the

systems. These surfactants and stabilizers help to reduce the interfacial tension between

oil and water by adsorbing to the interface and thus surfactants assist in the formation and

stabilization of microemulsions. Depending on the type of surfactant used, two types of

microemulsions can be formed. Typically, a hydrophilic surfactant (high HLB value) aids

in emulsifying oil throughout the continuous water phase to form an oil-in-water (o/w)

microemulsion. Similarly, a lipophilic surfactant (low HLB value) aids in emulsifying

water throughout the continuous oil phase and forms a water-in-oil (w/o)

microemulsion112.

By virtue of their lipophilic nature, small size and good physical stability,

microemulsions have been explored as delivery systems to enhance uptake across the

nasal mucosa. Several studies showed improved transfer of drugs to the brain and

systemic circulation when delivered in microemulsions compared to aqueous solutions.

Some of these studies are summarized in Table 1-7. For example, risperidone

microemulsion consisting of Capmul MCM, Tween 80, and propylene glycol showed

~14% higher blood-brain ratio of drug 30 min after delivery via intranasal administration

in rats compared to intravenous administration of same formulation. Incorporation of

chitosan, a mucoadhesive polymer in the microemulsion system, resulted in a further

increase in the brain-blood ratio for resperidone when delivered via the intranasal

route113. Microemulsions enable the formulation of high concentration dosage forms

suitable for nasal administration. For example, the solubilization capacity of nimodipine,

35

a poorly water-soluble drug (2.3 μg/mL) was increased over 2700 fold to 6.4 mg/mL

when formulated in a microemulsion system consisting of Labrafil M 1944CS,

Cremophor RH 40, ethanol and water. Following intranasal administration of this

nimodipine microemulsion, a three-fold increase in nimodipine uptake in the olfactory

bulb compared with intravenous injection of drug in a solution containing ethanol, PEG

400 and water was reported 114.

In spite of their advantages, there are no commercial microemulsion drug delivery

systems available in the market designed for the intranasal route. Their drug loading

capacity and use of high concentrations of excipients limits the use of microemulsions.

Surfactants and cosolvents can be toxic at high doses; formulators are limited in the range

of compositions in development of microemulsion systems with high drug loading

capacities while minimizing use of the surfactants and cosolvents.

Formulation of lipophilic drugs as non-aqueous concentrates in soft gelatin

capsules is another alternative delivery system that gained interest of pharmaceutical

industry. These non-aqueous drug concentrates are designed to form microemulsions

upon dilution with water immediately before administration or following administration

and dilution with gastric fluids115. In cases where these drug concentrates form clear

transparent microemulsions upon dilution, the concentrates are called self-

microemulsifying drug delivery systems (SMEDDS). While most of the systems being

developed are intended for oral delivery116-118, SMEDDS have the potential to be used in

intranasal delivery, where the drug concentrates can be diluted immediately before

administration and then sprayed into the nasal cavity.

36

Table 1-7 Summary of studies showing intranasal uptake of drugs/vaccines using microemulsion systems.

Microemulsion Composition Vesicle

size (nm) Drug

Experimental model

Findings Reference

Labrafac PG, Labrasol, Transcutol HP, water

60 Saquinavir mesylate

Rabbits 12-fold increase in bioavailability of microemulsion

formulation compared to commercial tablet taken orally Hosny et

al.119

Medium chain triglyceride, polyoxytheylene-35-

ricinoleate, polysorbate 80, propylene glycol, water

15 Clonazepam Rats ~2-fold increase in brain/blood drug ratio with

microemulsion compared to intravenous administration Vyas et

al.120

Capmul MCM, Labrasol, Tween 80, Transcutol P,

water 35

Quetiapine fumarate

Rats 3.8-fold higher bioavailability in brain with microemulsion

compared to drug solution Shah et

al.121

Capmul MCM, Tween 80, Propylene glycol, Transcutol

P, water 15 Risperidone Rats

~14% higher brain-blood ratio of drug after 30 min compared to intravenous administration of same

formulation

Kumar et al.113

Labrafil M 1944CS, Cremophor RH 40, ethanol,

water 25 – 30 Nimodipine Rats

3-fold increase in uptake of drug to olfactory bulb compared with intravenous injection of drug in solution

Zhang et al.114

37

Summary

The nasal route has been widely studied for the delivery of drugs and vaccines

using a variety of colloidal delivery systems. Even though significant progress has been

made, the commercial availability of colloidal nasal dosage forms for systemic delivery

and delivery to the brain are very limited. Many questions about the safety and efficacy

of the colloidal delivery systems remain elusive. In addition, poor drug loading capacity

and high production costs limit usage of colloidal delivery systems. Although significant

progress has been made, the effect of particle/vesicle size and surface characteristics on

particle uptake is still unclear. Several reports suggest that uptake of smaller diameter

nanoparticles into nasal tissues is greater compared to larger particles of similar

composition. Also, there are reports which show smaller size nanoparticles with positive

surface charge show significantly reduced uptakes into nasal tissues, compared to larger

particles with negative surface charge. Additionally, it is unclear how these colloidal

systems are internalized or absorbed into the nasal mucosa. Most of the studies that report

improved delivery of drugs from nasal mucosa fail to investigate or report on the

underlying mechanisms of particle uptake into the nasal respiratory and olfactory mucosa

or provide sufficient information about the release of the drug contents from the

nanoparticle to determine how the particle-associated drug will be presented once

absorbed or transferred into the mucosa.

The studies designed in this research aim to address the existing knowledge gap

regarding ultrafine nanoparticle (< 20 nm) transfer into the nasal mucosa by investigating

the following important questions: 1) How much of the applied nanoparticle load is

transferred into the nasal tissues? 2) Where do the absorbed nanoparticles distribute

38

within the nasal cavity and throughout the body? 3) What uptake pathways can

nanoparticles access? and 4 ) Does the physical state of the nanoparticles (solid vs semi-

solid) affect the efficiency of nasal uptake?

These questions will be addressed in the following chapters: Chapter 3 addresses

Questions 2 & 3 and describe the investigation of the uptake mechanisms of ultrafine

nanoparticles following measurement of the concentration of nanoparticles in various

tissue regions. These are some of the first studies to use chemical analytical methods to

investigate nanoparticle uptake and distribution in tissues. The localization of such

extremely small nanoparticles within those tissue regions will also be shown using

confocal and electron microscopy techniques. Chapter 4 addresses Questions 2 & 3 by

describing the results following the use of non-invasive in vivo imaging methods to

demonstrate the fate of intranasally instilled ultrafine nanoparticles. The potential use of

whole animal imaging and micro-CT techniques for qualitative and semi-quantitative

analysis of nanoparticle biodistribution provides additional methods to investigators to

understand the fate of nanoparticles following intranasal administration. Finally, in

Chapter 5, an attempt was made to study the effect of the physical state of the

nanomaterials on their uptake mechanisms in nasal mucosa by using a drug-containing

microemulsion system. This research overall was aimed at addressing Question 4 and

investigates a new type of colloidal nasal delivery system.

39

CHAPTER II

OBJECTIVES

Colloidal dispersions, irrespective of physical state (solid vs semi-solid) have

been shown to enhance the delivery of therapeutic agents via the nasal mucosa to the

brain and/or the systemic circulation. Although there is evidence that intact nanomaterials

transverse the respiratory and olfactory epithelial barrier, the nasal mucosa represents an

effective barrier against the uptake of many other particulates. Careful characterization of

the uptake and distribution of particles into the olfactory and respiratory tissues, the

brain, and the systemic vasculature is needed to identify particle characteristics that can

be leveraged for advanced drug delivery strategies.

The central hypothesis of this work focuses on nanoparticulate uptake in the nasal

mucosa is that: extremely small nanoparticles (1-20 nm) have access to multiple uptake

pathways through the nasal mucosa, including convective transport within the

paracellular spaces, endocytic uptake by the epithelial cells, and additional neuronal-

associated pathways. A secondary hypothesis is that: the physical state of the

nanomaterials (solid vs semi-solid) and surface characteristics of the nanomaterials

influence their uptake mechanism.

The rationale for the pursuit of this research is that the determining the particle

characteristics which contribute to nasal uptake can result in improved targeted therapies

for a multitude of diseases. Knowledge of nanoparticle uptake can also be applied to

avoid potential neurotoxicities through the appropriate selection of nanomaterials. These

hypotheses will be tested the following specific aims:

40

Specific Aim 1: Investigate the extent of quantum dots (< 20 nm) uptake into

bovine respiratory and olfactory mucosal explants using inductively coupled plasma-

optical emission spectroscopy to analyze uptake and evaluate their localization in nasal

tissues using confocal and electron microscopy.

Specific aim 2: Evaluate the extent of PEGylated quantum dots uptake into bovine

nasal mucosa and compare the uptake with carboxylate-modified quantum dots to

determine the effect of surface modification on nanoparticle transit into nasal tissues.

Specific aim 3: Identify the pathways involved in the uptake of quantum dots into

the bovine nasal respiratory and olfactory mucosal explants by probing energy-dependent

and energy-independent uptake pathways into the bovine nasal mucosa.

Specific aim 4: Investigate the effect of physical state (solid vs semi-solid) on the

uptake of nanomaterials into nasal mucosa by comparing the uptake properties of solid

quantum dots (QDs) nanoparticles and semi-solid (microemulsion) nanodroplets. Prepare

diazepam-containing microemulsions and investigate diazepam permeation into/across

bovine respiratory and olfactory mucosal explants. Identify the pathways involved in the

uptake of microemulsion into nasal mucosa by probing the energy-independent pathways.

Specific aim 5: Investigate and characterize the biodistribution of QDs with

different surface modifications (carboxylated and PEGylated) after intranasal

administration in mice using non-invasive in vivo fluorescence imaging and determine

involvement of endocytic pathways in the uptake of quantum dots in mice. Identify the

transverse pathways of gold nanoparticles from nasal cavity of mice using micro-CT

whole animal imaging.

41

CHAPTER III

UPTAKE AND TRANSPORT PATHWAYS FOR ULTRAFINE

NANOPARTICLES (QUANTUM DOTS) IN THE NASAL MUCOSA

Introduction

The applications of nanotechnology in the biomedical and pharmaceutical

industries have expanded rapidly during the past decade, particularly for the delivery of

drugs and vaccines and also in medical imaging for diagnosing and detecting disease11.

Studies have shown that both engineered nanoparticles (e.g. carbon nanotubes, quantum

dots) and naturally occurring nanoparticles (e.g. smoke, dust, viruses) appear in the

central nervous system (CNS) following intranasal exposure13-15. Additional reports are

also available showing that a variety of nanoparticles enter the systemic circulation

through the vascular bed in the nasal mucosa following intranasal administration37. As a

result, nanoparticulate systems are being used to exploit the nasal route to deliver drugs

locally, systemically and directly to the brain.

Particle size and surface chemistry are the important characteristics of particulate

systems that affect their extent of uptake through nasal tissues. An interesting study by

Brookings et al.37, demonstrated that the rate and extent of translocation of 125I-

radiolabelled latex nanoparticles (20, 100, 500 and 1000 nm) into the systemic circulation

following nasal administration in rats was dependent on the size and surface charge of the

nanoparticles. They observed that 20 nm sulfated polystyrene particles showed greater

uptake into the blood (3.25 % of administered dose) compared to larger particles.

Additional studies have shown the extent of uptake of small nanomaterials (< 50 nm) like

metals, metal oxides, and viruses was higher through the nasal mucosa when compared to

42

nanoparticles with diameters > 50 nm69, 72, 92. There are also several reports available in

the literature showing that nanoparticles with diameters less than 20 nm showed

increased deposition in the nasal cavity, compared to particles >100 nm, more typical of

the particle sizes most frequently studied as drug delivery systems70, 122. In general,

although ultrafine nanoparticles (< 20 nm) show higher translocation efficiencies,

limitations including low drug encapsulation efficiencies and high cost of production

continued to motivate investigators to select larger particles (>100 nm) for drug delivery

purposes. While there are a limited number of reports available in the literature

investigating the extent of uptake of ultrafine nanoparticles (< 20 nm) in the nasal

mucosa, better understanding of the uptake and transfer of these ultrafine particles is

necessary in order to identify solutions where these extremely small particle sizes may be

preferred for drug delivery and in contrast, when these ultrafine particles represent a

toxicological concern.

The major routes and the extent of uptake involved in the internalization of the

nanoparticles into tissues depend on the properties of the particles, including their size,

surface charge and shape123. Phagocytosis, macropinocytosis, clathrin-mediated

endocytosis and caveolae-mediated endocytosis are currently the major pathways which

have been identified in the regulation of entry of physiologically relevant

materials/macromolecules into cells46. Nanoparticles can also be entrapped within these

endocytic vesicles and enter into cells. Previously, Huang et al. reported that 10 nm

carboxylate-modified polystyrene nanoparticles and 200 nm amine-modified polystyrene

particles were found in both the paracellular spaces and in trancellular locations

(endocytosis into interior of cells) of excised rabbit nasal mucosa41. More recently,

43

multiple size-dependent pathways within the bovine nasal mucosa have been investigated

using fluorescently-labeled polystyrene particles (20 nm and 100 nm). It was observed

that 100 nm particles were primarily taken up by macropinocytosis in both the respiratory

and olfactory mucosa whereas a clathrin-mediated pathway appeared to be primarily

involved in the uptake of 20 nm nanoparticles across the respiratory mucosa38. Uptake

was dependent not only on the size and surface chemistry of the particles but also on the

epithelial region within the nasal cavity. Further understanding of the underlying

mechanisms of the membrane transport and distribution of nanoparticles (< 20 nm) is

essential to assist in the design and development of advanced nanoparticulate drug

delivery systems with improved efficiency for nasally-administered therapeutics.

The aim of the present study was to investigate the uptake of ultrafine

nanoparticles in the bovine nasal mucosa and to study the pathways involved in that

uptake. Quantum dots (QDs) were chosen as the model for ultrafine nanoparticles for the

purpose of this study. QDs are readily-available, inorganic nanocrystals (1 – 10 nm)

composed of a heavy metal core (e.g. CdSe) with an inorganic shell (e.g. ZnS). The

unique, size-specific optical properties of QDs have encouraged their application in a

number of in vivo and in vitro systems in biomedical and pharmaceutical sciences124-125.

Compared to conventional organic dyes and fluorescent proteins, QDs offer

unique advantages, such as size dependent optical properties, large absorption

coefficients across a wide spectral range and high levels of brightness and

photostability124. These unique properties make QDs very attractive probes for diagnostic

purposes using molecular imaging and in vivo imaging. QDs were observed to be rapidly

cleared from the blood circulation following intravenous injection and have been detected

44

in the reticuloendothelial tissues, including liver, spleen, lymph nodes and bone marrow

of animals for up to 2 years after administration126-129. There are also reports showing QD

transfer through the blood-brain barrier to reach the brain following intraperitoneal

injection in mice130. Yet, Choi et al. reported that QDs of < 6 nm with cysteine coating

were rapidly cleared through the kidneys131. Additional investigations of the

biodistribution of quantum dots in the body are needed to fully understand the importance

of their properties, including size and surface chemistry, on their fate within the body.

Materials and Instrumentation

Fluorescent quantum dots (QDs) with cadmium selenide (CdSe) core and zinc

sulfide (ZnS) shell were purchased from NN-Labs, LLC (Fayetteville, AR). These

particles are available with a variety of surface modifications; QDs with a carboxyl

surface group (COOH-QDs) or polyethylene glycol surface-modified QDs (PEG-QDs)

were used in these studies. Nanoparticles ~ 7 nm diameter in size with emission

wavelength of 620-640 nm were selected. A sample optical spectrum of COOH-QDs is

shown in Appendix-A. PEG-QDs were used to study the effect of hydrophilic surface

modification only and COOH-QDs were used for all other studies.

2, 4-Dinitrophenol, chlorpromazine, methyl-β-cylcodextrin, amiloride,

glutaraldehyde solution (50% in water), osmium tetroxide solution (4% in water), and

sodium glycocholate hydrate were purchased from Sigma Aldrich Co. (St. Louis, MO).

Sucrose, potassium ferrocyanide and uranyl acetate were obtained from Spectrum

Chemicals (New Brunswick, NJ). Vertical Navicyte® diffusion chambers were obtained

from Harvard Apparatus (Hollison, MA) and were used to perform transport studies. The

exposed tissue area was 0.64 cm2, and each chamber reservoir capacity was 2 mL.

45

Krebs Ringer Bicarbonate buffer (KRB) consisting of 123.8 mM sodium

chloride, 1.5 mM monobasic sodium phosphate, 4.56 mM potassium chloride, 10 mM

dextrose, 15 mM sodium bicarbonate were purchased from Research Products

International Corp (Mt. Prospect, IL). KRB also contained 1.67 mM magnesium chloride

(Sigma Aldrich, St. Louis, MO), 0.7 mM dibasic sodium phosphate (Sigma Aldrich, St.

Louis, MO), and 1.2 mM calcium chloride (EM Science, Fibbstown, New Jersey). The

buffer pH was adjusted to between 7.2 and 7.5 with 1 N hydrochloric acid or 1 N sodium

hydroxide solution (VWR, Radnor, PA).

Experimental Procedures

Preparation of Quantum Dot Dispersions

Commercially available QDs were purchased as 1 mg/mL dispersions in water.

These particles were shaken by hand for 1 min and an aliquot was pipetted and diluted to

0.05 mg/mL using KRB buffer and mixed well for an additional 1 min.

Determination of Particle Size and Zeta Potential

The mean diameter of the QDs (0.05 mg/mL) in KRB was determined using

dynamic light scattering (Nicomp Particle Sizer, Model 380 ZLS, Santa Barbara, CA)

using a volume-weighted distribution analysis, their surface zeta potentials were

measured using a Malvern Nano ZS Zetasizer (Worcestershire, UK).

Preparation of Bovine Respiratory and Olfactory Mucosal

Tissues

Bovine nasal mucosal tissues (olfactory and respiratory tissues) were obtained

from a local abattoir (Bud’s Custom Meats Co, Riverside, IA). The animals were

46

decapitated and longitudinal incisions along the lateral walls of the nasal cavity were

made along with a vertical incision along the ocular plane to expose the nasal respiratory

and olfactory regions. Both the nasal respiratory and olfactory tissues were harvested,

rinsed thoroughly with KRB and transported to the lab after placing them in fresh, ice-

cold KRB. Tissue sections were carefully peeled away from the underlying cartilage and

mounted between the donor and receiver chambers of a Navicyte® diffusion chamber

system. Tissues were mounted such that the mucosal side of the tissue faced the donor

solution and transport took place in the direction from the mucosal to the submucosal

surface. The explants were equilibrated for 20-30 min with 1 mL of prewarmed (37 ˚C)

KRB in both donor and receiver chambers. The tissues were kept aerated with carbogen

(95% O2 + 5% CO2), and the temperature was maintained at 37 ˚C throughout the

experiment.

