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General Method for Producing Biodegradable Nanoparticles and Nanofibers Based on Nanoporous Membranes

Permanent Link: http://ufdc.ufl.edu/UFE0042515/00001

Material Information

Title: General Method for Producing Biodegradable Nanoparticles and Nanofibers Based on Nanoporous Membranes
Physical Description: 1 online resource (127 p.)
Language: english
Creator: GUO,PENG
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2011

Subjects

Subjects / Keywords: BIODEGRADABLE -- NANOFIBERS -- NANOPARTICLES -- NANOPOROUS
Chemistry -- Dissertations, Academic -- UF
Genre: Chemistry thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy GENERAL METHOD FOR PRODUCING BIODEGRADABLE NANOPARTICLES AND NANOFIBERS BASED ON NANOPOROUS MEMBRANES By Peng Guo May 2011 Chair: Charles R. Martin Major: Chemistry Biocompatible and biodegradable nanostructured materials have attracted more and more attention since they offer numerous exciting possibilities in medical sciences, such as drug delivery and tissue engineering. The increasing need for novel drug delivery systems with enhanced specificity/activity and reduced side toxicity has led to the development of nano-sized drug vehicles, which provide the advantage of delivering small molecular drugs, as well as macromolecules, via both targeted delivery and controlled release. For tissue engineering, considerable effort has been made to develop three dimensional artificial nanofibrous scaffolds, which closely resemble the natural protein nanofiber network in the extracellular matrix (ECM) . The goal of this thesis is to develop a simple and efficient method to produce nanostructured biomaterials for drug delivery or tissue engineering applications. We present here a novel strategy, referred to as nanopore-injection (N-I) method, based on the use of a nanoporous membrane that separates the feed and receiver solutions. By pumping one solution into the other, through the membrane, one can generate nanostructured materials at the exits of the membrane nanopores. The first part of the dissertation (Chapters 2, 3, 4) involves the fabrication of biodegradable nanoparticles, including hydrophobic drugs and drug-encapsulated polymeric nanoparticles, as well as exploring their applications in drug delivery. The N-I approach was designed by either anti-solvent or pH-sensitive features, and the particle size was found to be affected by the flow rate and viscosity of the feed solution and the pore size of the membrane. These nanoparticles exhibit excellent biocompatibility and sustained release capabilities. The second part of the dissertation (Chapters 5, 6) focuses on the fabrication of biodegradable nanofibrous scaffolding and their applications in tissue engineering. By using a modified N-I setup, polymers and organic/inorganic hybrid nanofibers were generated with a controlled morphology and size. The obtained nanofibrous scaffolds could support stem cell proliferation and differentiation.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Statement of Responsibility: by PENG GUO.
Thesis: Thesis (Ph.D.)--University of Florida, 2011.
Local: Adviser: Martin, Charles R.
Electronic Access: RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2012-04-30

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2011
System ID: UFE0042515:00001

Permanent Link: http://ufdc.ufl.edu/UFE0042515/00001

Material Information

Title: General Method for Producing Biodegradable Nanoparticles and Nanofibers Based on Nanoporous Membranes
Physical Description: 1 online resource (127 p.)
Language: english
Creator: GUO,PENG
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2011