Quantum Dot Uptake Studies

Methods to measure QD uptake using bovine nasal mucosal explants were

adapted and modified from previous studies reported by Kandimalla et al.132. After

equilibration of tissues with KRB, the buffer was replaced with 1 mL of QD suspension

(0.05 mg/mL) in the donor chamber and 1 mL of prewarmed KRB buffer in the receiver

chamber. At various time points (30, 60 and 120 min) tissues were removed from the

system and the donor and receiver solution were collected and stored at 4 ˚C for future

analysis. The mucosal surface of the tissues was washed with deionized water and the

washing fluids were also collected for analysis. The tissue sections exposed to the QD

suspension were carefully cut to isolate the tissue region exposed to the quantum dots and

the weight of the tissues was recorded using an analytical balance (Model: XS105DU,

47

Mettler Toledo, Columbus, OH). All the samples (tissues, donor, receiver, and tissue

washings) were stored at 4 ˚C until further analysis for QD content. The integrity of the

tissues was monitored by measuring transepithelial electrical resistance (TEER) of the

tissues before and after the completion of the transport study using an EVOM2 epithelial

voltohmmeter (World Precision Instruments, Sarasota, FL). Previous studies have shown

that bovine nasal tissue explants were viable for up to 4 h after harvesting132, and all

experiments were performed within this 4 h window. A typical range of TEER values

reported in literature is 120 -180 Ω cm2 for bovine nasal mucosa132. Initial studies

showed average TEER values for bovine respiratory mucosa and olfactory mucosa were

~ 180 Ω cm2 and ~140 Ω cm2 respectively. Any values less than 100 Ω cm2 were

indicative of tissue damage and tissue explants with resistances less than 100 Ω cm2 were

discarded.

The effect of hydrophilic surface modification of QDs was tested using PEG-

QDs. For these experiments, the particles and tissues were prepared in an identical

manner as for the COOH-QDs, but tissues were exposed to PEG-QDs (0.05 mg/mL) in

KRB for 2 hr. The extent of PEG-QD uptake was compared to that for COOH-QDs by

comparing the amounts of QDs in the donor and receiver chamber and within the tissue

segment exposed to the donor chamber containing the QD dispersion.

Quantification of Quantum Dots

The uptake of QDs into the nasal tissues was studied by quantifying the particle

mass transferred into the tissues as a function of time. QDs have fluorescent properties

and several studies have shown the use of fluorescent spectroscopy to quantify quantum

dots. However, initial attempts to quantify QDs with this technique were unsuccessful

48

because of the high variation within repeated samples. Also, many of the concentrations

used in these tissue uptake studies were below the quantification limit of fluorescent

spectroscopy. QD emission spectra are sensitive to the size of the particles and slight

changes in the particle size changes their emission spectra133. For accurate quantification

using fluorescent spectroscopy methods extraction of intact QDs from the tissue samples

was required. During the course of a transport study and during the extraction procedure,

however, the size of the QDs could change, which would result in either over or under

prediction of the true QD content134. In order to avoid these discrepancies, several

researchers used alternate methods for quantification of QDs that are independent of

particle size, including atomic spectroscopy, mass spectrometry, and inductively coupled

optical emission spectroscopy (ICP-OES). Unlike fluorescence spectroscopy, ICP-OES

quantifies the elemental content (Cd, Zn, and Se) of QDs from which the total amount of

QDs in the sample can be quantified. Due to the high sensitivity of this technique, ICP-

OES analysis of Cd was used to determine the amount of nanoparticles in the tissue

samples.

ICP-OES is a powerful and popular analytical tool for the quantification of

metals. A schematic of the working principle of ICP-OES is shown in Figure 3.1. An

inert gas, argon, continuously flows through a source of concentric quartz tubes called a

torch where the argon gas is ionized by a radio-frequency (RF) generator (0.5 to 2 kW).

49

Figure 3.1 Schematic showing the working principle of ICP-OES135. Metal-containing samples are nebulized using a peristaltic pump and an auxiliary argon flow directly into the plasma. The metal-containing droplets are atomized, ionized and finally exited to higher energy levels. The characteristic emissions from metal ions are separated using a high precision prism and a photomultiplier tube detector captures the intensity of each emission.

The resulting ions and electrons interact with a magnetic field produced by an

induction coil (placed around the torch) to accelerate the ions and electrons in specific

annular paths. The ions and electrons collide with additional argon to generate a high-

temperature (5000 – 10000 K), inductively-coupled plasma (ICP). Metal-containing

samples are nebulized using a peristaltic pump and an auxiliary argon flow directly into

the plasma. The metal-containing droplets are atomized, ionized and finally exited to

higher energy levels. When returning to their ground state, metallic ions emit photons

characteristic of each specific element. These emissions are separated using a high

50

precision prism and a photomultiplier tube detector captures the intensity of each

emission. The measured intensity of the emission is proportional to the concentration of

the element, and elemental concentrations can be calculated using a standard curve

generated using known concentration standards.

The accuracy and efficiency of ICP-OES depends on various parameters

including the RF power, the argon gas flow rate into the plasma, the peristaltic pump rate,

and the auxiliary argon flow rate. These parameters were optimized for the detection of

cadmium, which emits characteristic photons at a wavelength of 226.5 nm136. The

intensities of the corresponding photon wavelengths were measured to quantify the

cadmium content. After the method was developed, it was tested for linearity using a

series of elemental cadmium standards (Sigma Aldrich, St. Louis, MO) The experimental

plan was to measure the concentration of the Cd present in the quantum dots contained in

the digested samples and thereby determine the QD concentration using stoichiometric

calculations. A Varian ICP-OES 720 ES system (Agilent Technologies, Santa Clara, CA)

was used for these studies.

Extraction of Cadmium from QDs

Extraction of cadmium from the core of the quantum dots is necessary in order to

accurately quantify QDs using ICP-OES. Because of their inert nature, QDs exhibit

greater thermal and chemical stability both in solution state and in vivo conditions

compared to other polymeric nanoparticles137-139. The procedure to extract cadmium from

the quantum dots was adapted from a previous report using an acid digestion method134.

QD-containing dispersions were digested with 70 %w/w nitric acid in borosilicate culture

test tubes for about 24 h at 80 °C maintained using a heat block, followed by dilution

51

with deionized water. The efficiency of the digestion method in extracting Cd was

investigated by correlating the theoretical Cd concentration based on the initial QD

dispersion concentration from aliquots of the QD dispersion to the Cd concentration

measured following acid digestion of the dispersion aliquot.

Extraction of Cadmium from QDs in Bovine Respiratory

and Olfactory Tissues

It was previously reported that a variety of mouse tissues could be completely

digested using nitric acid134. Adaptation of the nitric acid digestion method to extract Cd

from QDs in bovine nasal tissues was carried out using the same method as described in

above section. Bovine respiratory and olfactory tissues exposed to QDs were treated with

70 %w/w nitric acid for about 24 h at 80 °C and diluted with water before analysis using

ICP-OES. Since cadmium is present in very low concentrations in tissues, there is the

possibility of interference from endogenous Cd. In order to test for the interference of the

endogenous Cd, control bovine nasal tissues (without QD exposure) were treated under

similar conditions and analyzed using ICP-OES.

To extract cadmium from the QDs dispersed in the donor, receiver and washing

fluids obtained during transport studies, 0.2 mL of a QD donor sample, or 0.5 mL of a

receiver sample, 0.5 mL of a washing fluid were placed in borosilicate glass test tubes

and 1 mL, 0.5 mL and 0.5 mL of 70% w/w nitric acid were added, respectively, to the

test tubes. Tissue explants exposed to the QD dispersion were weighed and placed in a

borosilicate test tube and 1 mL of 70%w/w nitric acid was added to the tube. All the test

tubes with nitric acid were placed in a 80 °C heating block for 24 h. Following Cd

extraction, the samples were diluted to 5 mL with deionized water and analyzed for

52

cadmium content using the Varian ICP-OES 720 ES system. Each set of samples

included a blank (no QDs), a set of elemental Cd standards of known concentrations, a

positive control (0.05 mg/mL QD dispersion in KRB) and a control tissue (tissue not

exposed to QDs) as control samples.

Visualization of QDs in Tissues Using Confocal

Microscopy

The unique, size-dependent fluorescent spectral properties of QDs make them

good alternatives to fluorescent dyes for visualization. Compared to conventional dyes,

QDs have improved photo-stability and have broad excitation spectra and narrow

emission spectra124. Following the QD transport studies, the tissue sections were

visualized for the localization of QDs within the tissues using confocal laser scanning

microscopy.

Following a QD transport study, donor and receiver solutions were collected, and

the mucosal side of the tissue was washed with deionized water. The tissue sections

exposed to QDs were carefully cut and placed into a fixative (4% paraformaldehyde) for

24-36 h followed by treatment with a series of increasing concentrations of sucrose (10%,

20% and 30% w/v). The tissues were snapfrozen using liquid nitrogen and stored at

-20 ˚C. Vertical cross-sections (10 µm) were cut using a cryostat microtome (Lecia

Microm HM 505 E Cryostat, Buffalo Grove, IL) at -25 ˚C and stained with DAPI, a blue-

fluorescent nuclear stain. A Zeiss 710 confocal laser-scanning microscope (Oberkochen,

Germany) was used with a UV laser (405 nm) and an argon ion laser (458 nm) to excite

the DAPI and QDs, respectively. Emission bands of 460 ± 40 nm and 620 ± 20 nm were

53

used to capture DAPI and QD fluorescence, respectively. The captured images were

processed using ImageJ software (Freeware from National Institute of Health, USA).

Visualization of QDs in Tissues Using Transmission

Electron Microscopy (TEM)

In addition to examining the tissues for fluorescence indicating QD uptake,

additional visualization of the QDs in the tissues was attempted using transmission

electron microscopy. Since confocal microscopy is limited by the wavelength of the

excitation laser (~460 nm in these studies), visualization of individual, extremely small

QDs within the cells is difficult. On the other hand, resolution can be increased

substantially using TEM, which is not limited by the wavelength range of light, and

instead, is dependent on the energy of the electron beam. TEM was utilized to obtain

high-resolution images of electron-dense QDs within the cells140.

Similar to the confocal studies, following the completion of the transport studies,

the tissues mounted between the diffusion cells were washed with deionized water and

trimmed to the exposure area between the diffusion cells and further sectioned into

~1-2 mm3 sections which were chemically fixed with 2.5% glutarldehyde for ~ 24 h. The

tissues were rinsed with 0.2 M sodium glycocholate and treated with fresh 1 % OsO4

(prepared by adding 2 parts of 0.2 M sodium glycocholate to 1 part of 4% osmium and 1

part of 6% potassium ferocyanide) followed by treatment with 2.5% uranyl acetate. After

30 min, the tissues were dehydrated with a series of increasing acetone concentrations

(50%, 75%, 95% and 100%). Following complete dehydration, the tissues were placed in

epoxy resin (Epon-812TM, Sigma Aldrich, St Louis, MO) and molded at 70 ˚C for at least

2-3 days. Ultrathin sections (~ 80 nm) were prepared using an ultramicrotome (Leica EM

54

UC6 Ultramicrotome Mz6, Buffalo Grove, IL) and placed on a grid for TEM analysis. A

JEOL JEM-1230 TEM (Tokyo, Japan) was used to obtain images of the QDs in the tissue

sections.

Investigation of the Endocytic Pathways Involved in the

Uptake of Quantum Dots

Particulate matter is most commonly taken up into cells using energy-dependent

endocytic pathways141-142. However, due to the small sizes of the quantum dots, these

particles may be able to enter the tissues through energy-independent processes,

including transport through the paracellular spaces41, 71. The extent of passive QD uptake

was investigated by blocking all energy-dependent endocytic pathways using 0.18 µg/mL

2, 4-dinitrophenol (2,4-DNP) (Sigma Aldrich, St Louis, MO). The involvement of

specific endocytic pathways was identified by using chemicals reported to be specific

inhibitors for each pathway. The pharmacological inhibitors chlorpromazine143 (CPZ, 10

mg/mL), amiloride144 (10 µg/mL) and methyl-β-cyclodextrin145 (MBC, 5 mg/mL) have

been shown to inhibit the clathrin-mediated endocytosis, macropinocytosis and caveolae-

mediated endocytosis, respectively.

The inhibitor of interest was dissolved in KRB and after tissues were equilibrated

for 30 min with KRB, 1 mL of inhibitor in KRB was replaced in the donor and receiver

chambers for 60 min. This step allows the inhibitors to permeate into the tissues prior to

QD exposure. The donor solution was then replaced with 1 mL of a QD suspension (0.05

mg/mL) containing the inhibitor in KRB and the receiver chamber was replaced with a

fresh 1 mL of the KRB with inhibitor. Transport studies were carried out for an additional

55

60 min, and tissue samples along with donor, receiver and washing fluids were collected

and analyzed for QD content using the method described in previous section.

Statistical Analysis

Each experiment was repeated at least 3 to 6 times with tissues obtained from

different animals and the data are presented as mean ± standard deviation. Statistical

significance was tested using either one-way or two-way analysis of variance (ANOVA),

where appropriate. Multiple comparisons were conducted using the uncorrected Fischer’s

LSD test. A Student’s t-test was used when comparing two sample sets. Differences were

considered significant at p < 0.05. GraphPad Prism, Inc. (La Jolla, CA) software was

used to perform the statistical testing.

Results

Particle Size Analysis

Three lots of COOH-QDs and one lot of PEG-QDs of ~ 7nm size were purchased

as 1 mg/mL dispersions in water. Both COOH- QDs and PEG-QDs were observed to

show a bimodal size distribution with the majority (> 99.5 %) of particles of < 20 nm and

a small fraction (~0.1 - 0.4%) of larger size (~800 nm) (See Figure B.1 in Appendix-B).

These larger particles may be minor contaminants, most likely dust particles. Purchased

dispersions were diluted in KRB to a concentration of 0.05 mg/mL and the particle size

was measured (Table 3-1). The viscosity and refractive index values of the QD

dispersions used for particle size analysis were 0.933 cPs and 1.333, respectively. These

values were selected based on the assumption that the viscosity and refractive index of

the QD dispersions was similar to water at 25 °C. The larger particle sizes measured,

56

compared to the manufacturer’s specifications, may be the result of either not correcting

for the increased viscosity or altered refractive index of the QD dispersions. Aggregation

of the particles is also a possibility, but patterns typical of aggregate formation (multi-

modal size distribution) are not noted with the observed particle size distributions. Zeta

potential measurements revealed that the surfaces of the carboxyl-QDs and PEG-QDs

were negatively charged. Since PEG is a hydrophilic, neutral polymer, an increase in the

surface charge towards neutrality was observed when compared to the anionic COOH-

QDs.

Table 3-1 Quantum dot (~ 7nm) particle size distribution (n=3, mean ± standard deviation) and surface charge for 0.05 mg/mL samples in KRB.

Surface Group Lot Mean Particle

Diameter (nm) Zeta potential (mV)

COOH 1 ( LW074414A21104) 11.4 ± 0.4 -18.05 ± 2.05

COOH 2 (Not available) 12.8 ± 1.7 Did not test

COOH 3 (Not available) 14.5 ± 2.1 Did not test

PEG 1 (LW134414A23108) 14.6 ± 0.3 -9.23 ± 1.18

Quantification of Quantum Dots

The developed ICP-OES instrument operational conditions and measurement

parameters for quantification of Cd are provided in Table 3-2. The ICP-OES method was

calibrated using certified Cd elemental standard solutions (Sigma Aldrich, St. Louis, MO)

and the method showed good linearity over a range of 10 ng/mL to 1000 ng/mL of Cd (r2

> 0.999) (Figure 3.2). The lowest calibration concentration was 10 ng/mL and this was

taken as the quantification limit; any values below 10 ng/mL showed high variation and

were considered as zero values in the data analysis.

57

Table 3-2 Operating conditions and measurement parameters of Varian ICP-OES 720 ES.

Figure 3.2 Sample calibration curve for elemental Cd using ICP-OES (n=3). Calibration equation was Intensity (a.u) = 3.4984 * Cd Conc. (ng/mL), r2 = 0.9999.

0

1000

2000

3000

4000

5000

0 200 400 600 800 1000 1200

Inte

nsi

ty U

nit

s (a

.u.)

Concentration of Cd (ng/mL)

Sample Introduction

Auxiliary Argon Flow (L/min) 1.5

Sample Uptake (s) 30

Rinse Time (s) 10

Pump Rate (rpm) 15

Nebulizer Gas Flow (L/min) 0.75

Replicates 3

Plasma Properties

RF power (kW) 1.00

Plasma Gas Flow (L/min) 15.0

Cadmium (Cd) emission line (nm) 226.5136

58

The extraction of Cd from the core of the QDs was shown to be achievable with

digestion in nitric acid. Digestion of QDs (0.143 μg to 143 μg) with 1 mL of 70 %w/w

nitric acid for 24 h at 80 °C was shown to be sufficient to extract all the Cd from the core

of the QDs. Calculation of the QD concentrations from the Cd content in the digested

samples showed a good correlation to the theoretical concentrations of QDs with a good

correlation (r2>0.999) (Figure 3.3).

Figure 3.3 Correlation between theoretical QD concentrations versus ICP-OES measured QD concentration. A correlation equation of Intensity (a.u) = 0.9697 * Cd. Conc (ng/mL) was observed (r2 =0.9992).

The same acid digestion conditions used to extract Cd from QDs in KRB were

used to extract Cd from QDs present in nasal tissues. The average weight of the bovine

respiratory and olfactory tissues exposed to the QDs was 110 ± 24 μg and 80 ± 24 μg,

respectively. When representative bovine respiratory and olfactory tissues were exposed

0.00

0.02

0.04

0.06

0.08

0.10

0.12

0 0.02 0.04 0.06 0.08 0.1 0.12

Ca

lcu

late

d c

on

c o

f Q

D (

mg

/m

L)

Theoretical conc of QD (mg/mL)

59

to blank KRB for 2 h in the Navicyte® diffusion apparatus, followed by digestion in 1 mL

of 70 %w/w nitric acid at 80 °C, within 24 h a clear, faint yellowish color solution was

observed without any visible particulate content. This suggested that both respiratory and

olfactory tissues were completely digested. To investigate the endogenous Cd content in

the blank tissues, digested tissue samples were analyzed for Cd content with ICP-OES

and a negligible amount of endogenous Cd was observed.

To test if QDs present in the tissues can be digested completely using the acid

digestion conditions, known concentrations of QDs (data provided in Appendix-C) were

spiked with blank tissue samples and digested in nitric acid for 24 h at 80 °C followed by

analysis of samples using ICP-OES. A good correlation between the measured Cd

concentration and the theoretical concentration of added QDs was obtained. From Figure

3.4 and 3.5 it can be observed that the measured mass of QDs per gram of bovine nasal

respiratory and olfactory tissues correlated well (r2>0.99) with the theoretical loaded

mass of QDs. A sample calculation is shown in Appendix-D.

Figure 3.4 Correlation of the theoretical mass of Cd as added QD dispersion to blank respiratory tissues and the Cd concentration measured from digested samples of these tissues. A good correlation between added mass of QDs and measured mass of QDs was observed (y=0.997x, r2=0.999).

0

35

70

105

140

175

210

0 50 100 150 200

Me

asu

red

Ma

ss o

f Q

Dp

er

g o

f R

esp

ira

tory

Tis

sue

(μg/g

)

Calculated Mass of QD per g of Respiratory Tissue …

60

Figure 3.5 Correlation of the theoretical mass of Cd as added QDs to blank olfactory tissues and the Cd concentration measured from digested samples of these tissues. A good correlation between added mass of QDs and measured mass of QDs was observed (y=996x, r2=0.999).

From these studies it was observed that the developed acid digestion conditions

were sufficient to digest the bovine nasal tissues and to extract the Cd completely from

the core of the QDs. The lowest detectable concentration of QDs using the developed

ICP-OES method was found to be 0.143 μg/mL, which corresponds to 10 ng/mL of Cd.

QD Translocation into Nasal Respiratory and Olfactory

Mucosa

Following 120 min incubations of QD dispersions with respiratory and olfactory

tissues in Navicyte® diffusion cells, the uptake of COOH-QDs by the olfactory mucosa

was found to be ~2.5-fold higher compared to the respiratory mucosa (Figure 3.6).

0

75

150

225

300

375

450

0 75 150 225 300 375

Me

asu

red

Ma

ss o

f Q

Dp

er

g o

f O

lfa

cto

ryT

issu

e(μg/g

)

Calculated Mass of QD per g of Olfactory Tissue

(μg/g)

61

T im e (m in )

Am

ou

nt o

f Q

Ds

pe

r

gra

m o

f t

iss

ue

(u

g/g

)

30

60

120

0

5 0

1 0 0

1 5 0

R e s p ira to ry

O lfa c to ry

Figure 3.6 Comparison of QD uptake into full thickness olfactory and respiratory tissues after a 120 min incubation period. A) Column graph showing the mean and standard deviation of the groups Uncorrected Fischer’s LSD test showed significant difference (p<0.05) in uptake of QD between respiratory and olfactory tissues after 120 min incubation B) Box Whisker plots of the same data showing the median and range of the data. (n=3).

30 60 120

0

10

20

30

40

Time (min)

Am

ou

nt

of

QD

s p

er

gra

m o

f ti

ssu

e (

mg

/g)

Respiratory Tissue

Olfactory Tissue

*

A

B

62

The transfer of QDs (expressed as the percent of the original donor concentration

in Table 3-3) into the olfactory mucosa was increased from 1.1 % in 30 min to 4.4 % in

120 min. These findings suggest entrapment of particles within nasal tissues as soon as

30 min of exposure. Transfer into the respiratory mucosa remained low (~ 0.7 – 1.8 %)

throughout the incubation period. The uptake of QDs into respiratory and olfactory

tissues after 60 min exposure was found to be similar (~ 1.2 %), however exposure for

longer times (120 min) resulted in greater uptake into olfactory tissue (~ 4.4 %)

compared to respiratory tissue (~1.8 %). Approximately 1.2 – 5 % of the QDs were

recovered from the tissue washings which implies some QDs were adsorbed to the tissue

surface. Maximum adsorption of QDs was observed with respiratory tissues (~ 5%)

exposed for 120 min. The recovery of QDs (in µg) from the donor chambers, receiver

chambers, tissue washing fluids and tissues at all time intervals is summarized in Table 3-

4. Most of the QDs remained in the donor chamber, and a negligible mass of the QDs (<

0.11 µg) was recovered from the receiver chamber and tissue washings. The localization

within tissues was further studied using microscopy techniques described in following

sections. The overall recovery of QDs following transport studies was found to be greater

than 79 %.

63

Table 3-3 Measurement of the percent transport (relative to the donor QD loading) of quantum dots across bovine respiratory and olfactory mucosal explants. Experiments were initiated by placing 1 mL of QD dispersion containing approximately 44.5 ± 3.9 µg of QD in the chamber facing the mucosal surface of the tissue. Incubations of 30, 60 and 120 min were conducted and 3 tissues were evaluated at every time period. Recovery of QDs in donor chamber, receiver chamber, mucosal tissue, and from tissue washings following the transport studies are provided as percentage of the initial dose. The values are given as mean (n=3) ± (standard deviation).

Sample Time (min) Weight of tissue (g) Tissue (%) Donor (%) Receiver (%) Washings (%)

Respiratory Tissue

30 0.1140 (0.021) 0.7 (0.6) 84.4 (7.4) 0.2 (0.0) 1.2 (1.0)

60 0.0999 (0.009) 1.3 (0.5) 85.1 (7.0) 0.2 (0.1) 1.7 (0.6)

120 0.1160 (0.039) 1.8 (0.9) 82.3 (7.6) 0.2 (0.1) 4.9 (4.7)

Olfactory Tissue

30 0.0627 (0.020) 1.2 (0.4) 81.2 (2.1) 0.2 (0.1) 1.1 (0.3)

60 0.0807 (0.017) 1.9 (0.9) 81.2 (2.9) 0.2 (0.1) 1.6 (0.8)

120 0.0954 (0.025) 4.4 (1.7) 73.6 (9.4) 0.1 (0.1) 1.1 (0.3)

64

Table 3-4 Measurement of the transport of quantum dots across bovine respiratory and olfactory mucosal explants. Experiments were initiated by placing 1 mL of QD dispersion containing approximately 44.5 ± 3.9 µg of QD in the chamber facing the mucosal surface of the tissue. Incubations of 30, 60 and 120 min were conducted and 3 tissues were evaluated at every time period. Recovery of QDs in donor chamber, receiver chamber, mucosal tissue, and from tissue washings following the transport studies are provided in the table. The values are given as mean (n=3) ± (standard deviation).

Sample Time (min)

Weight of tissue (g)

Amount in tissue (µg)

Amount in Donor (µg)

Amount in Receiver (µg)

Amount in Washings

(µg)

Total Mass of QDs recovered

(µg)

Total Recovery

(%)

Respiratory

Mucosa

30 0.1140 (0.021) 0.29 (0.2) 37.39 (1.3) 0.10 (0.0) 0.54 (0.4) 38.05 (1.3) 86.6 (8.2)

60 0.0999 (0.009) 0.54 (0.2) 37.70 (1.5) 0.10 (0.0) 0.78 (03) 38.74 (1.8) 88.2 (6.5)

120 0.1160 (0.039) 0.75 (0.4) 36.41 (1.0) 0.07 (0.1) 2.16 (1.9) 38.30 (2.4) 89.0 (11.7)

Olfactory

Mucosa

30 0.0627 (0.020) 0.49 (0.2) 36.09 (2.7) 0.11 (0.0) 0.48 (0.1) 36.93 (2.4) 83.7 (2.6)

60 0.0807 (0.017) 0.80 (0.4) 36.19 (4.2) 0.08 (0.1) 0.73 (0.4) 37.42 (4.1) 84.8 (2.6)

120 0.0954 (0.025) 1.87 (0.7) 32.52 (1.7) 0.07 (0.0) 0.48 (0.2) 34.69 (2.3) 79.1 (10.9)

65

The effect of surface properties of QDs on the extent of uptake into nasal mucosa

was studied by comparing the uptake of PEG-QDs and COOH-QDs. A 2 h exposure of

bovine nasal mucosa to PEG-QDs at the same concentration used for the COOH-QDs

(0.05 mg/mL) and at twice the concentration (0.1 mg/mL), did not show any uptake of

the PEG-QDs into either respiratory or olfactory tissues. This is significantly different

than the COOH-QDs which showed ~1 % to 5 % uptake into bovine nasal tissues (Table

3-3). More than 90% of the PEG-QDs remained in the donor chamber and ~5% were

recovered from the tissue washings. Since PEG-QDs did not show any uptake into the

nasal tissues, only COOH-QD uptake using imaging techniques was further pursued.

Visualization of QDs in Tissues Using Confocal and

Electron Microscopy

The fluorescent images of the respiratory and the olfactory tissues following

exposure to QDs are shown in Figures 3.7 and 3.8, respectively. The blue color in the

images shows the position of the nucleus in each cell stained with DAPI, and a clear

distinction between the intact epithelium (showed with green color lines in the figures)

and the submucosal region of the nasal tissues following exposure to QDs shows that the

integrity of the tissue remained unaffected for the entire incubation period. As additional

tissue viability evidence, the TEER values, which are a measure of the tissue integrity,

remained within an acceptable range for the entire period of the incubation (See

Appendix-E). These observations show that the transfer of the QDs into the tissue is not

simply a result of epithelial barrier disruption.

66

Figure 3.7 Confocal laser scanning microscopic images of respiratory tissues showing the

transport of QDs. Column I shows the nuclear stain (DAPI) channel. The

epithelial region is indicated by a solid line and the submucosal region by a

double arrowed line. Column II shows the QD channel, and column III shows

merged images from both channels. Each row of images is labeled with the

exposure time of the respiratory tissues to QDs or to a control samples with no

QD exposure. White arrows highlight the QD localization in the merged

image of the respiratory tissue. (Scale bar = 20 µm).

I II III

Control

no QD

exposure

30 min

60min

120 min

67

Figure 3.8 Confocal laser scanning microscopic images of olfactory tissues showing the transport of QDs. Column I shows the nuclear stain (DAPI) channel. The epithelial region is indicated by a solid line and the submucosal region by a double arrowed line. Column II shows the QD channel, and column III shows merged images from both channels. Each row of images is labeled with the exposure time of the olfactory tissues to QDs or to a control samples with no QD exposure. White arrows highlight the QD localization in the merged image of the olfactory tissue. (Scale bar = 20 µm).

I II III

Control no

QD

exposure

30 min

60min

120 min

68

The fluorescent signal intensity corresponding to the QDs (red color, Figures 3.7

and 3.8) in the epithelium and submucosal region of both the tissues gradually increased

over the 30 min to 120 min measurement intervals, demonstrating that the QDs were

continuously translocated into the respiratory and the olfactory tissues. As early as

30 min after incubation, QDs were visually observed in the submucosal regions of both

the respiratory and olfactory mucosa. The increased fluorescence intensity in the

submucosal region between 60 min and 120 min compared to the early 30 min incubation

shows that the QDs accumulate in these regions.

Transmission Electron Microscopy (TEM)

The TEM images displayed in Figures 3.9a & 3.10a show epithelial cell images

from respiratory and the olfactory samples exposed to quantum dots for 120 min. QDs

were observed in the intercellular spaces between the epithelial cells. At higher

magnification (Figures 3.9b & 3.10b) showing portions of two epithelial cells and their

associated intercellular junctions revealed the presence of QDs in various endocytic

vesicular structures with variety of morphologies. QDs were also present as small

aggregates in the cytoplasm. These results suggest that the uptake of QDs is primarily by

endocytic pathways, with some contribution from paracellular transfer.

69

Figure 3.9 Transmission electron micrographs of bovine respiratory epithelial cells exposed to COOH-QDs for 120 min. Distinct nucleus (N), mitochondria (M), cellular junction (CJ), golgi apparatus (G) can be observed. a: magnification x7000. b: Enlarged mucosal region (orange circle) showing dispersed, electron-dense particles in cytoplasm (green circles) and in vesicle structures (red circles) of the epithelial region (magnification x21000).

70

Figure 3.10 Transmission electron micrographs of bovine olfactory epithelial cells exposed toCOOH-QDs for 120 min. Distinct nucleus (N), mitochondria (M), cellular junction (CJ) can be observed. a: Image with magnification of x7000. b: Enlarged mucosal region (orange box) showing dispersed, electron-dense particles in cytoplasm (green circles) and vesicular structures (red circles) in the epithelial region(magnification: x24000).

71

Identification of Endocytic Pathways

Assessment of QD tissue uptake in the presence of inhibitors of endocytosis indicated

that the uptake of QDs into the nasal mucosa involves multiple pathways. Incubation with

2, 4-DNP a general inhibitor of ATP-dependent activities, resulted in a significant

reduction in the uptake of COOH-QDs into respiratory tissues (Figure 3.11). Similarly, in

the presence of specific endocytic inhibitors, either amiloride, CPZ or MBC, a reduction

in COOH-QD uptake was also observed. These findings suggest that the primary uptake

pathways of QDs in the respiratory tissue involve macropinocytosis, along with clathrin-

mediated and caveolae-mediated endocytic pathways. Energy-independent pathways in

the respiratory tissue do not appear to play major role in the uptake of ultrafine

nanoparticles. However, extremely small size QDs may have access to energy-

independent pathways through the intercellular spaces that are evident in the TEM

images (Figures 3.9a & 3.9b).

In the case of olfactory tissues (Figure 3.12), only chlorpromazine was able to

significantly reduce the uptake of the COOH-QDs, suggesting that the major endocytic

uptake pathway for these ultrafine particles in the olfactory mucosa might be via clathrin-

mediated endocytosis. The inclusion of any other inhibitor, including 2, 4-DNP, did not

reduce the uptake to a statistically significant level, which implies that energy-

independent pathways also exist for the uptake of quantum dots into the olfactory tissues.

TEM images (Figures 3.10a & 3.10b) show the presence of QDs in the intercellular

spaces between the olfactory epithelial cells, providing additional evidence for the

importance of these pathways in the olfactory mucosa.

72

A

B

Co

ntr

ol

Wit

h 2

,4 D

NP

Wit

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milo

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Wit

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BC

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PZ

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2

4

6

8

Am

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QD

s p

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ra

m

of

tis

su

e (

g/g

)

Figure 3.11 Uptake of QDs in the nasal respiratory tissue in the presence of inhibitors: 2,4-dinitrophenol (DNP), amiloride, methyl-β-cyclodextrin (MBC) and chlorpromazine (CPZ). A) Bar graph showing mean and standard deviation, * indicates significant difference between the control and inhibited uptake compared used Student’s t-test, n=3 or 6, p<0.05. B) Box Whisker plot showing the median and range of the same data.

Con

trol

With

2,4

DN

P

With

Am

ilori

de

With

MBC

With

CPZ

0

2

4

6

8A

mou

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QD

s p

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ram

of

tiss

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(µg/g

)

*

*

* *

73

A

B

Co

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ol

Wit

h 2

,4 D

NP

Wit

h A

milo

r id

e

Wit

h M

BC

Wit

h C

PZ

0

5

1 0

1 5

2 0

Am

ou

nt

of

QD

s p

er g

ra

m

of

tis

su

e (

g/g

)

Figure 3.12 Uptake of QDs in the nasal olfactory tissue in the presence of inhibitors: 2,4-dinitrophenol (DNP), amiloride, methyl-β-cyclodextrin (MBC) and chlorpromazine (CPZ).A) Bar graph showing mean and standard deviation, * indicates significant difference between the control and inhibited uptake compared used Student’s t-test, n=3 or 6, p<0.05. B) Box Whisker plot showing the median and range of the same data.

Con

trol

With

2,4

DN

P

With

Am

ilori

de

With

MBC

With

CPZ

0

5

10

15

20

Am

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s p

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tiss

ue

(µg/g

)

*

74

Discussion

Although the science of nanotechnology has advanced immensely in recent years,

the transport properties of nanoparticles into and across tissues, and specifically the nasal

mucosa are still not well understood. In these studies the uptake of quantum dots, a

model for ultrafine nanoparticles, into or across the bovine nasal mucosa was evaluated.

While it is unlikely that QDs will be effective drug delivery systems due to their heavy

metal contents, their unique, size-specific optical properties have encouraged their

investigation in a number of different applications in the biomedical and pharmaceutical

sciences124.

After 2 h of QD exposure to nasal tissues, the majority of the COOH-QDs were

recovered from the donor chamber with more limited accumulation in nasal tissues and

only negligible amounts of the QDs were transported through the whole thickness of the

tissues and entered the receiver chamber. Visualization of the COOH-QDs using confocal

microscopy also showed an accumulation of aggregates of QDs in the tissues. Unlike

small drug molecules, quantum dots are particles that are likely to be entrapped inside

vesicles in subcellular regions. They also appear to be trapped in the collagen fibers in the

submucosal regions of the nasal tissues, and this entrapment likely hinders the transfer of

QDs into the receiver side of the diffusion apparatus.

The red fluorescence signal intensity corresponding to the QDs observed in

epithelial and submucosal regions (Figures 3.7 and 3.8) suggest that a greater number of

particles accumulated in the submucosal region. These results are somewhat consistent

with previous report by Garzotto et al., who showed similar accumulations of carboxyl

surface modified QDs in the submucosal region, surrounding the blood vessels and nerve

75

bundles of the olfactory epithelium after 4 h of intranasal administration in CD1 mice71.

In a different study, Chen et al., showed accumulation of 20 nm and 100 nm carboxylate-

modified polystyrene particles in the submucosal regions of bovine olfactory and

respiratory mucosa after only 60 min of exposure38. Contrary to these findings, however,

in a recent study, Mistry et al. reported that, even after 90 min exposure of either 20 nm,

100 nm or 200 nm carboxylate - modified polystyrene particles to excised porcine nasal

mucosa. The majority of the particles either remained in the donor chamber or associated

with the apical edge of olfactory epithelium and none of the particles, irrespective of size,

showed accumulation in the submucosal region when visualized using fluorescence

microscopy40. These conflicting results could be due to use of different ex vivo tissue

models that may show different rate of particle uptake.

Similarly, there are reports showing transfer of 20 – 1000 nm nanoparticles into

systemic circulation following nasal exposure37, 146. Most of the current and previous

reports lead to the same conclusions that ultrafine nanoparticles accumulate in the

submucosal region of nasal tissues. Accumulation of nanoparticles in the nasal tissues

can be advantageous for drug and vaccine delivery. The submucosal region of the nasal

tissues is highly vascularized, innervated with olfactory neuronal cells (olfactory tissue)

and also contains lymphoid tissues. Drug and/or antigen encapsulated nanoparticles

accumulated in this region may release the encapsulated drug/vaccine into both lymph or

blood circulation and provide the desired therapeutic effect.

Since the olfactory mucosa is highly innervated with olfactory neuronal cells, any

accumulation of particles near these cells may enhance the potential to transfer the

nanoparticles to brain via olfactory neuronal pathways or through perineuronal spaces

76

along the neuronal axons. Some earlier studies showed that colloidal particles of

elemental metals including iron oxide82, manganese72, and gold73 reach the brain in small,

but potentially toxic quantities following nasal exposure.

Unlike COOH-QDs, when PEG-QDs of similar size were exposed to the bovine

nasal tissues, uptake into these tissues was considerably reduced compared to the COOH-

QDs. It is not surprising that PEG-QDs were not internalized into the cells, since the

more neutral, non-interactive PEG surface hinders interactions with proteins, other

macromolecules, and cell-based receptors in the mucosa. Particle interactions with

components in the nasal secretions or at the cellular surfaces lead to in-situ surface

modifications that may promote uptake by a variety of endocytic pathways. However,

PEG, being a large, flexible neutral polymer is believed to camouflage the surface of the

nanoparticle which decreases the particle interactions with the components of the

secretions and cell surfaces, and thus limits uptake via the endocytic pathways147.

Similar observations have been reported by other investigators148-149 , and it has been

repeatedly shown that surface chemistry plays an important role in cellular

internalization.

Researchers have previously shown that ultrafine nanoparticulate matter is

typically taken up by three major endocytosis processes45, 150: clathrin-mediated

endocytosis, caveolae-mediated endocytosis and macropinocytosis. Several studies report

additional uptake mechanisms for QDs, including those with carboxylated surface

characteristics, in other tissues. Zhang et al. showed that quantum dots (655 nm emission

QDs) surface modified with carboxyl groups were taken up into human epidermal

keratinocytes by caveolae/lipid raft- mediated endocytosis involving LDL receptors and

77

scavenger receptors145. Xiao et al. used similar, 655 nm emission QDs with carboxyl-

derived surfaces, and showed that the mechanism of uptake into human mammary cells

(MCF-7 and MCF-10A cells) was through clathrin-mediated endocytosis151. The results

from nasal mucosal uptake studies show that the COOH-QDs were internalized in the

respiratory tissues via multiple endocytosis pathways including clathrin-mediated,

caveolae-mediated endocytosis and macropinocytosis. QDs exposed to bovine olfactory

mucosa, in comparsion, were internalized primarily by clathrin-mediated endocytosis. It

is not uncommon for particles to be taken up via multiple pathways, for example in a

recent study reported by Jiang et al.152 the involvement of both clathrin-mediated

endocytosis and macropinocytosis was observed in the uptake of D- penicillamine coated

quantum dots (8 nm diameter) into HeLa Cells.