Subjects

Subjects / Keywords: BIODEGRADABLE -- NANOFIBERS -- NANOPARTICLES -- NANOPOROUS
Chemistry -- Dissertations, Academic -- UF
Genre: Chemistry thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy GENERAL METHOD FOR PRODUCING BIODEGRADABLE NANOPARTICLES AND NANOFIBERS BASED ON NANOPOROUS MEMBRANES By Peng Guo May 2011 Chair: Charles R. Martin Major: Chemistry Biocompatible and biodegradable nanostructured materials have attracted more and more attention since they offer numerous exciting possibilities in medical sciences, such as drug delivery and tissue engineering. The increasing need for novel drug delivery systems with enhanced specificity/activity and reduced side toxicity has led to the development of nano-sized drug vehicles, which provide the advantage of delivering small molecular drugs, as well as macromolecules, via both targeted delivery and controlled release. For tissue engineering, considerable effort has been made to develop three dimensional artificial nanofibrous scaffolds, which closely resemble the natural protein nanofiber network in the extracellular matrix (ECM) . The goal of this thesis is to develop a simple and efficient method to produce nanostructured biomaterials for drug delivery or tissue engineering applications. We present here a novel strategy, referred to as nanopore-injection (N-I) method, based on the use of a nanoporous membrane that separates the feed and receiver solutions. By pumping one solution into the other, through the membrane, one can generate nanostructured materials at the exits of the membrane nanopores. The first part of the dissertation (Chapters 2, 3, 4) involves the fabrication of biodegradable nanoparticles, including hydrophobic drugs and drug-encapsulated polymeric nanoparticles, as well as exploring their applications in drug delivery. The N-I approach was designed by either anti-solvent or pH-sensitive features, and the particle size was found to be affected by the flow rate and viscosity of the feed solution and the pore size of the membrane. These nanoparticles exhibit excellent biocompatibility and sustained release capabilities. The second part of the dissertation (Chapters 5, 6) focuses on the fabrication of biodegradable nanofibrous scaffolding and their applications in tissue engineering. By using a modified N-I setup, polymers and organic/inorganic hybrid nanofibers were generated with a controlled morphology and size. The obtained nanofibrous scaffolds could support stem cell proliferation and differentiation.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Statement of Responsibility: by PENG GUO.
Thesis: Thesis (Ph.D.)--University of Florida, 2011.
Local: Adviser: Martin, Charles R.
Electronic Access: RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2012-04-30

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2011
System ID: UFE0042515:00001


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GENERAL METHOD FOR PRODUCING BIODEGRADABLE NANOPARTICLES AND NANOFIBERS BASED ON NANOPOROUS MEMBRANES By PENG GUO A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORID A IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2011 1

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2011 Peng Guo 2

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To my family, for thei r continued support and love 3

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ACKNOWLEDGEMENTS The work presented in this dissertat ion was conducted between August 2006 and May 2011 in graduate school at the University of Florida. I would like to acknowledge every individual who gives me help, advice, and encouragement. I would like to sincerely thank my advisor Prof. Charles Martin, who supports and guides me in both my research and life. Prof. Martin is always helpful and prepared to give me guidance. Besides research, I also matu red as an individual in many areas; in relationships, such as trusting coworkers, self-confidence, and desire of exploration. My future career and life have and will continue to significantly benefit from working with Prof. Charles Martin. I am very grateful to have had the opportunity to work in the Martin lab, where I met and collaborated with many fantastic students and postdoctora l fellows. Dr. Pu Jin and Dr. Jillian Perry helped me begin my thesis res earch. Both taught me many experiments and how to use instruments during my early gr aduate years. Dr. Jiahai (Jay) Wang also worked with me in the first three years and provided many valuable suggestions. Dr. Hitomi Mukaibo, Dr. Lloyd Horne, Dr. Lindsay Sexton, Dr. Kaan Kececi, Dr. Dooho Park, Dr. Fan Xu, Gregory Bishop, Funda Mira, W illiam Hardy and Li Zhao are group members who worked with me and provided valuable support and suggestions during my research. I also would like to sincerely thank Prof. Richard Zare at Stanford University. During my visiting study in the Zare lab between Ap ril 2009 and May 2011, Prof. Zare took me in as one of his own graduate st udents, generously helping and training me to pursue my research. I also thank all the Zare lab group members for their sup port in my visiting research at Stanford University. 4

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I also acknowledge our collaborators in Stanf ord Univeristy: Prof. Gerald Fuller and Dr. Michael Maas in Dept. of Chemical Engi neering; Prof. Fan Yu and Dr. Michael Keeney in Dept. of Bioengineering; Prof. Ramin Beygui and Dr. Evgenios Neofytou in Medical Center Line: Cardiothoracic Surgery; Prof. A. C. Matin and Matthew Sylvester in Dept. of Microbiology and Immunology; and Prof. Chri stopher Contag and Dr. Tobi Schmidt in Dept. of Pediatrics. Without these great mind s I could not accomplish close to as much. My family has provided the most important support during my research. I am very fortunate to have such a loving and supportive fa mily. I want to thank my lovely wife Jing Huang. During the long and sometimes challenging graduate study, Jing has always stood beside me, providing enc ouragement and support, through both the highs and lows. Her delicate care and tenderness sustained me through every step of my research. I owe much to her. I want to thank my mother Shuying Wang for taking care of me, her kindness and support never wavered. 5