Currently, proposed mechanisms for nanoparticle uptake include the adsorption of

proteins/ligands from the extracellular environment on the nanoparticle surface that serve

as triggers for specific endocytic processes. Macropinocytosis has not been previously

reported as an uptake mechanism for carboxyl-surface quantum dots. However,

macropinocytosis is a non-specific endocytosis process in which cell membrane ruffling

causes formation of large troughs, which can subsequently form large size vesicles

(1-5 μm diameter) taken into the cells and it is very likely that QDs could be included in

the materials entrapped within these vesicles153.QD uptake by bovine respiratory tissues

was observed to involve macropinocytosis, but this pathway did not seem to play a

significant role in QD uptake into olfactory tissues. Similar results were observed in

previous studies with polystyrene nanoparticles38, 56. Macropinocytosis also seemed to

play less significant role in uptake of 40 nm and 100 nm carboxylate polystyrene particles

78

from bovine olfactory mucosa, whereas this pathway played a major role in uptake of

these nanoparticles in bovine respiratory mucosa38, 56. Though macropinocytosis is a non-

specific endocytic pathway, it is associated with actin-dependent ruffling of the plasma

membrane. Factors that increase actin polymerization have been shown to elevate

macropinocytosis154. While it is not clear that the absence of these factors in bovine

olfactory mucosa is responsible for the minor involvement of macropinocytosis in uptake

of QDs in olfactory mucosa compared to the respiratory mucosa, it may be one direction

of future investigations. Also, it was shown that antigen-presenting cells like dendritic

cells and macrophages of the immune system internalize solutes and antigens using

macropinocytosis144. The absence of NALT in the olfactory tissue compared to

respiratory tissue could be another reason for lower involvement of macropinocytosis in

the olfactory mucosa.

An interesting observation from the present study is that although multiple

endocytic processes were involved in QD uptake into the respiratory tissue, only ~ 2 % of

the exposed QD load was taken up into the tissue. In comparison, ~ 5% of the QD load

was internalized into the olfactory tissues, primarily via clathrin-mediated endocytosis. A

potential explanation for the increased uptake into the olfactory tissues may be due to the

involvement of additional non-energy dependent mechanisms, including greater

utilization of paracellular routes in olfactory tissues compared to respiratory tissues. The

structural differences between respiratory and olfactory mucosa, including the presence

of Bowman’s glands in the olfactory epithelium may make olfactory mucosa more

permeable compared to respiratory mucosa. This is also evident from the lower TEER

values measured in the olfactory mucosa (~140 Ω cm2) compared to respiratory mucosa

79

(180 Ω cm2). Surprisingly, PEG-QDs did not show any uptake into the olfactory tissues,

which calls into question the role of the passive intercellular pathways in the uptake of

nanoparticles by the olfactory tissue.

Conclusion

The uptake behavior of ultrafine nanoparticles (<20 nm) was demonstrated to be

dependent on the region of the nasal epithelium involved (respiratory vs. olfactory) where

the extent of uptake was observed to be greater in the olfactory epithelium compared to

the respiratory epithelium. While both respiratory and olfactory tissues seem to be

morphologically similar, the uptake pathways utilized by these tissues were found to be

different. In respiratory tissues, clathrin-dependent, macropinocytosis and caveolae-

dependent endocytosis process were all involved in the uptake of QDs. Whereas in

olfactory tissues clathrin-dependent endocytosis was the major endocytic pathway

involved in uptake of QDs. Additional energy-independent pathways appeared to be

active in the internalization of QDs into the olfactory mucosa, however the effect of

surface chemistry on these pathways requires further investigation. The significantly

higher uptake of ultrafine nanoparticles by the olfactory mucosa might be advantageous

for the delivery of therapeutic materials into the CNS, but this also suggests that there is

an increased risk to the CNS from the transfer of airborne nanoparticulate substances

deposited on olfactory region.

80

CHAPTER IV

DISTRIBUTION OF QUANTUM DOTS AFTER INTRANASAL

ADMINISTRATION IN MICE IN VIVO LIVE ANIMAL IMAGING

Introduction

A number of reports of research conducted in animals have demonstrated

improved delivery of therapeutic agents to brain from the nasal cavity when delivered via

particulate systems155-157. For example, Al-Ghananeem et al. reported higher brain

concentrations of diadanosine in rats after intranasal administration of diadanosine-

carrying chitosan microparticles (269 – 382 nm) compared to concentrations achieved

after intravenous or intranasal administration of diadanosine solution158. There are also

reports showing potential transfer of extremely small, non-therapeutic nanomaterials to

the brain following nasal exposure in animal models70, 72-73, 82, 159. For example, Elder et

al. reported a 3.5 fold increase in manganese levels in the olfactory bulb after a 12 day

inhalation exposure to 30 nm manganese oxide nanoparticles72. The direct nose-to-brain

delivery via olfactory and trigeminal neuronal pathways and convective transport through

perineuronal spaces along olfactory axon bundles are believed to assist in the transfer of

substances into the brain while bypassing the blood- brain barrier30, 71. It is likely that

ultrafine small nanoparticles, those with diameter less than 100 nm, can both be

internalized into epithelial cells and transported via neuronal pathways to reach the brain.

There are also reports showing the transfer of inert particles/microparticles from

the nasal cavity into the systemic circulation. Almeida et al. reported low levels (0.96 %

of administered dose) of 510 nm carboxylated polystyrene particles into the circulatory

system of rats within 10 min following intranasal administration160. In another study,

81

Alpar et al. also reported evidence of 830 nm carboxylated polystyrene microparticles in

blood of rats even after 24 h of intranasal dosing146. The highly perfused respiratory

mucosa and the presence of nasal associated lymphoid tissue may provide pathways for

the subsequent transfer of internalized particles to systemic circulation29. However, it is

difficult to conceive of such large microparticles passing through the small fenestrated

capillary openings (diameter ~13-17 nm161) in the nasal mucosa, thus uptake into the

lymphatic system by scavenger cells seems to be the more likely method for larger

nanoparticle/microparticle transfer into the systemic circulation.

In Chapter 3, it was shown that carboxylate surface modified quantum dots (7 –

10 nm) are taken up into the nasal mucosa and lodge within these tissues. Understanding

the fate and biodistribution of the nanoparticles accumulated within these tissues is

essential for the development of efficient targeted delivery systems. The purpose of these

experiments was to study the distribution of nanoparticulate matter from the nasal

mucosa in live animals using fluorescent imaging and computed tomography techniques.

Quantum dots with carboxyl surface modifications (COOH-QDs) were used as model

nanoparticulate systems. While it is unlikely that quantum dots will be effective delivery

systems due to their heavy metal contents, they are an excellent model for initial

investigations to characterize nanoparticle biodistribution patterns without the need for

further bioconjugation with fluorescent dyes. Since QDs are also available with a variety

of surface modifications, the effect of surface properties on the biodistribution from the

nasal cavity can also be studied. In the current study, the biodistribution of PEG-coated

QDs (PEG-QDs) was compared to the distribution of carboxylated QDs to probe some of

the initial, potentially surface-dependent, uptake mechanisms of these particles. An

82

attempt to visualize the uptake of fluorescent quantum dots from the nasal cavity in

presence of inhibitors of endocytosis mechanisms was the focus of these initial

investigations.

The ability to quantitatively or semi-quantitatively study the biodistribution of

intranasally administered nanoparticles is a difficult process potentially involving

invasive techniques enabling the measurement of the resulting particulate content in the

harvested organs. Simple, non-invasive in vivo imaging of biodistribution in preclinical

animal models is a rapidly emerging field utilizing techniques including optical

fluorescence imaging, computed tomography, magnetic resonance imaging and positron

emission tomography.

In Vivo Fluorescence Imaging

New whole-body animal fluorescence imaging techniques, especially those

utilizing the near infrared emission region (NIR), now enable the detection of agents with

intrinsic fluorescence properties or those tagged with biocompatible fluorescence dyes in

live animals. A number of optical imaging approaches that rely on fluorescence,

absorption, reflectance, or bioluminescence as the source of contrast have recently been

described162. In general, selected wavelength photons irradiate a whole animal, passing

through the tissues to excite the fluorescent contrast agent possibly deposited in the

tissue. The fluorescent agents absorb the photons and then emits a characteristic

fluorescence light, which travels back to the surface of the animal. The application of an

emission filter allows for the selected detection of the desired wavelength of emission. A

detector (typically a Charge-Coupled-Device (CCD) camera) captures the signal.

Detectors process the photon signals into a digital image. The resolution and sensitivity

83

of fluorescence bioimaging is limited due to the interference from the endogenous

tissues. The thick, opaque animal tissues absorb and scatter emitted photons and generate

strong autofluorescence, as a result, the intensity of the emitted signal is attenuated and

becomes diffuse or blurred163. Several components within the tissues including small

molecules (sugars, fatty acids, amino acids, nucleotides, ions, water), macromolecules

(proteins, phospholipids, RNA, DNA, polysaccharides), organelles, and cell membranes

collectively absorb and emit light in the ultraviolet through the visible wavelength region

of electromagnetic spectrum162. Absorption by tissue components in this wavelength

range limits effective light penetration. However, these interferences can be minimized

using light in the far-red or near-infrared wavelength ranges as the absorption of light in

this region is limited to deoxyhemoglobin, oxyhemoglobin, water and lipids162. As a

result, use of NIR agents in bioimaging has been used by several investigators and

commercial imaging technology companies, and NIR imaging agents including organic

NIR dyes and nanocrystals with intrinsic NIR emission properties have been developed

for animal imaging164.

Quantum Dots in Small Animal Fluorescence Imaging

In recent years, semiconductor quantum dots (QDs) have proven to be the best

NIR imaging agents for in vivo animal studies. Quantum dots with inherent fluorescent

emissions in the NIR region are readily available enabling their direct imaging without

the need for additional conjugation. NIR-QDs offer advantages over fluorescence

techniques for deep-tissue imaging because both scattering and autofluoresence from

endogenous tissues are reduced in the NIR region124. QDs have size-tunable optical and

electronic properties. The particle size determines the wavelength of fluorescence

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emission. By altering the QD size and its chemical composition, fluorescence emission

may be tuned from the near ultraviolet throughout the visible and into the near-infrared

spectrum, spanning a broad wavelength range from 400-2000 nm137.

Micro-Computed Tomography (Micro-CT) Small Animal

Imaging

Unlike fluorescence/NIR imaging, CT offers imaging using three-dimensional

modalities that help to visualize the distribution pattern of high contrast, electron dense

material in the deeper tissue regions. A typical laboratory micro-CT scanner will consist

of a tungsten-anode X-ray tube coupled to a high resolution X-ray detector system. To

produce a truly 3D dataset using CT, X-ray projection views are acquired at hundreds of

equally spaced angular positions around the object of interest. These views are then used

to reconstruct a CT image, typically using proprietary image processing software

programs165. Several studies have reported the benefits of gold nanoparticles as contrast

agents in computed tomography studies in preclinical animal models166-168. The

commercial availability of gold in extremely small particulate sizes allowed for their use

in the investigation of nanoparticle biodistribution patterns from the nasal cavity using

micro-CT techniques.

Although fluorescence and CT imaging offer several advantages, interference

from endogenous fluorescence or attenuation of CT signals from soft tissues and bone

provides limitations to their overall use in live animal imaging. The weak signal

provided from single nanoparticles or small aggregates of nanoparticles remains difficult

to distinguish from these background signals, and, as a result, the observations made

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using these techniques are primarily limited to stronger signals emitted from

accumulated, aggregates of nanoparticulate materials.

Materials and Instrumentation

Near-infrared dye, IRDye (IRDye® 800CW carboxylate, Ex/Em-774 nm/789 nm)

was purchased from LI-COR, Lincoln NE). Near-infrared quantum dots (NIR-QDs),

COOH-QDs (8 μM Qdot® 800 ITK carboxyl quantum dots in 50 mM borate buffer) and

PEG-QDs (2 μM Qtracker® 800 vascular labels in 50 mM borate buffer with PEG surface

coating) were purchased from Molecular Probes (Life Technologies Inc., Eugene, OR).

The hydrodynamic diameter of these NIR-QDs was ~ 20 nm and they have a broad

excitation range (405-760 nm) and narrow emission range with emission maxima of ~

800 nm. Gold nanoparticles (AuNPs) (Mvivo AU, colloidal suspension of gold

nanoparticles (200 mg/mL) in 10 mM phosphate buffered saline, core size 14±2 nm)

were purchased from MediLumine Inc. (Montreal, Canada). 2, 4-Dinitrophenol,

chlorpromazine, methyl-β-cyclodextrin and amiloride were purchased from Sigma

Chemical Co. (St.Louis, MO).

Animals

Male BALB/C mice were purchased from Harlan Sprague (Indianapolis, IN).

Animals were maintained on a twelve-hour light/dark cycle and allowed access to food

and water ad libitum. Mice were 5-7 weeks of age and weighed 20 to 25 g at the time of

the experiments. Animals were acclimated to the animal facility for at least 48 h prior to

use. All the experimental protocols were approved by The Institutional Animal Care and

Use Committee at the University of Iowa (protocol number: 1403044). Prior to dosing,

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animals were anesthetized by intraperitoneal injection of pentobarbital (50 mg/mL, 20 μL

per animal).

Administration of Quantum Dots

For intranasal administration, anesthetized mice were placed on a lab bench in the

supine position with head elevated slightly and a 5 μL volume of QD dispersion

(30 μg/animal of COOH-QDs and PEG-QDs) was slowly dropped into one nostril of

each mouse using a micropipette. For intravenous administration, 50 μL of QD dispersion

(60 μg/animal of COOH-QDs) were administered as a bolus dose via a retro-orbital

injection in each mouse. For studies including inhibitors, a cocktail of methyl-β-

cyclodextrin (10 mg/mL), amiloride (0.1 mg/mL) and chlorpromazine (0.1 mg/mL) was

prepared in normal saline. The stock solution (12 μg/ μL) of COOH-QDs was diluted to 6

μg/μL with this cocktail of inhibitors solution and a 5 μL was administered into the

nostril of each mouse (30 μg/animal dose of COOH-QDs).

Initial animal imaging studies were performed using intranasal instillation of

IRDye® solutions. A 5 μg dose of IRDye® (5 μL volume of 1 μg/μL IRDye® solution in

normal saline) was instilled into one nostril of an anesthetized mouse in supine position.

For studying QD distribution, animals were divided into five groups (n=3 each group);

the first group of animals were administered COOH-QDs intranasally, the second group

were administered PEG-QDs intranasally and the third group of animals were

administered COOH-QDs intravenously, the fourth group received COOH-QDs in

endocytic cocktail inhibitor solution and the fifth group served as a control group, those

animals received an intranasal instillation of normal saline (5 μL ).

87

In Vivo Fluorescence Imaging

A Carestream In Vivo Multispectral Imaging system (In Vivo MS Fx Pro,

Carestream Health, Rochester NY) was used for obtaining the whole-animal fluorescence

images along with X-ray images. The multimodal imaging capability of the MS Fx Pro

imaging system enabled the acquisition of X-ray and fluorescence images from the

animals simultaneously. Super imposition of X-ray images with fluorescence images

enable the visualization of the anatomical locations of the fluorescence signals in the

mice. The Carestream In Vivo MS Fx Pro system operates on an inverted illumination

and detection platform. The animal resides in a temperature controlled translucent sample

tray/scanning box with an optional anesthesia supply feature. For fluorescence imaging, a

xenon excitation source is used to irradiate the animal from the bottom of the sample tray

and the emitted light is collected at a right inverted angle (90°) and detected by the CCD

detector. For X-ray imaging, the X-rays are irradiated overhead of the scanning box and

the mouse; the radiographic phosphor screen placed beneath the scanning box capture the

transmitted X-ray energy and sends a digitalized signal to the CCD camera placed at the

bottom of scanning box.

After instilling the test QD dispersion, anesthetized animals were placed in either

a prone (animal lies flat with head in upright position) or side position (animal lies on

side position with one side of the body resting on the tray) within the scanning box inside

the in vivo imaging system. The system was configured and a protocol was developed to

acquire an X-ray image as the first step with subsequent acquisition of a fluorescence

image. For X-ray imaging animals were irradiated with high energy X-rays (f-stop=2.86,

filter=0.4 mm, field of view=190 mm, acquisition time = 180 sec) and the resulting signal

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was captured by the CCD camera and processed to an image file using Multimodal®

imaging software. For fluorescence imaging, the animals were irradiated with filtered

light (770 nm filter for IRDye®; 450 nm filter for NIR-QDs) and the fluorescence

emission was captured using an emission filter (830 nm for IRDye®; 790 nm filter for

NIR-QDs) with an acquisition period of 60 sec (f-stop=2.51, field of view=190 mm,

binning=4, acquisition time = 60 sec). Mice were imaged before (baseline images) and

immediately after (time 0) administration of the test sample and at 2, 4, and 24 h for

IRDye® and 1, 2, 5,7 and 24 h for NIR-QDs. The fluorescence signal intensity from the

deeper tissue regions is attenuated and the intensity of emission will be low and cannot be

captured by this system, so at each time point, images were acquired with mice in supine,

prone and side positions in order to improve signal sampling from various positions. For

later sampling times (>15 min), the mice were briefly anesthetized during imaging by

supplying isoflurane vapors (1-2 % in oxygen) through the nose cones inside the

scanning box.

After the 24 h measurment, animals were euthanized by injecting a lethal dose of

pentobarbital (50 mg/mL, 50 μL per animal) intraperitoneally. Various organs from the

euthanized mice, including the brain, olfactory bulb, liver, kidneys, heart, lungs and

spleen were collected, washed with 1X phosphate buffer saline (Sigma Aldrich, St.Louis,

MO), and imaged for QD fluorescence within these organs. To capture any QD

fluorescence from the remaining body tissues, each body was imaged in the supine

position. Also, to measure any remaining QD signal in the nasal cavity, an incision along

the nasal septum of each mouse was made such that the nasal cavity was exposed and the

animals were imaged in the prone position.

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Image Analysis

Images were analyzed using ImageJ software (Openware from NIH). The raw

files (.bip image files) from the MS Fx Pro system were converted to 16-bit .tif files using

the Carestream Molecular Imaging software (Carestream Health, Rochester NY)

provided with the imaging system. These 16-bit .tif image files were opened using

ImageJ software and the fluorescence signal was pseudocolored with “red”. These

pseudocolored fluorescence images were co-registered with the corresponding X-ray

images to better visualize the anatomical location of the fluorescence signal from various

anatomical regions. For whole animal images and images following organ removal, a

region of interest (ROI) analysis was performed by manually drawing a fixed area ROI

(area= 55 mm2) to outline the fluorescence signal in the nasal region. The mean

fluorescence intensity in each ROI was quantified. The background mean fluorescence

intensity was measured by drawing three ROIs with the same area in a background area

where no fluorescence was visually observed, and that mean intensity was subtracted

from the mean fluorescence intensity from the QD signals. Similarly, ROI analyses were

performed by drawing ROI to outline each tissue and the mean fluorescence was

measured after subtracting the background intensity. The resulting ROI values were

plotted using Graphpad Prism software to semi-quantitatively describe the biodistribution

of the nanoparticles.

Distribution of Gold Nanoparticles: Micro-CT Imaging

The biodistribution of gold nanoparticles (AuNPs) following intranasal

administration was studied using Micro-Computed Tomography (Micro-CT). In one set

of experiments, two mice were administered 1 mg gold nanoparticles as a single dose via

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the intranasal route (5 μL of 200 mg/mL) and one mouse received a 20 mg dose of gold

nanoparticles (100 μL of 200 mg/mL) as an intravenous injection via the retro-orbital

route. In another set of experiments, multiple doses of 1 mg AuNPs were administered to

mice (n=2) by administering a 1 mg (5 μL of 200 mg/mL) dose every hour for 4 h via the

intranasal route. Before instilling the dose, animals were anesthetized using an

intraperitoneal injection of pentobarbital (50 mg/mL, 20 μL per animal) for the first dose

and isoflurane inhalation (1-2 % in oxygen) for the 2nd, 3rd and 4th dose. The mice were

scanned with a Micro-CT scanner (Siemens Inveon High-Resolution CT scanner,

Munich, Germany). Two dimensional x-ray images of the mice were acquired by

rotating the source around the animal (50 kV, 500 mA, 99 microns, 360 rotation in 120

steps). The 2D images obtained were used to generate a 3D virtual model for the head

region and abdominal regions. Images were displayed in three orthogonal planes of the

mouse body placed in a prone position as shown in Figure 4.1 below. The transverse

plane (axial view) divides the mouse body into anterior and posterior regions; the sagittal

plane (lateral view) bisects the mouse body as a right and left side; the frontal plane

(dorsal view) divides the body into ventral and dorsal sections. These 3D models were

analyzed using ImageJ software. For animals receiving a single intranasal dose, CT scans

were acquired 24 h after dosing. For animals receiving multiple doses, CT scans were

acquired at 6 h and 24 h after administration of the first dose. For the control mouse

(intravenous dosing), a single CT scan was taken 24 h after administration. Animals

remained anesthetized during the CT scanning period (~30-60 min) using isoflurane (2.5

% for induction and 1-2% for maintenance).

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Figure 4.1 Anatomical planes of mouse placed in prone position, showing transverse, sagittal and frontal planes169. Reproduced with permission.

Results and Discussion

IRDye® Distribution Following Intranasal Administration

The purpose of this pilot study was to investigate the feasibility of using an

optical imaging system for studying the distribution of the NIR-QDs. This study assisted

in the design of the experiments utilizing quantum dots with respect to the selection of

sampling time points, positions of imaging and ease of detection of the signal. IRDye®

800CW fluorescent dyes are widely used for optical imaging of tumors in animal

models170-171. These dyes are available with several functionalities, and the carboxylate

form of this dye has been described to be non-reactive within the body, hence it is widely

used in animal imaging studies as a control for understanding retention of the dye in

specific organs or locations in the body along with clearance of the dye itself172. The

structure, molecular weight and chemical formula of the IRDye is shown in Figure 4.2.

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Figure 4.2 Structure of IRDye 800CW, chemical formulation: C46H50N2Na4O15S4, molecular weight of 1091.1 g/mol173.

Figure 4.3 is the montage of overlapped images of X-ray signal and IRDye

fluorescence signal in mice using prone and side views at 2, 4 and 24 h after intranasal

administration of the dye. High levels of IRDye were observed in the urinary tract and

digestive system of the mice within 2 h following nasal administration; an increase in the

fluorescence intensity in these regions was observed during the 24 h observation period.

The fluorescence intensity in the nasal region of the mice decreased during the 24 h

post-administration period, but from the images it can be observed that most of the

IRDye remained in the nose during this 24 h period. The decrease in the fluorescence

intensity in the nasal region suggests the absorption of the dye from the nasal mucosa into

the systemic circulation, and the strong fluorescence in the urinary system (urethra and

bladder) suggests the dye is primarily eliminated via the kidney.

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Figure 4.3 Co-registered fluorescence and x-ray whole animal images of a representative mouse showing biodistribution of IRDye after intranasal administration. Green color represents the pseudo-colored NIR emission signal from the IRDye. Presence of IRDye in the nasal region can be observed in both dorsal and side views up to 4 h and only in the dorsal view at 24 h. In the 2 h side view image, a strong presence of dye in the throat region can be observed. The majority of the dye seemed to reside in the abdominal region and was likely associated with the digestive and urinary systems.

Prone view

Side View

2 h 4 h 24 h

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A study by Marshall et al. also reported the accumulation of IRDye in the kidneys

and urinary system of rats following intravenous administration (5 mg/kg dose), along

with additional accumulation in the lungs, liver, spleen, and reproductive organs 174.

However in the present study, when the IRDye was administered intranasally

(0.25 mg/kg) in mice, no such accumulation in the lungs, liver and spleen was observed,

instead a diffuse signal from the abdominal cavity and urethra was observed which might

be the result of intranasal absorption into the systemic circulation followed by elimination

from blood through the urinary system. Also, the fluorescence signal from the throat

region 2 h after intranasal administration suggest that a portion of the IRDye was

swallowed or cleared from nasal cavity through mucociliary clearance into nasopharynx,

and subsequently into the GI tract. The IRDye from GI tract may also be absorbed into

the systemic circulation and eliminated through urinary system. Some portion of the

administered dye also appeared to be lost from the nose when the mice rubbed their paws

on their noses (fluorescence from the paws can be seen in Figure 4.3).

Distribution of COOH-QDs Following Intranasal

Administration

Figure 4.4 shows typical NIR fluorescence images co-registered with x-ray

images of whole mice (dorsal view) after intranasal and intravenous administration of

COOH-QDs (carboxylate-modified quantum dots with 800 nm emission). Fluorescence

intensities reported in arbitrary units (a.u.) in the nasal region as a function of time are

depicted in Figure 4.5. Following intranasal dosing, a strong fluorescence signal was

observed only in the nasal region of the mice. The intensity became stronger from 5 min

to 5 h post injection and eventually dissipated from 7 h to 24 h.

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Figure 4.4 Composite fluorescence images co-registered with x-ray images of mice showing the distribution of COOH-QDs after intranasal (top row) and intravenous (middle row) administration. The control group received normal saline is shown in the bottom row. The red color represents the fluorescence signal from COOH-QDs. A gradual decrease of fluorescence intensity in the nasal region (yellow circle in top row image) from 5 min to 24 h can be observed after intranasal dosing, whereas intravenous dosing resulted in high fluorescence intensities in the abdominal region within 2 h. Gradual decreasing intensities can be seen up to 24 h. Sequential images for an individual mouse are provided in Appendix-F.

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0 2 4 6 8

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N o rm a l S a lin e i.n .*

*

Figure 4.5 Mean fluorescence intensity from nasal regions (55 mm2 ROI; circled region shown in insert) of animals administered COOH-QDs via intranasal (i.n.), or intravenous (i.v.) routes. * represents statistical significance between i.n. and i.v. fluorescence intensities when tested with Student’s t-test at p<0.05 (n=3).

The dissipation of fluorescence intensity in the nasal cavity within 24 h might be

the result of redistribution of the nanoparticles into other regions of the body from the

deeper tissue regions of the nasal mucosa, but no measurable signal was observed in any

other anatomical region of the animal at the 24 h imaging session. Following intravenous

dosing, a weak fluorescence was observed initially (5 min) in the nasal region which

dissipated within 1 h of dosing (Figure 4.5). This might be due to close proximity of

retro-orbital sinus to the nasal regions, where a portion of the signal from the

nanoparticles administered to the eye could “spill over” and be measured in the identified

nasal ROI. In one of the animals (Mouse 1), a strong fluorescence was observed in eye

region following the retro-orbital injection (Figure F.3 in Appendix-F) for the entire

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study period (24 h), this might be due to accumulation of the nanoparticles in the retro-

orbital sinus. Transfer of particles to the nasal region through the dorsal nasal vein from

the retro-orbital sinus may also occur and contribute to the initial fluorescence signal in

the nasal region following retro-orbital injection175. Strong fluorescence in the abdominal

region, likely liver and spleen regions of the mice following administration via the

intravenous route shows the typical systemic distribution pattern of the nanoparticles. The

cells of the reticuloendothelial system in the liver and the spleen effectively remove the

great majority of particles from blood176. Immune cells including macrophages,

specialized endothelial cells lining the sinusoids of the liver, spleen and bone marrow

may internalize administered QDs and present them to the reticuloendothelial system

(RES), specifically the liver and spleen. Intranasal administration of normal saline in a

control group of animals did not show any signs of fluorescence in either the nasal region

or in any other anatomical region, but enables the quantification of the baseline

autofluorescence from these tissues.

To investigate any accumulation of particles in the deeper nasal tissues, animals

were sacrificed 24 h after dosing and the nasal mucosa was exposed by making an

incision through the nasal septum and the nasal cavity was opened to prior to imaging

along the incision. A strong fluorescence signal in the nasal tissues was observed, which

can be seen in Figure 4.6 and Figure 4.7. The signal was quite diffuse, the exact location

of the nanoparticles was unclear. The fluorescence signal observed in the deeper nasal

tissues of animals following intravenous QD administration or in the control group was

very weak compared to the signal from animals administered QDs via intranasal route

(Figure 4.7).

98

These results demonstrate the accumulation of the nanoparticles in the deeper

nasal tissues over long (up to 24 h) post-exposure. To further investigate the distribution

of particles in other anatomical regions, various organs of interest (liver, spleen, lungs,

kidneys, heart, brain and olfactory bulb) were harvested from euthanized animals 24 h

after intranasal and intravenous dosing and the organs were imaged using the same

instrument settings as described for the whole animal imaging.

Figure 4.6 Fluorescence images co-registered with x-ray images of mice 24 h after COOH-QD administration. Upper panel shows images of anesthetized, live mice with intact nasal cavity and lower panel shows images of euthanized mice with exposed nasal cavity. Opening of the nasal cavity enabled visualization of the strong fluorescence signal from COOH-QD accumulation in the deeper nasal tissues following intranasal administration that was not visible in the mice with intact nasal cavities. Intravenous administration did not show nasal tissue accumulation, even in the exposed nasal cavity images. Images of all mice after opening nasal cavity are provided in Appendix-F.

99

Figure 4.7 Mean fluorescence intensities of COOH-QDs from the nasal regions of mice with intact nasal cavities and exposed nasal cavities after 5 min and 24 h following intranasal (30 ug/animal) and intravenous (60 ug/animal) administration. * represents statistical significance between i.n. and i.v. fluorescence intensities when tested using Student’s t-test at p < 0.05 (n=3).

Figure 4.8 shows the QD fluorescence images co-registered with the x-ray signal

from the harvested organs; images from the control animals are also included. Figure 4.9

shows the numerical comparison of the fluorescence intensities (a.u) from the harvested

organs based on treatment group. From these images it can be observed that the

distribution of QDs beyond the nasal cavity to more distant regions is negligible as the

fluorescence intensities from all of the organs, including brain and olfactory bulb, were

similar to the intensities measured from the control group. In contrast, quantum dots

administered via the intravenous route accumulated in the liver and spleen of the animals,

even up to 24 h after dosing. The transfer of particles to the liver and spleen following

intravenous dosing is expected, based on known distribution patterns for nano-and

microparticles to the reticuoloendothelial organs147.

5 m

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*

100

Figure 4.8 Fluorescence images co-registered with X-ray images from various harvested organs of mice 24 h following intranasal administration (A) and intravenous administration (B).

101

Figure 4.9 Mean fluorescence intensities from various organs of mice 24 h after intranasal (30 ug/animal) and intravenous administration (60 ug/animal) of QDs.* represents statistical significance between i.n. and i.v. fluorescence intensities when tested using Student’s t-test at p < 0.05 (n=3).

The imaging results suggest that the majority of the quantum dots remained in the

nasal tissues even after 24 h of intranasal administration. This is in good agreement with

observations of the in vitro transport of quantum dots through the excised bovine nasal

epithelium (Chapter 3), where during a 2h of study period, nanoparticles were observed

to accumulate in the epithelial and submucosal regions of respiratory and olfactory

epithelium, and in no cases were the particles observed to cross the full-thickness tissues

and transfer into the receiver chamber.

Intranasal uptake of nanoparticles (diameter < 100 nm) have been reported by

several investigators70, 72, 82, however there are very few studies showing evidence of

distribution of intact particles from the nasal mucosa to distant tissues. Brooking et al.

previously reported that small, 20 nm 125I-radiolabelled latex nanoparticles showed a

cumulative uptake of ~ 3.25% of the administered dose from the nasal mucosa of rats into

Liver

Lungs

Bra

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the systemic circulation while larger, 100 nm particles showed ~2% transfer37. However,

the fate of the remaining dose was not discussed in the report; It is likely that the majority

of these nanoparticles remained in the nasal cavity for longer periods of time. Brooking et

al. used liquid scintillation gamma counters to quantify the concentration of radiolabeled

particles in blood and tissues. Gamma counters have a higher sensitivity and are able to

detect trace amounts of the radioactivity, whereas fluorescence imaging has limited

sensitivity and the quantification of the present COOH-QDs uptake using whole animal

images will be difficult. Brooking et al. also reported higher levels of radioactivity in the

liver (~0.05 % of administered dose) and kidneys (~0.1% of administered dose) 3 h after

intranasal instillation of 20 nm nanoparticles in rat, which is not in agreement with the

findings from this QD intranasal uptake study. QDs, which are ~20 nm might be expected

to reach the systemic circulation and further accumulate in more distant organs like the

liver and spleen due to their extremely small sizes and likely enhanced mobility. No such

distribution was observed even when the harvested organs were imaged (Figure 4.8) to

improve the accuracy. This may be the result of too low of an amount of QDs transferred

to be detected with the fluorescence imaging, whereas the radioactive labeling of the

particles enabled Brooking et al. to quantify amounts as low as 0.1 μg particle mass in

tissues.

Numerous studies have been reported about the nose-to-brain transport of

materials via the olfactory and trigeminal neuronal pathway pathways70, 72, 82. In one

study, Oberdorster et al. reported significant transfer of 35 nm 13C radiolabeled graphite

particles to the lungs, cerebrum, and cerebellum within 24 h after nasal inhalation in

rats70. However, these authors did not clearly differentiate whether the uptake into the

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brain was primarily via neuronal pathways or, instead, via the systemic circulation. De

Lorenzo reported the transfer of silver-coated gold nanoparticles (~50 nm) from the

olfactory mucosa to the olfactory bulb of squirrel monkeys within 30-60 min of intranasal

administration using the olfactory neuronal pathway74. These investigators were able to

visualize individual gold particles in the cytoplasm of axons of olfactory neurons using

transmission electron microscopy73. In another study, Mistry et al., used confocal

microscopy and showed that larger, either 100 or 200 nm, fluorescent polystyrene

microparticles did not have access to the olfactory neuronal pathways in mice even after

repeated intranasal administration up to 4 days. Instead, the majority of the polystyrene

microparticles were accumulated in the olfactory and respiratory epithelium177.

The average diameter of the olfactory axons in 2-month-old rabbits (age

equivalent to 8 yr old human) was shown to be around 200 nm, and since the diameter of

the olfactory axons reduces as they pass through the basement membrane, the passage of

particles > 100 – 200 nm within the axon is difficult73. The QDs used in these studies are

smaller (~20 nm) compared to 50 nm gold nanoparticles used by De Lorenzo and 35 nm

carbon particles used by Oberdorster et al., which might enable relatively easy access to

olfactory neuronal pathways and the olfactory bulb. Unlike the previously discussed

reports, however, when the quantum dots were administered intranasally, no signs of QD

transfer to the brain or olfactory bulb were observed either using whole animal images or

images of the extracted brain. This finding again may be the result of the limited

resolution and sensitivity of fluorescence whole animal imaging. Utilization of electron

microscopy enabled DeLorenzo to visualize either individual particle or small aggregates

in the cells; such visualization with whole animal fluorescence imaging would be

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difficult. However, all of the previously published studies used extensive microscopy

techniques including confocal and electron microscopy to qualitatively determine the

biodistribution of particles. The fluorescence whole animal imaging used in the present

study enabled an examination of the distribution of the intranasally administered

nanoparticles, both qualitatively and semi-quantitatively using a simple non-invasive

method and was able to show that majority of nanoparticles remained in the nasal cavity

after 24 h following intranasal administration.

Effect of Particle Surface Modifications on Intranasal

Uptake

The surface properties of nanoparticles also play a major role in determining their

uptake across the nasal mucosa177. In this study, the distribution of ~20 nm PEGylated

quantum dots (PEG-QD) was compared with the distribution of ~20 nm carboxylate

surface modified quantum dots (COOH-QD) following intranasal administration in mice.

The fluorescence images co-registered with x-ray images of mice which were

administered PEGylated QDs (PEG-QD) are shown in Figure 4.10 and the mean

fluorescence intensities in the nasal region as a function of time are depicted in Figure

4.11. From these images it appears that the majority of the PEG-QDs remained in the

nasal cavity for only 2h, whereas COOH-QD particles remained in the nasal tissues for

longer periods (up to 24 h).

105

Figure 4.10 Composite fluorescence images of mice co-registered with corresponding x-ray images. These images compare the distribution of (upper panel) PEG-QDs (30 μg/animal in 5 μL volume) and (lower panel) COOH-QDs(30 μg/animal in 5 μL volume) at various time points after intranasal administration. Sequential images for all individual mice are provided in Appendix-F.

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P E G -Q D s i.n .

Figure 4.11 Comparison of mean fluorescence intensities from the intact nasal region of mice (from a 55 mm2 area ROI as shown in Figure 4.4) following intranasal administration of COOH-QDs and PEG-QDs at same dose of 30 μg/animal in 5 μL volume. (n=3).

Interestingly, no fluorescence signal was observed when the deeper nasal tissues

of mice receiving PEG-QDs were exposed by opening nasal cavity and directly

visualized after 24 h of dosing (Figure 4.12 and Figure 4.13), whereas a significant

accumulation of COOH-QDs was observed. It is likely that the PEG-QDs on the nasal

tissues may have been emptied into gastro intestinal tract through either mucocilary

clearance or swallowing, and are either in the GI mucosal tissues (difficult to detect using

whole animal fluorescence) or excreted. After 24 h, no fluorescence signal from PEG-

QDs was observed either in whole animal images or in the excised organs or in the nasal

tissues. This observation is in agreement with the previous in vitro studies, where PEG-

QDs showed negligible uptake into excised bovine respiratory and olfactory tissues

(Chapter 3.4.3), and the majority of particles remained in the donor chamber of the

Navicyte® diffusion apparatus.

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Figure 4.12 Fluorescence images of mice co-registered with corresponding x-ray images 24 h after intranasal administration of PEG-QDS and COOH-QDs. The upper panels are images of mice with intact nasal cavities. Lower panels are images of mice following the opening of the nasal cavity along the septum prior to imaging. Images of all mice after opening nasal cavity are provided in Appendix-F.

108

Figure 4.13 Mean fluorescence intensities from the deeper nasal tissues of mice with intact and exposed nasal cavities 24 h after intranasal administration of PEG-QDs and COOH-QDs, at same dose of 30 μg/animal * represents statistical significance between COOH-QDs and PEG-QDs fluorescence intensities tested using Student’s t-test at p<0.05 (n=3).

Several previous studies have investigated the effect of surface modification of

micro/nanoparticles with PEG on the uptake of particles across the nasal mucosa178-179.

The results reported are inconsistent or unclear regarding whether PEGylation improves

the uptake of intact nanoparticles across the epithelial tissues. For example, a study

reported by Vila et al. showed improved antibody levels in the blood of intranasally

administered tetanus toxoid when delivered in PEG-PLA microparticles (~ 196 nm and

1500 nm) compared to the administration of the toxoid in PLA particles of similar size148.

There was no clear evidence in the report if the PEG-PLA particles were taken up to a

5 m

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109

greater extent or whether the dissociated/released tetanus toxoid was taken up as free

protein, instead. In comparison, the observations from the present study using PEG-QDs

(~10 - 20 nm) showed no such uptake/presence of these ultrafine nanoparticles when they

were either incubated with bovine nasal tissue explants or after their intranasal

administration in mice. This disagreement in results may be related to the presence of

unentrapped antigen (tetanus toxoid) on PEG-PLA microparticles (~200 and 1500 nm)

surfaces which potentially may trigger immune cells including M cells in the NALT and

assist in internalization of these microparticles into nasal epithelial cells. Whereas in the

present study, the surfaces of QDs might be completely passivized by PEGylation which

might result in improved stealth properties with resulting lack of uptake by any cells.