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TABLE OF CONTENTS page ACKNOWLEDGEMENTS ...............................................................................................4 LIST OF TABLES ............................................................................................................9 LIST OF FIGURES ........................................................................................................10 ABSTRACT ...................................................................................................................14 CHAPTER 1 INTRODUCTION AND BACKG ROUND.................................................................16 Introduction .............................................................................................................16 Current Progress in Biodegradable Nanostructures ...............................................16 Biodegradable Na noparticles in Drug Delivery .................................................16 Biodegradable Nanofibers in Tissue Engineering .............................................21 Methods for Producing Biodegradable Nanoparticles and Nanofibers ....................24 Nanoparticle Formation Method .......................................................................24 Nanoprecipitation .......................................................................................24 Nano-Emulsion ..........................................................................................26 Ionic Gelation .............................................................................................28 Nanofiber Formation Method ............................................................................29 Electrospinning ..........................................................................................29 Phase Separation ......................................................................................31 Self-Assembly ............................................................................................33 Dissertation Overview .............................................................................................34 2 FORMULATING HYDROPHOBI C DRUG NANOPA RTICLE S...............................40 Aim ..........................................................................................................................40 Experimental ...........................................................................................................42 Materials ...........................................................................................................42 Formation of Hydrophobic Drug Nanoparticles .................................................42 Analysis of Nanoparticles by Electron Microscope ...........................................43 Dynamic Light Scattering (DLS) Measurement ................................................43 X-Ray Diffraction (XRD) Analysis .....................................................................43 Dissolution Rate Measurement ........................................................................44 Results and Discussion ...........................................................................................44 Perspective .............................................................................................................48 3 GENERAL METHOD FOR PRODUCING POLYMERIC NANOPARTICLES USING NANOPOROUS MEMBRA NES..................................................................55 6

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Aim ..........................................................................................................................55 Experimental ...........................................................................................................56 Materials ...........................................................................................................56 Formation of Ultrafine Chitosan Nanoparticles .................................................56 Characterization of Chitosan Nanoparticles .....................................................57 Encapsulation of Rhodamine 6G in Chitosan Nanoparticles ............................57 Results and Discussion ...........................................................................................58 Perspective .............................................................................................................61 4 BIODEGRADABLE POLYMERIC NANOPARTICLES AS DRUG DELIVERY VEHICLE ................................................................................................................68 Aim ..........................................................................................................................68 Experimental ...........................................................................................................70 Materials ...........................................................................................................70 Synthesis and Characterization of PLGA-PEG Diblock Copolymer ..................70 Formation of Drug Encapsulated Nanoparticles ...............................................71 Characterization of Drug Encapsulated Nanoparticles .....................................72 Sustained Release Study of Drug Encapsulated Nanoparticles .......................72 Fluorescent Microscopy Imaging ......................................................................72 In Vitro Cytotoxicity Study by Clonogenic Assay ..............................................73 In Vivo Cytotoxicity Study by Bioluminescence Imaging ..................................73 Results and Discussion ...........................................................................................74 PLGA-PEG/MCHB Nanoparticle ......................................................................74 Chitosan/luciferin Nanoparticles (CS/Luc NPs) ................................................77 Perspective .............................................................................................................79 5 FORMATION OF BIODEGRADABL E NANOFIBERS BY NANOPOROUS MEMBRANE...........................................................................................................86 Aim ..........................................................................................................................86 Experimental ...........................................................................................................87 Materials ...........................................................................................................87 Formation of Collagen Nanofibers ....................................................................87 Characterization of Collagen Nanofibers ..........................................................88 Isolation and Culture of Cardiac Stem Cells (CSCs) ........................................88 Scaffold Seeding ..............................................................................................89 Microscopy Fluorescent Imaging ......................................................................90 Bioluminescence Imaging (BLI) ........................................................................90 Statistical Analysis ............................................................................................91 Results and Discussion ...........................................................................................91 Perspective .............................................................................................................94 6 ORGANIC/INORGANIC HYBRID NANO FIBERS FOR TISSUE ENGINEERING...99 Aim ..........................................................................................................................99 Experimental .........................................................................................................101 7