There are also additional reports that show negligible or reduced uptake of PEGylated

nanoparticles across other cells. An in vitro investigation by Kumaraswamy et al.

reported that 45 – 70 nm polystyrene nanoparticles with PEGylated surfaces showed

almost no uptake into microglial cells following 1 h incubation with neural cells whereas

carboxylated surface functionalization showed significantly higher uptake into those

cells149. This result is somewhat in agreement with the findings from presenty study using

PEG-QDs administered intranasally in mice, which showed no accumulation in deeper

nasal tissues compared to COOH-QDs.

Passivation of nanoparticles by surface modification with a hydrophilic PEG is a

widely used technology to avoid rapid uptake by scavenger macrophages or other cells of

the immune system or RES. The PEGylated particles are believed to avoid non-targeted,

non-specific binding of proteins which can result in their increased circulation time in

blood147. It has been shown that charged nanoparticles, especially cationic nanoparticles

110

interact with proteins present on the cell surfaces and can trigger endocytic uptake which

increases uptake into the epithelial cells through the vesicular pathways45. PEGylation of

nanoparticles may decrease these interactions with proteins and thus the extent of uptake

of these particles into epithelial cells through endocytic pathways. This suggests that

most of the PEG-QDs in the nasal cavity likely cleared rapidly from the nasal cavity via

mucociliary clearance rather than being transferred into tissues.

Mechanistic Evaluation of COOH-QD Uptake from Nasal

Tissues Using Whole Animal Imaging

In above sections it was shown that the majority of the intranasally administered

COOH-QDs accumulated in the deeper nasal tissues in mice. In chapter 3 using an ex

vivo bovine nasal tissue model, it was shown that the pharmacological inhibitors

chlorpromazine (CPZ), methyl- β- cyclodextrin (MBC) and amiloride inhibited clathrin-

mediated endocytosis, caveolae-mediated endocytosis and macropinocytosis pathways

affecting the uptake of COOH-QDs across bovine nasal tissues, respectively. The

involvement of these endocytic pathways was investigated in live animals using NIR

imaging techniques along with the administration of a cocktail of inhibitors of

endocytosis to halt the these uptake processes. A 5 μL volume of inhibitor solution

containing 0.1 mg/mL CPZ, 0.1 mg/mL amiloride and 10 mg/mL MBC was administered

to one nostril of each mouse. After 15 min, 30 μg of COOH-QDs dispersed in the

inhibitor cocktail was administered into the same nostril. Figure 4.14 shows the

fluorescence images of whole animals at various time points after dosing and Figure 4.15

shows a graphical representation of the mean fluorescence intensities of COOH-QDs in

the nasal region as a function of time.

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Figure 4.14 Whole animal fluorescence images of mice co-registered with corresponding x-ray images comparing the distribution of COOH-QDs (top row) in the presence of a cocktail of endocytic inhibitors (bottom row) at various time points after intranasal administration. The red fluorescence in the nose region represents signal from QDs. Inclusion of inhibitors resulted in no difference in QD fluorescence signal when compared to QDs without inhibitors. Sequential images for all individual mice are provided in Appendix-F.

112

0 2 4 6 8

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N o rm a l S a lin e i.n .

In h ib ito r c o c k ta il + C O O H -Q D s i.n .

Figure 4.15 Mean fluorescence intensities from nasal regions (from a 55 mm2 area ROI as shown in Figure 4.4) of live animals in presence and absence of inhibitor cocktail. No difference in fluorescence signal from QDs in presence of inhibitor cocktail is observed. (n=3).

The nanoparticles appeared to remain in the nasal cavity for 2 h to 5 h and then

from 7 h to 24 h the fluorescence signal from the particles slowly dissipated. The

endocytic inhibitors did not appear to affect the transport behavior of COOH-QDs in the

nasal region. The fluorescence images of mice that received inhibitor cocktail were

observed to be similar to the images of mice which received COOH-QDs without

inhibitors.

From Figure 4.16 and Figure 4.17, it can be observed that 24 h of post

administration, particles accumulated in the deeper nasal tissues, even in presence of

inhibitors. These results are somewhat different from the results from in vitro studies,

where uptake of particles into the nasal tissues was shown to be negligible in presence of

these inhibitors. A possible explanation for these inconsistent results may be an

113

insufficient sustained concentration of inhibitors used in whole animal studies where

endocytic inhibition was not sustained over the 24 h imaging interval. The inhibitors

administered to the mice may have been eliminated via mucociliary clearance or, more

likely, were absorbed into the systemic circulation. In the case of in vitro studies, the

concentration of the inhibitor was maintained throughout the entire study period.

Figure 4.16 Images of mice showing the fluorescence from the deeper nasal regions 24 h after intranasal administration of COOH-QDs in the presence and absence of an endocytic inhibitor cocktail. The upper panels are images of mice with intact nasal cavities. Lower panels are images of mice following the opening of the nasal cavity along the septum prior to imaging. Images of all mice after opening nasal cavity are provided in Appendix-F.

114

Figure 4.17 Mean fluorescence intensities from the nasal tissues of mice with intact and exposed nasal cavities 24 h after intranasal administration of COOH-QDs in presence and absence of an endocytic inhibitor cocktail (n=3).

Several previous investigations have used in vitro models, including cell lines and

excised tissue models, to study the cellular internalization or tissue uptake of

pnanoarticulate matter145, 152, 180-181. This study attempted to investigate if a live animal

model could be used to evaluate the uptake of quantum dots following nasal

administration. However, it appears to be difficult to probe endocytosis processes in in

vivo models due to continuous clearance/elimination of the inhibitors from the nasal

cavity either by mucociliary clearance or by absorption into systemic circulation. This

suggests that care should be taken when extrapolating in vitro results to in vivo processes

when evaluating uptake mechanisms in the nasal mucosa.

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115

Translocation of Gold Nanoparticles Measured Using

Micro-CT

Using quantum dots, it was shown that these extremely small nanoparticles

accumulate in the deeper nasal tissues following nasal administration. However, due to

the limitations of poor spatial resolution and the diffusive signal strength of the

fluorescence whole animal imaging technique, it was difficult to determine the exact

anatomical location or the dynamics of tissue distribution of the quantum dots following

intranasal administration. An alternative attempt to visualize the distribution of

intranasally administered ultrafine nanoparticles in live animals was made using

computed tomography (CT) and readily available, 15 nm gold nanoparticles (AuNPs).

Figure 4.18 shows micro-CT images of the mouse head region in three-

dimensional axial, dorsal and lateral views, obtained 24 h after administration of a single

dose of AuNPs. Particles were observed in the deeper middle and superior turbinate

regions of the nasal mucosa. Reconstruction of the head region in 3D mode (Figure 4.19)

revealed the presence of AuNPs as an accumulated mass in the anterior nasal tissues;

small amounts of the nanoparticles were also observed to transverse into the posterior

region of the nasal cavity and into the superior turbinate region of the nasal mucosa.

Nanoparticles in the superior turbinate region were observed to be in closer proximity to

the olfactory bulb, however, there was no clear indication of the presence of particles in

the olfactory bulb or in any regions of the brain. This finding is similar to observations

from the fluorescence imaging of mice administered COOH-QDs. The high contrast

signal from the micro-CT images enabled the visualization of the discrete location of the

nanoparticle mass unlike the nasal cavity and associated tissues.

116

Figure 4.18 In vivo micro-CT imaging of a mouse head region 24 h after intranasal (A) and intravenous (B) administration of 15 nm gold nanoparticles. Axial (left column), dorsal (middle column) and lateral views (right column) are depicted. Yellow colored arrows show the accumulation of gold particles in the nasal conche following intranasal administration, whereas no such accumulation was observed in the same regions after intravenous administration.

117

Figure 4.19 In vivo micro-CT 3D lateral view of the head regions of a mouse showing the accumulation of AuNPs in anterior and posterior regions of the nasal cavity (yellow arrows) 24 h after intranasal administration.

Following intravenous administration of AuNPs, a greater contrast was observed

in the blood vessels, spleen, heart and liver of the animals (Figure 4.20), which are the

primary RES organs involved in the uptake of particulate matter from blood. Immediately

after the retro-orbital injection of AuNPs, the mice exhibited a bluish skin color that

remained over the following 24 h period. This change is due to presence of gold

nanoparticles in the blood, trapped in the endothelium, or leached into interstitial spaces.

No accumulation of particles was observed in the deeper nasal tissues of animals

following intravenous administration, however excised organs including the liver, spleen,

and heart appeared bluish color compared to the control organs, which also support the

conclusion of particle transfer to these organs.

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Figure 4.20 In vivo micro-CT images of mouse in axial (left), dorsal (middle) and lateral (right) views 24 h after intravenous administration of AuNPs. High contrast CT signal in heart (yellow arrow), liver (purple arrow) and spleen (blue arrow) can be observed.

119

In a follow up study in order to increase the total dose of gold nanoparticles, mice

were given four intranasal doses of AuNPs every 1 h and imaged 6 h and 24 h after the

first dose. Six hours post administration of the first dose accumulated masses of gold

particles were visible as high contrast regions in the deeper nasal tissues of mice (Figure

4.21a). A gradient of accumulation of the gold particles from the anterior to posterior

regions of the mouse nasal mucosa showed the gradual transfer of particles from the

nostrils to the deeper nasal tissues. However, a 24 h-post administration micro-CT image

(Figure 4.21b) showed that the majority of the nanoparticles has disappeared from the

nasal tissues while a small quantity remained in the anterior and posterior regions of the

nasal mucosa (Figure 4.21). There was no evidence of the gold nanoparticles transferring

or remaining in the brain or other head regions of the mice. A previous report by De

Lorenzo showed that ~50 nm gold particles can be transported to the olfactory bulb

following intranasal administration and appeared as small aggregates74. It is likely that

the smaller gold particles (~15 nm) should also have access to this pathway and most

likely, some portion of the gold nanoparticles may be either distributed to parts of the

brain or distant tissues as small aggregates or individual particles and are not detectable

using the micro-CT technique. While, no evidence of high contrast discrete signal from

gold nanoparticles was observed either in throat region or the GI, some portion of the

gold nanoparticles might have also been cleared from the nasal cavity through

mucociliary clearance to the GI tract.

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Figure 4.21 In vivo micro-CT images of the head region of a mouse 6 h (A) and 24 (B) after multiple dosing of AuNPs via the intranasal route. Accumulation of AuNPs (colored in red) in the nasal mucosa can be observed.

121

While both fluorescence imaging and micro-CT imaging are both non-invasive

methods used for studying ultrafine nanoparticle uptake in whole animals, the signal from

quantum dots in whole animal fluorescence images is very diffuse and scattered, which

increases the difficulty of identifying the discrete anatomical locations of the

nanoparticles. Additional imaging of harvested organs is able to confirm that QDs

accumulate in a variety of tissues and organs. In comparison, micro-CT enabled

visualization of the nanoparticle accumulation in animal organs without the need for

organ isolation and the signal obtained with micro-CT was more resolute compared to the

diffuse signal from the fluorescence images. The distribution and elimination of ultrafine

nanoparticles from the nasal tissues is better demonstrated using AuNPs with micro-CT

compared to QDs and NIR imaging, yet with regard to the understanding of drug-

containing ultrafine nanoparticles distribution, it is unlikely that gold nanoparticles will

be a suitable matrix for drug delivery, thus further use of these nanoparticles may have

limited application in drug delivery.

Conclusions

From these studies it can be concluded that extremely small nanoparticles are

transferred from the nasal cavity into the posterior turbinate region and in close proximity

to the olfactory bulb. Both NIR imaging and micro-CT imaging were useful tools for

visualization of in vivo nanoparticle distribution. However, neither of these techniques

was able to visualize a discrete signal associated with nanoparticles in the brain or other

parts of the body, due to the inability to capture the signal from small aggregates or

individual nanoparticles.

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The accumulation of nanoparticles in the nasal tissues is dependent on the

physical properties of the particles, where surface modifications providing negatively-

charged particles (COOH-QDs) increased the uptake of nanoparticles from the nasal

cavity compared to neutral, PEGylated surface modifications.

These studies suggest that ultrafine nanoparticles instilled in the nose can transfer

to posterior regions of the nasal cavity and accumulate in the tissues associated with the

turbinate regions. The accumulation of extremely small nanoparticles in the turbinate

regions can result in release of the encapsulated cargo into the local external

environment, which may eventually result in transfer to the brain via olfactory neuronal

pathways, including the perineuronal spaces, or to other distant tissues following

absorption and transfer to the systemic circulation. There is great interest in transnasal

vaccine delivery using nanoparticulate systems and the accumulation of these

nanosystems in and around lymphatic vessels in the deeper nasal mucosa may result in

the release of antigens from the particles, transfer to circulating immune cells, and uptake

into the lymphatics for subsequent antigen processing and antibody production.

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CHAPTER V

UPTAKE OF MICROEMULSIONS FROM NASAL TISSUES

Introduction

In most current drug delivery application, nanomaterials are assumed to be solids

composed of single or multiple materials. There are other nano-sized systems that exist in

liquid form, and some of these systems have also been evaluated for their potential as

drug delivery systems. Microemulsions, for example are two phase liquid dispersions

where the droplet sizes of the emulsified materials are typically 10 – 100 nm.

Microemulsion dosage forms have been developed for nasal administration, and

numerous reports exist regarding their success in improving nasal bioavailability182-187.

Unlike solid nanoparticles, drugs in microemulsions do not require a nanoparticle support

for conjugation, instead these systems form drug-loaded microemulsions spontaneously

while simultaneously increasing the solubulization capacity of drugs in the

microemulsions188-189. Microemulsions are thermodynamically stable, isotropic,

translucent systems consisting of a lipid phase, and an aqueous phase with surfactants and

stabilizers at the interface between phases. The dispersed phase in a microemulsion is

typically 10-100 nm in size190. The high lipid content in microemulsions enable improved

bioavailability of lipophilic drugs with low water solubility and, also unlike macro-

emulsions, preparation of microemulsions use relatively simple methods without the need

of high sheer mixing190. These advantages encouraged investigators to explore

microemulsions as alternative delivery systems for various routes of administration

including the oral188, 191-192, transdermal193-194, ocular195-196 and intranasal197-201. Several

studies have shown improved delivery of drugs across the nasal mucosa to the brain when

124

delivered in microemulsions184-187. However, very little information is known about the

mechanisms of drug transfer from microemulsions across the nasal mucosa or the

interaction of the microemulsions with the mucosal tissues. Some researchers have

suggested that the oil droplets in microemulsions act as drug reservoirs and improve drug

absorption by maintaining a constant concentration gradient182-183. There are also reports

that describe components of the microemulsions as permeation enhancers which can

improve the absorption of drugs by altering the mucosal membrane permeability202. It is

well known that particulate matter is taken into cells by endocytic mechanisms, and the

previous chapters in this dissertation describe the endocytic uptake of small, solid

nanoparticles like quantum dots. Similar to solid nanoparticles, the dispersed phase in

microemulsions could act as a fluid-phase nanoparticle with greater mechanical

flexibility, which may change the droplets/particles access to various endocytic pathways.

However, the involvement of any energy-dependent endocytic pathway in the uptake of a

drug-rich dispersed phase of a microemulsion has not been previously investigated. The

purpose of this work is to investigate pathways involved in the transfer of drug from an

oil-in-water microemulsion across the bovine nasal mucosa to evaluate the importance of

the material phase (solid vs liquid) on the uptake of nanomaterials.

Figure 5.1 Chemical structure of diazepam203.

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Diazepam, (7-chloro-1-methyl-5-phenyl-3H-1,4-benzodiazepin-2-one) is a

benzodiapine derivative with anti-anxiety, sedative, hypnotic and anticonvulsant

properties. It is a lipophilic compound with low water solubility (0.05 mg/mL), a pKa of

3.4 and a logP of 2.8203. The structure of diazepam is shown in Figure 5.1. Currently,

intravenous or rectal administration of diazepam can be used in the initial treatment of

status epilepticus204. Even though these methods are effective, more convenient

administration would improve patient comfort and potentially enable more rapid

treatment. Diazepam intranasal delivery has gained the attention of several investigators

recently. A variety of formulation approaches are reported in literature for intranasal

delivery of diazepam, including biodegradable PLGA nanoparticles65 and

microemulsions199. Acorda Pharmaceuticals developed a proprietary diazepam-

containing microemulsion formulation using diethylene glycol monoethyl ether,

propylene glycol monocaprylate, methyl laurate, N-methyl-2-pyrrolidone, ethanol and

water as inactive ingredients205. In a pilot clinical study in healthy human volunteers, the

intranasal administration of microemulsion formulation resulted in similar bioavailability

results of diazepam to that of commercial rectal formulation. However, the formulation

did not show bioequivalent results when administered in epilepsy patients and further

development has been discontinued206. Also, intranasal administration of the

microemulsion resulted in moderate to severe adverse events like nasal discomfort, and

lacrimation205. Another investigational intranasal diazepam formulation (NRL-1)

developed by Neurelis, Inc. using proprietary technology showed ~ 97% absolute

bioavailability (compared to intravenous diazepam administration) in healthy human

volunteers in a pilot clinical study with good tolerability and reasonable variability207.

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Though the composition of the new NRL-1 formulation is not disclosed, the company has

previously reported another diazepam intranasal formulation consisting of permeation

enhancers like glycofurol, which showed poor tolerability and moderate to severe nasal

discomfort in healthy human volunteers208.

The nasal delivery of diazepam is challenging because of its poor water solubility,

which limits the amount of diazepam able to be administered into the nasal cavity and the

resulting amount absorbed across nasal mucosa due to the low concentration gradient.

Although, use of permeation enhancers and microemulsions show good bioavailability of

diazepam after intranasal administration, the components may cause temporary or

permanent discomfit to nasal mucosa, which further reduces the tolerability and safety of

these formulation.

Materials and Methods

Diazepam, isopropyl myristate, sorbitol, polysorbate 80 (Tween 80), and 2, 4-

dinitrophenol (2,4-DNP) were obtained from Sigma Aldrich (St. Louis, MO). High

performance liquid chromatography (HPLC) grade methanol and water were purchased

from Fisher Scientific (Pittsburgh, PA).

Composition and Preparation of Krebs Ringer Bicarbonate buffer (KRB) is

described in previous chapter (Chapter 3).

Solubility Studies

The solubility of diazepam in individual components of the microemulsion was

measured by adding an excess amount of diazepam to individual scintillation vials, each

containing 2 mL of either water, KRB, isopropyl myristate, Tween 80, 70 %w/v sorbitol

solution. The vials were mixed vigorously using a vortex mixer (Analog Vortex Mixer,

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Fisher Scientific, Hampton NH) for 2 min, followed by shaking at 100 rpm using a

VWR®

incubating orbital shaker (Henry Troemner LLC, Thorofare, NJ) for 48 hours at

25 °C. The contents of the vials were centrifuged for 15 min at 3000 g using an

Eppendorf®

AG5810R centrifuge (Hamburg, Germany). The resulting supernatant was

filtered using Millex®

-GS (0.22μm) filter units into HPLC vials and analyzed using the

HPLC method described in above section.

Microemulsion Formulation

The selection of microemulsion system was based on obtaining an oil-in-water

system with dispersed phase diameters <50 nm in size. In a series of papers published by

Ktistis et al.209 and Attwood et al.210, the properties and phase behavior of o/w

microemulsions prepared using polysorbate/sorbitol/IPM/water systems were described.

The key factors influencing the formation of microemulsion were identified to be the

IPM fraction and the polysorbate to sorbitol ratio. In an initial study, Attwood and Ktistis

reported that a 1:2 ratio of polysorbate 60: sorbitol at 40% by weight with 2.7 – 6.7 %

IPM by weight resulted in largest region of stable microemulsion formation with 20-30

nm diameter dispersed phase210. In another report, Ktistis et al., showed that replacement

of polysorbate 60 with polysorbate 80 a higher ratio of sorbitol was needed. A ratio of 1:

2.5 polysorbate 80/sorbitol at 40% by weight with 2.7-6.7% IPM resulted in largest

region of stable microemulsion region in phase diagram209.

Preparation of the microemulsion was modified slightly as published by Attwood

and Ktistis210. The composition of the microemulsion is shown in Table 5-1.

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Table 5-1 Composition of microemulsions with and without drug and 2,4-DNP.

Ingredient Function Composition of

Blank Microemulsion (%)

Composition of DZME (%)

Composition of DZME with 2,4

–DNP (%)

Diazepam Drug 0 0.4 0.4

Isopropyl myristate Lipid 5 5 5

Tween 80 Surfactant 11.6 11.2 11.2

Sorbitol Stabilizer 28.4 28.4 28.4

2,4-Dinitrophenol Inhibitor 0 0 0.018

Water Diluent 55 55 ~55*

* For 2,4-DNP containing microemulsion, an excess amount of 1mM 2,4-DNP in water

solution was prepared and the required amount of this solution was used for preparation

of the microemulsion.

Sorbitol was weighed and ~3/4 of the total water for the microemulsion was

added to sorbitol and stirred using a magnetic stirrer (VWR, Radnor, PA) on a stir plate

(120A, 1000W capable hot plate stirrer, VWR, Radnor, PA) at 400-500 rpm while

heating the mixture to ~55 °C. In a separate beaker, IPM and Tween 80 were heated to

~55 °C and diazepam was added to the mixture and stirred until all the diazepam

dissolved. To this IPM/Tween 80/diazepam mixture, the sorbitol solution was added

dropwise with stirring at 400 - 500 rpm while maintaining the temperature at ~55 °C. The

resulting concentration of diazepam in the microemulsion was 4 mg/mL. Initially, the

system appeared turbid. The remaining amount of water was added dropwise until the

system turned translucent. The diazepam-containing microemulsion was cooled to room

temperature while maintaining the stirring rate at 400-500 rpm. If the system became

turbid during cooling more water was added (always < 1mL) until the system appeared

translucent. A schematic of the preparation method is shown in Figure 5.2. For endocytic

inhibition studies, a 2,4-dinitropehnol (2,4-DNP) containing microemulsion with

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diazepam was prepared using water which contained 1mM 2,4-DNP in solution. A blank

microemulsion was prepared using the same procedure without the addition of diazepam.

The prepared microemulsion systems were stored for not more than 24 h at ambient

temperature.

Figure 5.2 Schematic showing the preparation of diazepam-containing microemulsion. Microemulsions were prepared by adding the required amount of a sorbitol solution of known concentration to a mixture containing IPM, Tween 80, and diazepam in a stirred beaker at 55 °C followed by cooling to room temperature.

130

Characterization of Microemulsions

The mean diameter of the dispersed phase in the blank and diazepam-loaded

microemulsions was measured using dynamic light scattering (Nicomp Particle Sizer,

Model 380 ZLS, Santa Barbara, CA). Size measurements were made at room temperature

using a HeNe laser at a wavelength of 632.8 nm, viscosity of 0.933 cPs and refractive

index of 1.333. Approximately 3 mL of the microemulsions were transferred into a

disposable cuvette (Fisher Scientific, Hanover Park, IL) and placed into the sample

holder followed by a 2 min equilibration period. The neutral density filter was adjusted

until the scattered intensity oscillated around a setting point of 300 kHz. Data processing

software (Nicomp® ZW380, Ver. 1.51) was used for data analysis; channel width,

detector sensitivity, and baseline were automatically adjusted. The zeta potential of the

microemulsion system was measured using a Malvern Nano ZS Zetasizer

(Worcestershire, UK). Approximately 2 mL of the microemulsion were placed into a

disposable folded capillary cell (DTS1070, Malvern, Worcestershire, UK). Measurements

were made using freshly prepared microemulsion prior to any exposure to nasal tissues.

Preparation of Bovine Nasal Tissues

Bovine nasal respiratory and olfactory tissues were obtained from a local abattoir

and prepared as explained in Chapter 3. Small pieces of fresh respiratory and olfactory

tissue was cut and immediately placed in 4% formalin solution for histological

evaluation. Remaining tissues were transported to lab in cold KRB maintained on ice.

Uptake Studies of Diazepam-containing Microemulsions

The underlying cartilage from the mucosal tissues was peeled off, and the fresh

tissues were used immediately for transport studies. Respiratory and olfactory mucosal

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tissues were mounted between the donor and receiver chambers of a Navicyte® diffusion

chamber system (Harvard Apparatus, Holliston, MA), such that the mucosal side of the

tissue faced the donor solution and transport took place from the mucosal to the

submucosal surfaces. The membranes were equilibrated for 20-30 min with 1 mL of pre-

warmed (37 ˚C) KRB in both donor and receiver chambers. The tissues were kept aerated

with carbogen (95% O2 + 5% CO2) at a rate of 2-4 bubbles/second, and the temperature

was maintained at 37 ˚C throughout the experiment. Aeration helps with mixing the

contents of the chambers and supplies oxygen to the tissues. Transepithelial electrical

resistance (TEER) was measured at the beginning and at the end of transport studies

using an EVOM volt-ohmmeter (Model: EVOM2; World Precision Instruments Inc.,

Sarasota, FL). TEER values below 100 Ω*cm2 were indicative of a compromise in

mucosal integrity and those tissues were excluded from use.

After equilibration, buffer in the donor chamber was replaced with 1 mL of the

diazepam-loaded microemulsion (4 mg/mL) and 1 mL of fresh KRB buffer in the

receiver chamber. The transport of diazepam across the tissues was measured by

withdrawing aliquots (200 μL) from the receiver chamber at regular intervals (20, 40, 60,

80, 100 and 120 min). The aliquots removed were replaced with fresh buffer (200 μL) to

maintain a constant volume in the receiver chamber. After 120 min, a 200 μL aliquot of

microemulsion from the donor chamber was taken to determine the amount of diazepam

that remained in the donor chamber. All samples were refrigerated until further analysis.

Total recovery of diazepam after the 120 min transport study was calculated by

adding the amount remaining in the donor chamber and the cumulative amount recovered

from the receiver chamber. Any unaccounted diazepam was assumed to be present

132

within the mucosal tissues. A control study using a synthetic artificial membrane was

performed to justify this assumption and to determine if any diazepam loss was due to

experimental technique. A synthetic, polytetrafluoroethylene (PTFE) membrane of

~ 90 μm thickness was selected as the artificial membrane to eliminate transport across

the membrane so any diazepam transfer or loss would not be due to failures in the

apparatus or sampling error. The PTFE membrane was too thin and did not fit snugly

between the diffusion chambers, hence it was wrapped around a silicone rubber (Silastic®

membrane) of 1.27 mm thickness to avoid any loss of material due to leakage. PTFE

wrapped Silastic® membranes were placed between the donor and receiver chambers of

the Navicyte® diffusion chambers and 1 mL of 4 mg/mL diazepam microemulsion was

added to the donor side and 1 mL of fresh KRB to the receiver side. The transport of

diazepam across the artificial membranes was measured using the same methods as

described above. Total diazepam was measured after 120 min by summing the amount of

diazepam recovered from donor chamber and the cumulative amount of diazepam

recovered from the receiver chamber.

In another control study, diazepam was dissolved in isopropyl myristate and the

uptake of diazepam from the isopropyl myristate solution and the microemulsion were

compared. In this experiment, instead of adding 1 mL of diazepam-containing

microemulsion to the donor chamber, 1 mL of diazepam dissolved in isopropyl myristate

(4 mg/mL) was added to the donor chamber, and the transport of diazepam across the

bovine nasal tissues was measured using the same methods as described above.

133

Histological Evaluation of Bovine Nasal Tissues Exposed

to Microemulsions

Microemulsion-exposed nasal respiratory and olfactory tissues were subjected to

histological evaluations to observe any potential adverse effects caused by the

microemulsion on the tissue integrity. After a DZME transport study, the exposed nasal

tissue area was carefully cut from the mounted tissue and fixed in 10 mL of zinc formalin

solution (Sigma Aldrich, St Louis, MO) for 48 hours. The fixed tissues were treated with

10%, 20%, and 30% sucrose (Spectrum Chemicals, New Brunswick, NJ) solutions each

for 24 hours, successively. The tissue samples were placed into molds containing tissue-

freezing media (TFM-CTM) (Triangle Biomedical Science, Durham, NC) and cryo-frozen

in liquid nitrogen using a snap freezing system (Gentle Jane® Instrumedics Inc.,

Hackensack, NJ). Thin sections (10 μm) of frozen tissues were cut using a Microm®

Cryostat II (HM505E) with a CryoJane system (Microm Ineternational, Waldorf,

Germany) at -35°C. These thick sections were placed on Surgipath® adhesive-coated

glass microscope slides (CSFA-1X, Lecia Bioystems Richmond Inc., Richmond, IL) and

stained using hematoxylin and eosin (Sigma Aldrich, St Louis, MO) at room temperature

using an automatic staining machine (DRS-601 Sakura stainer, Sakura Finetec Inc.,

Torrance, CA). Coverslips were placed on the slides and the stained tissue sections were

imaged using bright field microscopy using an Olympus BX-61 motorized light

microscope (Olympus Microscope and Imagining System Inc., Melville, NY).

For control images, freshly harvested respiratory and olfactory tissue samples

were placed in zinc formalin immediately after harvesting and treated under similar

conditions as described above and examined using bright field microscopy.

134

Transport Inhibitor Studies

To investigate the involvement of energy dependent pathways in the uptake of

diazepam from microemulsions, bovine nasal respiratory and olfactory tissues were

exposed to 2,4-dinitrophenol (2,4-DNP), a metabolic inhibitor that interferes with

synthesis of ATP required for energy-dependent processes, including endocytosis. Nasal

tissues were exposed for 30 min with 1 mL of KRB containing 1 mM 2,4-DNP in the

donor and receiver chambers prior to the subsequent DZME transport study. After 30

min, the donor chamber fluid was replaced with 1 mL of DZME (4 mg/mL) also

containing 1mM 2,4-DNP and the receiver chamber fluid was replaced with1 mL of KRB

containing 2,4-DNP (1 mM). The transport of diazepam through the tissue and into the

receiver chamber was monitored as described in above section.

HPLC Analysis

The HPLC method for quantification of diazepam was developed based on a

previously reported method199. The HPLC system used was an Agilent1100 (Agilent

Technologies Co., Santa Clara, CA) consisting of a G1311A quaternary pump, G1313

ALS autosampler and G1315B diode array detector. Separation of diazepam was carried

out at ambient temperature with a Phenomenx Luna C18 (2) (256 x 4.6 mm, 5 μm

particle size) column protected by a Phenomenex Gemini–NX C18 (4 x 2.0 mm ID)

guard cartridge (Phenomenex Inc., Torrance, CA). The mobile phase consisted of

methanol: water (70: 30 %v/v) as an isocratic mobile phase at a flow rate of 1 ml/min

with a run time of 10 min and a post run time of 2 min between 10 μL sample injections.

Diazepam was eluted at ~ 7.6 min and detected at 254 nm. Concentrations of diazepam

were calculated from a calibration curve obtained using fresh diazepam standard solution.

135

Standard diazepam solutions were prepared by dissolving known amounts of diazepam in

methanol. The linearity of the HPLC method was established over a concentration range

of 1 μg/mL to 1000 μg/mL. A sample calibration curve is shown in Figure 5.3. Below 1

μg/mL diazepam was unquantifiable and this was chosen as the limit of quantification.

Each set of transport study samples consisted blank (mobile phase), standard (known

concentrations of diazepam in methanol) and blank microemulsion. The unknown

concentrations were determined using a calibration curve obtained using freshly prepared

standards in the sample set.

Figure 5.3 Sample calibration curve for analyzing diazepam using HPLC method (n=3). Good linearity of the method was observed from 1 μg/mL to 1000 μg/mL with AUC=35.913* Concentration (μg/mL) and r2=0.9999. Insert is presented for better visualization of the linearity in low concentrations (0 to 100 μg/mL).

Statistical Analysis

Each experiment was repeated at least three to six times and the data are presented

as mean ± standard deviation. Statistical significance was tested using either an unpaired

Student’s t-test or one-way ANOVA, where appropriate. Differences were considered

0

7,500

15,000

22,500

30,000

37,500

45,000

0 200 400 600 800 1000 1200

AUC

Concentra on(μg/mL)

0

1,000

2,000

3,000

4,000

5,000

6,000

0 20 40 60 80 100 120

AUC

Concentra on(μg/mL)

136

significant at p < 0.05. GraphPad Prism Inc. (La Jolla, CA) was used to perform the

statistical testing.

Results and Discussion

Solubility Studies

To develop a microemulsion of a highly hydrophobic molecule like diazepam, the

components selected should have ability to solubilize higher concentrations of diazepam

than water. Figure 5.4 shows the solubility of diazepam in IPM, Tween 80, sorbitol

solution (70 %w/v in water) and water. Among the microemulsion components

selected, Tween 80 and IPM showed the highest solubulization capacity for diazepam.

Figure 5.4 Solubility of diazepam in various components used in microemulsions. Column bars are labeled with the mean solubility value (mg/mL). Results are expressed as mean ± standard deviation of three replicates.

Characterization of Microemulsions

Diazepam-containing microemulsions and blank microemulsions were prepared

and characterized for dispersed phase size, zeta potential, and conductivity (Table 5-2).

The microemulsion reported by Attwood et al.210 contained IPM (4.7 %w/w), polysorbate

Tween 80 IPM Sorbitol (70%w/v) Water0.01

0.1

1

10

100

So

lub

ilit

y o

f d

iaze

pam

(m

g/m

L)

IPM

Sorbitol (70%w/v)

Tween 80

Water

0.05

64.55

0.19

10.23

137

60 (13.3 %w/w), sorbitol (26.7 %w/w) and water (55.3 % w/w), which resulted in

dispersed phase diameter of ~29 nm. In the current study, for the blank microemulsion

(without diazepam), the weight percent of surfactant: sorbitol was kept at 40 %w/w and

the mass ratio of surfactant: sorbitol was changed from 1: 2 (polysorbate 60: sorbitol) to

1: 2.5 (polysorbate 80: sorbitol), and the IPM content was slightly increased (5 %w/w).

The mean diameter of the dispersed phase in the control increased to ~ 48 nm. This

increase in size may be attributed to: 1) the greater mass ratio (1:2.5) of polysorbate

80/sorbitol (40% w/w) compared to the 1:2 polysorbate 60/sorbitol (40% w/w); 2)

polysorbate 80 may also occupy a greater area per molecule compared to polysorbate 60

thus increasing the droplet size; 3) Attwood et al. used a time-averaged light scattering

technique to determine droplet size. This technique measures the size using diffraction

principles, whereas in the current study, the size of the dispersed phase was determined

using a photon correlation spectroscopy autocorrelation function calculated based on the

Brownian motion of the particles; 4) the viscosity and refractive index of the

microemulsion was not accounted for in the particle size analysis and the autocorrelation

function depends on accurate values for these parameters to determine the correct particle

size. Since it is likely that the viscosity of the microemulsion was greater than water, the

size of the dispersed phase may have been over-predicted.

Inclusion of diazepam in the microemulsion resulted in a slight decrease in

dispersed phase diameter to ~42 nm. The zeta potential of blank and diazepam-containing

microemulsions were found to be ~ 0 mV due to the use of nonionic surfactant, Tween

80. No visual phase separation was observed after 7 days of storage at ambient

temperature (Figure 5.5). Since the chemical stability of the DZME formulation was

138

unknown, all studies were performed with DZME formulations stored for not more than

24 h.

Table 5-2 Size and zeta potential of microemulsions with and without drug and 2,4-DNP (mean ± std dev).

Formulation Droplet Size (nm) Zeta potential (mV)

Blank microemulsion 41.9 (9.8) -0.2(2.3)

Diazepam containing microemulsion 47.5 (4.5) -0.4 (3.2)

Diazepam containing microemulsion with 2,4-DNP

39.6 (10.2) 0.3 (4.2)

Figure 5.5 Appearance of diazepam-containing microemulsion.

139

Microemulsion Uptake Studies

The mass transfer of diazepam from the microemulsion and from an IPM solution

across both respiratory and olfactory tissues is shown in Figure 5.6. The transfer of

diazepam across both tissues appeared to be similar with only negligible quantities of

diazepam (<0.5% of administered dose) transferred across the tissues to reach the

receiver chamber after 120 min. Most of the diazepam (75 % - 90 %) remained in the

donor chamber after 120 min of exposure to the tissue explants (Figure 5.7). There was a

decrease of diazepam content (10 % - 25 %) in the donor chamber, but negligible

diazepam (<1%) was recovered from the receiver chamber. The unaccounted diazepam in

these experiments (15 % - 25 %) may be the result of accumulation within the tissues or

may be the result of experimental error. When the transfer of diazepam from DZME was

tested using an impermeable, artificial membrane, ~ 100% of the diazepam remained in

the donor chamber after 120 min of exposure (Figure 5.7), with negligible transfer to the

receiver chamber. As the total mass of loaded diazepam was recovered from the

impermeable membrane study, it appears unlikely that the experimental loss of diazepam

during the transport study was significant. As a result, any unaccounted for diazepam in

the tissue transport studies was assumed to accumulate within the tissues.

The unaccounted for diazepam was determined by taking the difference between

the initial diazepam in the donor fluids and the concentration of the diazepam recovered

from the donor and receiver chambers. Figure 5.8 shows the percent of unaccounted for

diazepam, which was assumed to accumulate within the respiratory and olfactory tissues.

140

A.

B.

Figure 5.6 Comparison of the cumulative percent of diazepam (relative to the donor chamber initial concentration) appearing in the receiver chamber as a function of time across bovine respiratory and olfactory tissue explants following exposure to DZS (diazepam IPM solution, 4 mg/mL) and DZME (diazepam-containing microemulsion, 4 mg/mL). A) Data shown are mean ± standard deviation (n=6 per tissue type). B) Mean percent diazepam transferred to receiver results shown without error bars.

0 50 100 1500.00

0.05

0.10

0.15

0.20

0.25

Time (min)

Perc

en

t D

iazep

am

Tra

nsfe

red

to R

eceiv

er

(%) DZME Respiratory

DZME Olfactory

DZS Respiratory

DZS olfactory

0 5 0 1 0 0 1 5 0

0 .0 0

0 .0 5

0 .1 0

0 .1 5

0 .2 0

0 .2 5

T im e (m in )

Pe

rc

en

t D

iaz

ep

am

Tra

ns

fere

d

to R

ec

eiv

er (

%)

141

Figure 5.7 Comparison of diazepam percent remaining in the donor chamber after 120 min exposure of diazepam IPM solution (DZS) and diazepam-containing microemulsion (DZME) to the respiratory, olfactory and artificial membrane. Dashed line represents 100%. Data shown are mean ± standard deviation (n=6 per tissue/membrane type).

Figure 5.8 Comparison of diazepam percent (relative to the initial diazepam concentration in the donor chamber) accumulated in the respiratory and olfactory tissues after 120 min exposure to diazepam-IPM solution (DZS) and diazepam-containing microemulsion (DZME). Data shown are mean ± standard deviation (n=6 for each tissue type). * Indicates a statistically significant difference when compared using an unpaired, two-tailed Student’s t-test with p<0.05.