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Materials .........................................................................................................101 Formation of Hybrid Nanofibers ......................................................................101 Analysis of Nanofibers by Electron Microscope ..............................................102 Stem Cell Preparation ....................................................................................102 In Vitro Cytotoxicity Study by Cell Titer Assay ................................................103 Fluorescent Microscopy Imaging ....................................................................103 Statistical Analysis ..........................................................................................103 Results and Discussion .........................................................................................104 Perspective ...........................................................................................................108 7 CONCLUSION S...................................................................................................114 LIST OF REFERENCES .............................................................................................116 BIOGRAPHICAL SKETCH ..........................................................................................127 8

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LIST OF TABLES Table page 2-1 Summary of the DLS analysis of hydrophobic nanoparticles. .............................54 3-1 Statistical size and encapsulation efficiency data for rhodamine 6G loaded chitosan nanoparticles. .......................................................................................67 4-1 DLS data for PLGA-PEG/MCHB and CS/Luc nanoparticles. ..............................85 9

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LIST OF FIGURES Figure page 1-1 Advantages of biodegradable nanopa rticles for drug deliver y............................37 1-2 (a) Scheme of nanofi ber network of natural ECM.118 (b) A typical SEM image of neural interconnect and ECM.119 Nerves and nerve bundles (yellow), ECM (red), and ganglion ce lls (bl ue)...........................................................................37 1-3 Illustration of fabricating bi odegradable nanoparticles through nano-emulsion method.120 (Reprinted with permission from Ref [120]; Copyright 2008 Elsevier .)............................................................................................................38 1-4 Illustration of fabricating biodegr adable nanoparticles through ionic gelation method.121 (Reprinted with permission from Ref [121]; Copyright 2004 Elsevier .)............................................................................................................38 1-5 Illustration of fabricating bi odegradable nanofibers thr ough electrospinning method.122 (Reprinted with permission from Re f [122]; Copyright 2008 Brill.)....39 2-1 Experimental set-up for the hy drophobic drug nanoparticl e preparation using nanoporous membrane. M1 Pressure meter. M2 Fl ow meter.............................49 2-2 Chemical structures of three hydrophobic compounds: silybin, beta-carotene, and butylated hydr oxytol uene............................................................................. 49 2-3 Photograph of a typical experimental setup........................................................50 2-4 SEM images of nanoporous membranes: anodized aluminum oxide (AAO) membrane with (a) 20 nm inlet and (b) 200 nm outlet........................................50 2-5 Photograph of 40 mg silymarin nanoparticles obtained within 20 min by using AAO nanoporous membrane. A penny se rves as a size marker........................51 2-6 SEM images of SM, BC, and BHT dr ug nanoparticles. a, d, and g are SM, BC, and BHT NPs via N-I method, respective ly; b, e, and h are SM, BC, and BHT NPs via SEDS, respectively; c, f, and i are untreated SM, BC, and BHT, respectively. The scale bar is 500 nm in all the figures ......................................51 2-7 Hydrodynamic diameters of (a) SM, (b) BC, and (c) BHT drug nanoparticles determined by DLS.............................................................................................52 2-8 Effect of flow rate on di ameter of the SM NPs obtained. ....................................52 2-9 XRD pattern of silymarin nanopar ticles and untreated silymarin powder............53 10