Res

pirato

ry

Tissu

e

Olfa

ctory

Tissu

eArtifi

cial

Mem

brane

0

50

100

150

% o

f D

iaze

pa

m

Re

ma

inin

g in

Do

no

r

DZS

DZME

Res

pirat

ory

Tissu

e

Olfa

ctory

Tissu

e

0

10

20

30

40

% o

f D

iaze

pa

m

in T

iss

ue

[by

diffe

ren

ce

]

DZS

DZME

* *

142

Although the tissues were exposed to equivalent diazepam concentrations, the

uptake of diazepam into the tissues from DZME was ~2-fold higher compared to uptake

from the diazepam-containing IPM solution. The increased uptake of diazepam from

microemulsion suggests that the components of the microemulsion may act as permeation

enhancers. There are reports that suggest that the dispersed phase in a microemulsion can

act as a reservoir for hydrophobic drugs which helps to maintain a constant concentration

gradient across the membrane182-183. For example, in a study by Patel et al.182, the authors

claimed that increased uptake of risperidone across sheep nasal mucosa from a

microemulsion compared to drug in solution was due to maintenance of constant

concentration gradient of drug across membrane where the dispersed phase acted as

reservoir of the drug. However, while the authors used a mathematical model to describe

the mechanism, no direct evidence of the mechanism of drug uptake from microemulsion

was provided.

The influence of structural differences between the bovine respiratory and

olfactory tissues may also influence the permeation of diazepam across these tissues.

Similar amounts (~25%) of diazepam were accumulated in these two different tissues. It

is well understood, however, that the two tissues are morphologically different, especially

their thicknesses. The respiratory tissue is ~1.6-fold thicker than the olfactory tissue

(average thickness of bovine respiratory mucosa: 0.096 cm, average thickness of bovine

olfactory mucosa: 0.059 cm). When the amount of diazepam permeated across each

tissue was normalized to the tissue thickness (Figure 5.9), due to the high variation of the

mass determined for each tissue, no statistical differences between the tissues could be

observed. In comparison, in Chapter 3 it was shown that quantum dots (QDs)

143

accumulated to a greater extent in olfactory tissues, and the higher uptake of QDs into the

olfactory tissues was attributed to the involvement of different uptake pathways for QDs

compared to those utilized by microemulsions.

Histological Evaluation

Images obtained using brightfield microscopy of hematoxylin and eosin stained

sections of bovine respiratory and olfactory mucosae are shown in Figures 5.10 and 5.11.

Two hours of exposure to microemulsion lead to damage to the epithelial layer of both

the respiratory and the olfactory mucosae when compared to the control tissues. The

submucosal layer did not show any signs of alteration, however. The epithelial changes in

both tissue types may explain the similar levels of diazepam uptake in both tissues

(thickness normalized) since the epithelial layer is the primary barrier for the permeation

of the drug across the nasal tissues.

Figure 5.9 Comparison of diazepam accumulation in the thickness normalized olfactory and respiratory tissue explants after 120 min exposure to a diazepam IPM solution (DZS) and a diazepam-containing microemulsion (DZME). Data shown are mean ± standard deviation (n=6 for each tissue type).

DZS

DZM

E

0

1000

2000

3000

4000

Am

ou

nt

of

Dia

zep

am

in

Th

ickn

ess N

orm

alized

Tis

su

e (

mg

)

[by d

iffe

ren

ce]

Respiratory Tissue

Normalized Olfactory Tissue

144

Figure 5.10 Brightfield microscopic images of control (left) and DZME exposed (right) respiratory tissue explant. Solid line arrow shows intact epithelium in control tissue and the dashed-line arrow shows the damaged epithelium in DZME-exposed respiratory tissue.

Figure 5.11 Brightfield microscopic images of control (left) and DZME exposed (right) olfactory tissue explant. Solid line arrow shows intact epithelium in control tissue and the dashed-line arrow shows the damaged epithelium in DZME-exposed olfactory tissue.

Mechanism of DZME Uptake into Nasal Tissues

Drug permeation from a microemulsion is a complex process involving several

pathways. Possible mechanisms of drug permeation from surfactant rich microemulsions

include passive diffusion of free drug from the continuous phase and uptake of the drug-

containing dispersed phase using vesicular uptake mechanisms. Diazepam, being a

145

hydrophobic molecule, has limited ability to access paracellular transport pathways. It

has not been reported to be a substrate for any transporter systems, so the likely primary

mechanism for uptake would be transcellular passive diffusion. In order to further

investigate whether any energy-dependent processes were involved in the uptake of

diazepam in the nasal tissue explants, diazepam uptake from DZME in the presence of

2,4-DNP was investigated (Figure 5.12). A slight decrease in diazepam uptake into the

tissues was measured in the respiratory tissues (p < 0.1) while no difference was observed

in diazepam uptake in the presence of 2,4 DNP in the olfactory tissues. These results

suggest that there may be additional, additional energy-dependent pathways that assist in

the uptake of diazepam from the microemulsion into the respiratory tissues. The

endocytic pathways would be the most likely energy-dependent mechanisms to be

involved; yet most comparison results to date suggest that the olfactory tissues are more

endocytically active than the respiratory tissues. Another potential pathway may be via

the paracellular route. In the presence of 2, 4 DNP, the junctional proteins are unable to

maintain their size-based regulation of the intercellular junctions. Hence, it suggests that

the drug transfer via passive diffusion through the transcellular route is likely the major

pathway in diazepam microemulsion uptake.

146

Figure 5.12 Comparison of diazepam accumulation in the respiratory and olfactory tissues after 120 min exposure to DZME in the presence and absence of the metabolic inhibitor 2,4 –dinitrophenol (2,4-DNP). Data shown are mean ± standard deviation (n=6 for each tissue type). * Indicates a statistically significant difference when compared using an unpaired two-tailed Student’s t-test with p<0.1.

In previous chapter 3, quantum dots were shown to internalize into the nasal

respiratory and olfactory tissues via multiple endocytic pathways (Chapter 3) along with

some energy-independent pathways in the olfactory mucosa. However, in the case of the

microemulsions, it is difficult to determine whether the fluid-phase oil droplets were

internalized via endocytic pathways, because the integrity of the tissue was compromised

and the endocytic processes were likely no longer operative (Figures 5.10 and 5.11).

Res

pirat

ory

Tissu

e

Olfa

ctory

Tissu

e

0

10

20

30

40

% o

f D

iaze

pa

m

in T

iss

ue

[b

y d

iffe

ren

ce

]

DZME*

DZME + 2,4-DNP

147

Conclusions

The diazepam-loaded, oil-in-water microemulsion formulation showed enhanced

drug transfer into both nasal olfactory and respiratory tissues. These results suggest that

microemulsions may improve the bioavailability of poorly water-soluble drugs

administered intranasally. However, formulators should be careful in the selection of the

components of the microemulsion system due to the potential to damage the mucosa with

high concentrations of surfactants and non-aqueous solvents. These studies showed that

drug transfer from a diazepam-containing microemulsion into bovine nasal tissues was

independent of tissue region and appeared to primarily involve energy-independent

pathways, likely passive diffusion. It is unclear if endocytosis of the fluid-phase

nanodispersions of oil droplets played a role in drug absorption from the microemulsions

in a manner similar to the uptake of solid-phase nanoparticles due to the significant loss

of the epithelial cell layer following exposure to the microemulsion formulation.

148

CHAPTER VI

CONCLUSIONS

Various types of nanomaterial have been shown to cross the nasal mucosa and

reach the brain or the systemic circulation. Exploiting the nasal route in delivering

therapeutic agents using colloidal dispersions presents a promising new strategy for

targeted or improved efficacy delivery systems. Despite the potential delivery

advantages, the percentage of the administered nanomaterials reaching their desired

targets is minimal. While it has been reported that the size and surface characteristics of

nanoparticles affect their translocation efficiency, there is still a significant knowledge

gap regarding how nanomaterials, especially ultrafine nanomaterials (< 20 nm) interact

with cells and specifically interact with the cells in the nasal mucosa following

administration of drug products or vaccines. Careful characterization of the uptake and

distribution of particles in the olfactory and respiratory tissues, subsequent transfer to the

brain, or the systemic vasculature, or the lymphatics is needed to identify particle

characteristics that can be leveraged in advanced drug delivery strategies.

The use of quantum dots (< 20 nm) as model ultrafine nanoparticles assisted in

the evaluation of the uptake mechanisms in bovine nasal respiratory and olfactory tissues.

The unique composition of QDs enabled the measurement of the concentration of QDs in

the nasal tissues using ICP-OES and their inherent optical properties also enabled their

visualization within the bovine nasal tissues using confocal and electron microscopy.

Based on these studies, it is suggested that ultrafine nanoparticles show greater uptake in

the olfactory tissues compared to the respiratory tissues. COOH-QDs showed

accumulation in both the epithelial and submucosal regions of the bovine nasal tissues.

149

The uptake pathways utilized by these two tissues were also found to be different. In

respiratory tissues, clathrin-dependent, macropinocytosis and caveolae-dependent

endocytosis processes were all involved in the uptake of QDs whereas in olfactory tissues

clathrin-dependent endocytosis was the major endocytic pathway utilized. Additional

energy-independent pathways also appeared to be active in the internalization of QDs

into the olfactory mucosa, however the effect of surface chemistry on these pathways still

requires further investigation.

Observations from in vivo biodistribution studies in mice following intranasal

administration of quantum dots suggest that extremely small nanoparticles are transferred

from the nasal cavity into the posterior turbinate region and subsequently in close

proximity to the olfactory bulb. The majority of the administered COOH-QDs appear to

remain in the deeper nasal regions for relatively long periods of time (up to 24 h). The

accumulation of nanoparticles in the nasal tissues was also dependent on the surface

characteristics of the nanoparticles. Both in vitro and in vivo studies demonstrated that

the internalization of PEGylated QDs was less than COOH-QDs.

Non-invasive in vivo biodistribution studies carried out in mice after intranasal

administration of quantum dots also showed that whole animal fluorescence imaging and

micro-CT techniques are useful tools in qualitative and semi-quantitative evaluation of

biodistribution in live animals. However, neither of these techniques was able to visualize

the discrete signals associated with individual nanoparticles in the brain or in other parts

of the body. Signal from aggregates of QDs was able to be visualized, however.

A novel attempt to study the effect of the physical state of the colloidal systems

on subsequent uptake was made using microemulsions. Unlike solid nanomaterials, it is

150

difficult to either visualize or measure oil droplets/dispersed phase of a microemulsion in

the nasal tissues. Hence, a diazepam-containing microemulsion was developed to

investigate the uptake mechanisms of microemulsions. The microemulsion system

showed similar permeation of diazepam into both the nasal respiratory and olfactory

tissues, unlike solid-nanoparticles (QDs) which showed greater uptake into the olfactory

tissues. Exposure of a diazepam-containing microemulsion to the bovine nasal tissues

resulted in damage to the epithelial barrier and may have contributed to the increased

permeation. Involvement of endocytosis for the internalization of fluid-phase oil

droplets/dispersed phase of the microemulsion for enhanced permeation of diazepam

across bovine nasal tissues is difficult with a surfactant rich microemulsion which caused

damage to the epithelial barrier.

Overall, these studies have increased the fundamental understanding of the

ultrafine nanoparticle uptake in nasal tissues and their biodistribution in the whole body.

While ultrafine nanoparticles may have limited application in the development of

efficient drug delivery systems, due to limitations of low drug loading efficiency and

potential toxic effects, these results contribute to the rational development of

nanoparticulate drug delivery strategies investigating the nasal and other routes of

administration.

151

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APPENDIX-A

Technical Specifications and Optical Spectrum of COOH-QDs

Table A-1 Specifications of COOH-QDs provided by NN-Labs, LLC (Fayetteville, AR). Catalog # CZW-R.

Composition Cadmium Selenide / Zinc sulfide

Stabilizing ligand Carboxylic acid ligand (mercaptoundecanoic acid)

Organic impurities <1% (not including ligands)

Solvent Water

Concentration 1 mg/mL

Cd content Each mg of QD contains 0.350 mg of Cd

Absorption and emission spectrum See Figure A-1

Figure A.1 Typical UV-Vis absorption (a) and emission (b) spectrum of COOH-QDs purchased from NN-Labs, Inc (Fayetteville, AR), Lot # LW074414A21104.

170

APPENDIX-B

Particle Size Distribution of 0.05 mg/mL COOH-QD and PEG-QD

Dispersion in KRB

Figure B.1 Representative particle size distribution of 0.05 mg/mL COOH-QDs (Lot# LW074414A21104) in KRB using a volume-weighted distribution analysis (Nicomp Particle Sizer, Model 380 ZLS, Santa Barbara, CA).

171

Figure B.2 Representative particle size distribution of 0.05 mg/mL PEG-QDs (Lot# LW134414A23108) in KRB using a volume-weighted distribution analysis (Nicomp Particle Sizer, Model 380 ZLS, Santa Barbara, CA).

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172

APPENDIX-C

Data Showing Efficiency of ICP-OES Measurements in Presence of Blank Tissues

Table C-1 Correlation between theoretical mass of Cd as added QDs to blank olfactory tissues and the Cd concentration measured from the digested samples of those tissues. QD dispersion (0.2 mL) spiked into known olfactory tissue weight and digested in 1 mL of nitric acid followed by dilution to 10 mL with DI water.

Concentration of QD spiked

(mg/mL)

Concentration of Cd spiked

(ng/mL)

Theoretical Mass of QDs

(μg)

Olfactory Tissue Weights

(g)


per g of Olfactory

Tissue (μg/g)

Measured concentration of Cd (ng/mL)

Measured Mass of QDs (μg)

Calculated Mass of QDs

per g of Olfactory

Tissue (μg/g)

0.01 70 2 0.0901 22.4 69.2 1.98 22.0

0.02 140 4 0.0608 69.7 136.7 3.90 67.7

0.05 350 10 0.0879 122.5 349.2 9.98 121.7

0.1 700 20 0.0586 347.4 698.8 19.96 346.7

From manufacturer specification each mg of QD contains 350 μg of Cd

Calibration curve equation using known standards of Cd was y=4.5567x, r2=0.994

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173

Table C-2 Correlation between theoretical mass of Cd as added QDs to blank respiratory tissues and the Cd concentration measured from the digested samples of those tissues. QD dispersion (0.2 mL) spiked into known respiratory tissue weight and digested in 1 mL of nitric acid followed by dilution to 10 mL with DI water.

Concentration of QD spiked

(mg/mL)

Concentration of Cd spiked

(ng/mL)


(μg)

Respiratory Tissue Weight

(g)


per g of Respiratory

Tissue (μg/g)

Measured concentration of

Cd (ng/mL) Measured Mass

of QDs (μg)

Calculated Mass of QDs

per g of Olfactory

Tissue (μg/g)

0.01 70 2 0.0906 22.2 69.7 1.99 22.1

0.02 140 4 0.1320 31.5 140.2 4.00 31.6

0.05 350 10 0.1073 93.2 346.6 9.90 92.3

0.1 700 20 0.1240 162.7 699.6 19.99 162.7


Calibration curve equation using known standards of Cd was y=4.5567x, r2=0.994

174

APPENDIX-D

Sample Calculation of Mass of QD from Measured Cd Concentration

Example:

Sample preparation for donor sample after 30 min exposure to respiratory tissue

0.2 mL of the donor sample was digested with 1 mL 70% of HNO3 for 24 h at 80 °C

followed by dilution to 5 mL with deionized water.

The dilution factor is 5.2/0.2=26

Intensity of Cd measured using ICP-OES = 1188

Calibration curve equation using known standards of Cd is y=2.277x with r2 = 0.9999.

The unknown concentration of Cd in the digested sample = 1188/2.277 = 521.7 ng/mL

Amount of Cd in donor sample (ng) = Measured Conc. of Cd (ng

mL) ∗ Dilution Factor

= 521.7 (ng

mL) ∗ 26 = 13565.2 ng = 13.56 μg


Mass of QD in donor chamber (mg)

=𝐴𝑚𝑜𝑢𝑛𝑡 𝑜𝑓 𝐶𝑑 𝑖𝑛 𝑑𝑜𝑛𝑜𝑟 𝑐ℎ𝑎𝑚𝑏𝑒𝑟 (μg) ∗ 𝑚𝑎𝑠𝑠 𝑜𝑓 𝑄𝐷 (𝑚𝑔)

350 μg of Cd

=13.56 μg ∗ 1 mg

350 μg= 0.0387 mg

Thus amount of QD measured in the donor sample after 30 min of exposure to respiratory

tissue is calculated to be 38.7 μg

Similarly for second and third replicates amount of QD remained in donor chamber was

calculated to be 36.1 and 37.4 μg respectively.

The average of three replicates is 37.4 μg with standard deviation of 1.3 μg (Table 3-3)

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175

APPENDIX-E

TEER Measurements Representing Respiratory and Olfactory Tissue Integrity

Table E-1 TEER values across respiratory and olfactory tissues exposed to COOH-QD dispersion measured at the beginning and end of the transport study for each time point. Values represent maintenance of tissue integrity before and after the transport.

Tissue Type Time (min)

TEER (Ω.cm2)

Experiment 1 Experiment 2 Experiment 3 Mean (std. dev)

Beginning End Beginning End Beginning End Beginning End

Respiratory

30 225 236 292 307 284 332 267 (37) 292 (50)

60 249 259 276 274 230 233 252 (23) 255 (21)

120 247 274 360 383 264 315 290 (61) 324 (55)

Olfactory

30 190 180 193 182 261 255 215 (40) 206 (43)

60 265 266 210 203 220 219 232 (29) 229 (33)

120 202 195 211 189 264 255 226 (34) 213 (36)

176

APPENDIX-F

Sequential Fluorescence Images from Individual Mice after Intranasal

Administration of COOH-QDs

Figure F.1 Fluorescence images co-registered with x-ray images from individual mice showing the distribution of COOH-QDs after intranasal administration. The red color represents the fluorescence signal from COOH-QDs.

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Figure F.2 Fluorescence images co-registered with x-ray images of euthanized individual mice with exposed nasal cavities observed 24 h after intranasal administration of COOH-QDs. The red color represents the fluorescence signal from COOH-QDs.

178

Sequential Fluorescence Images from Individual Mice after Intravenous

Administration of COOH-QDs

Figure F.3 Fluorescence images co-registered with x-ray images from individual mice showing the distribution of COOH-QDs after intravenous (retro-orbital injection) administration. The red color represents the fluorescence signal from COOH-QDs. Mouse 2 died after the 2 h time point.

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Figure F.4 Fluorescence images co-registered with x-ray images of euthanized individual mice with exposed nasal cavities observed 24 h after intravenous administration of COOH-QDs.

180

Sequential Fluorescence Images of Individual Mice after Intranasal

Administration of PEG-QDs

Figure F.5 Fluorescence images co-registered with x-ray images from individual mice showing the distribution of PEG-QDs after intranasal administration. The red color represents the fluorescence signal from PEG-QDs.

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Figure F.6 Fluorescence images co-registered with x-ray images of euthanized individual mice with exposed nasal cavities observed 24 h after intranasal administration of PEG-QDs.

182

Sequential Fluorescence Images of Individual Mice after Intranasal

Administration of COOH-QDs in the Presence of an Endocytic Inhibitor

Cocktail

Figure F.7 Fluorescence images co-registered with x-ray images of individual mice showing the distribution of COOH-QDs in the presence of an endocytic inhibitor cocktail following intranasal administration. The red color represents the fluorescence signal from COOH-QDs.

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Figure F.8 Fluorescence images co-registered with x-ray images of euthanized individual mice with exposed nasal cavities observed 24 h after intranasal administration of COOH-QDs in the presence of an endocytic cocktail inhibitor. The red color represents the fluorescence signal from COOH-QDs.

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