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2-10 Dissolution profiles for silymari n nanoparticles and untreated silymarin powder in PBS (pH 7. 4) at 37C. ........................................................................53 3-1 Method for producing chitosan nanoparticles by flow though a nanoporous membrane.......................................................................................................... 63 3-2 SEM images of nanoporous membrane s: (a) track-etched polycarbonate (PCTE) membrane with 10 nm pores; and anodized aluminum oxide (AAO) membrane with (a) 20 nm inlet and (c) 200 nm outlet........................................63 3-3 Typical TEM images of chitosan nanoparticles (CSNPs) prepared by using (a) the PCTE membrane; and (b) the AAO membrane. In these TEM images, the black area represents t he nanoparticle, and the grey area represents the backgroun d.........................................................................................................64 3-4 Comparison of size distributions of chitosan nanoparticles (CSNPs) prepared by using different nanoporous membranes determined by dynamic light scattering: (a) size of CSNPs obtained by PCTE membrane; and (b) size of CSNPs obtained by AAO memb rane. .................................................................64 3-5 Effect of solution flow rate on the diameter of the chitosan nanoparticle obtained. .............................................................................................................65 3-6 Effect of the viscosity of the ch itosan feed solution on the diameter of the nanoparticles obt ained.......................................................................................65 3-7 Typical TEM images of chitos an-rhodamine 6G nanoparticles prepared by using (a) the PCTE membrane and (b) the AAO membrane. In these TEM images, the black area represents the nanoparticle, and the grey area represents the backgr ound................................................................................. 66 3-8 Comparison of size distributions of chitosan-rhodamine 6G nanoparticles prepared by using different nanoporous membranes determined by dynamic light scattering: (a) PCTE memb rane; and (b) AAO membrane..........................66 4-1 NMR characterization of (a) PLGA (b) PLGA-PEG dibl ock copolymer..............80 4-2 Typical SEM image of PL GA-PEG/MCHB nano particle s....................................80 4-3 Hydrodynamic diameter of PL GA-PEG/MCHB NPs det ermined by DLS............81 4-4 In vitro sustained release profile of PLGAPEG/MCHB NPs...............................81 4-5 In vitro cytotoxicity st udy of PLGA-PEG /MCHB NP s..........................................82 4-6 Fluorescent image of PC-3 cell incubated with (a) PLGA-PEG/MCHB NPs, and (b) PLGA-PE G/CNOB NP s..........................................................................82 11

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4-7 Typical SEM image of CS/Luc nanopar ticles ......................................................83 4-8 Hydrodynamic diameter of CS/Luc NPs determined by DLS. .............................83 4-9 In vitro sustained releas e profile of CS /Luc NP s.................................................84 4-10 In vivo biotoxicity study of CS/ Luc NPs by biolumi nescence im aging.................84 5-1 Method for producing collagen nanof ibers by flowing though a nanoporous membrane.......................................................................................................... 95 5-2 Typical SEM images of collagen nanofibers prepared by using the PCTE membrane at (a) high magnificati on and (b) low m agnificati on...........................95 5-3 Typical TEM images of (a) a bundle of collagen nanofibers (b) a single collagen nanofiber. Inset is the relat ed selected area electron diffraction pattern (SAED pattern).......................................................................................96 5-4 Photograph of collagen na nofibrous scaffold prepared by N-I method in 2 h. A penny is used as a size mark er..........................................................................96 5-5 SEM images of (a) collagen nanofibers prepared by N-I method (b) collagen film prepared without nanopor ous membra ne....................................................97 5-6 Effect of nanopore size on the di ameter of the colla gen nanofibers...................97 5-7 Rheology study of co llagen nanofibrous scaffold................................................98 5-8 (a) Fluorescent microscope imaging of CSCs in (A) and (C) blank control (low and high magnification); (B) and (D) co llagen nanofibrous scaffold (low and high magnification). In fluorescent micr oscope images, bright area represents CSCs, and black area represents background. (b) bioluminescence image of CSCs proliferation on blank contro l and collagen nanofibr ous scaffold..............98 6-1 Experimental setup and proposed mo del for the formation of mineralized collagen fibe rs..................................................................................................110 6-2 (a, b) Unmineralized collagen fibers, (c, d) Mineralized collagen fibers (1 mM CaCl2), (e, f) Mineralized co llagen fibers (2.5 mM CaCl2) and (g, h) Mineralized collagen fibers (5 mM CaCl2). Inset images in (b, d, f, h) are SAED patterns..................................................................................................111 6-3 (a) Bundle of PAA/CaCO3 nanofibers (b) TEM mi crograph and SAED pattern of a PAA/CaCO3 (c) Flattened PILP droplets, (d) PAA/Calcium Phosphate nanofibers. ........................................................................................................112 6-4 Proliferation of hA DSC's on nanofibers. indicate statistical difference between groups at the sa me timepoi nt.............................................................112 12

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6-5 Alkaline phosphatase production fr om hADSC's cultured on nanofibers. indicates statistical differenc e at the same timepoint ........................................113 6-6 Fluorescent images of hADSCs cu ltured on nanofibers. The green indicates actin filaments while blue indicates cell nuclei. .................................................113 13

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Abstract of Dissertation Pr esented to the Graduate School of the University of Florida in Partial Fulf illment of the Requirements for t he Degree of Doctor of Philosophy GENERAL METHOD FOR PRODUCING BIODEGRADABLE NANOPARTICLES AND NANOFIBERS BASED ON NA NOPOROUS MEMBRANES By Peng Guo May 2011 Chair: Charles R. Martin Major: Chemistry Biocompatible and biodegradab le nanostructured materials have attracted more and more attention since they offer numerous exciting possibilities in medical sciences, such as drug delivery and tissue engineering. The increasing need for novel drug delivery systems with enhanced specificity/activity and reduced side toxicity has led to the development of nano-sized drug vehicles, which provide the advant age of delivering small molecular drugs, as well as macrom olecules, via both targeted delivery and controlled release. For tissue engineering, cons iderable effort has been made to develop three dimensional artificial nanofibrous scaff olds, which closely resemble the natural protein nanofiber network in t he extracellular matrix (ECM). The goal of this thesis is to develop a simple and efficient method to produce nanostructured biomaterials for drug deliver y or tissue engineerin g applications. We present here a novel strategy, referred to as nanopore-injection (N-I) method, based on the use of a nanoporous membrane that separates the feed and receiver solutions. By pumping one solution into the other, through the membrane, one can generate nanostructured materials at the exits of t he membrane nanopores. The first part of the dissertation (Chapters 2, 3, 4) involves t he fabrication of biodegr adable nanoparticles, 14

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including hydrophobic drugs and drug-encapsula ted polymeric nanoparticles, as well as exploring their applications in drug deliver y. The N-I approach was designed by either anti-solvent or pH-sensitive feat ures, and the particle size was found to be affected by the flow rate and viscosity of the feed solution and the pore size of the membrane. These nanoparticles exhibit excellent biocompatibil ity and sustained release capabilities. The second part of the dissertation (Chapters 5, 6) focuses on t he fabrication of biodegradable nanofibrous scaffo lding and their applications in tissue engineering. By using a modified N-I setup, polymers and organic/inorganic hybrid nanofibers were generated with a controlled morphology and size. The obtained nanofibrous scaffolds could support stem cell prolif eration and differentiation. 15

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CHAPTER 1 INTRODUCTION AND BACKGROUND Introduction The application of nanotechnolog y in medical sciences is changing the landscape of drug delivery and tissue engineering industry as a whole. 1-14 Nanostructured biomaterials, featuring a nanoscale morphology and size, exhi bit a wide range of advantages over the conventional biomaterials, such as high bioava ilability, improved cellular interaction, and specific designed functions. 1 It offers a promising solution to many difficulties in drug delivery and tissue engineering. 2,6 For example, a nano-sized drug vehicle has made significant progress in the delivery of conventionally undeliverable molecules, such as compounds with low water solubility and genetic biomolecules. 3-5 In tissue engineering, nanofibrous scaffolds ideally imitate the natur al extracellular matr ix (ECM) with cell behavior modulation functions. 7-14 In this thesis, we present a simple and efficient nanopore-injection (N-I) method to produce biodegradable nanoparticl es and nanofibers based on the use of nanoporous membranes, the function of which is to separate two liquids. 15 By pumping one liquid into the other, through the membrane, we can generate nanoparticles or fibers at the exit of the nanopore. We illustrated this technique for the preparation of a series of nanostructured biomaterials (e.g. chitosan, PLGA-PEG, and collagen) and also explored their applications in drug delivery and tissue engineering. Current Progress in Biodegradable Nanostructures Biodegradable Nanoparticles in Drug Delivery Drug delivery is the science that administe rs the drug compounds to improve their pharmaceutical and therapeutic properties Many drug delivery systems have been 16

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developed to improve drug rel ease, dissolution, biodistri bution and elimination; the delivery efficiency, convenience and safety could also be enhanced. 16 The developed drug delivery systems inclu de polymeric nanoparticles, 17 lipid particulate carriers, 18 drug loaded microspheres, 19 drug-polymer conjugates, 20 etc. Due to the advantages offered by sustained release and targeted delivery, biodegradable polym eric nanoparticle are currently under intense investigation. 17 Polymeric nanoparticles used for drug delivery are elicited as a solid particle with the polymeric matrix. The ther apeutic agent of interest is di spersed in this polymeric matrix with a preferable size between one na nometer to a few hundr ed nanometers. The use of polymeric nanoparticle as drug carriers began in the 1980s. In 1987, Aprahamian reported that polyalkylcyanoacrylate nanocapsu les could be used as a drug carrier via transmucosal passage in the small intestine. 21 Since then, various polymeric nanoparticles with different structures, co mponents and functions have been fabricated for drug delivery. 17,22-25 Compared with other drug carri ers, polymeric nanoparticles exhibit three major adv antages (Figure 1-1). First, polymeric nanoparticles demonstrate a higher surface area compared with conventional drug carriers, due to their nano scale size. This high surface area changes particle surface properties and the interactions with disperse phase, especially with respect to the dissolution rate. 26 More than 40 % of therapeut ic compounds are poorly water soluble, and their clinical usefulness greatly limited by their bioavailability. Formulating these hydrophobic drugs into a nanoparticle form could efficiently improve their dissolution rate, and eventually improve their performance. In the in vitro drug delivery study, the sub-cellular size of the particle improves its cellular uptake efficiency. 27 17

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For example, Desai reported that the Caco-2 cell uptake of 100 nm gold nanoparticles was 2.5-fold higher compared to that of 1 m microspheres and 6-fold higher compared to that of 10 m microspheres. Some cell lines (e.g. Hepa 1-6, HepG2, and KLN 205) express favored uptake of nanop articles over microspheres. 28 During in vivo drug delivery, nanoparticles can easily penetrate through fine capillaries into deep tissues, and cross biological barriers. The reason that nanoparticles coul d penetrate throughout the submucosal layers, whereas the microparticles are localized in the epithelial lining, is mainly due to their size difference. An impor tant research area that holds tremendous promise for treating diseases that affect the brain is using nanoparticles to deliver therapeutic agents across the blood-brain barrier (BBB). The BBB is an endothelial cell monolayer connected by tight junctions in t he brain capillaries for the separation of circulating blood and cerebrospi nal fluid. This barrier is tremendously challenging for therapeutics to cross. Devel opment of nanoparticulate vectors to provide efficient trans-vascular delivery of a ther apeutic drug for difficult-to-tr eat diseases like brain tumors could provide life-saving drugs and is under intense investigation throughout the world. Second, sustained release of therapeutic ag ent could be achieved by encapsulating it within different biopolymers. A variety of functional polymers could be used to encapsulate drug compounds, including naturally occurring (chitosan and serum) and synthetic polymers (polylactic acid (PLA) and polylactic-co-glycolic acid (PLGA)). By selecting different polymers, nanoparticles could gain specific functions, such as sustained or triggered release. Drug molecu les are physically dispersed into the polymeric matrix of the nanoparticle, and the release duration of drug molecule from these nanoparticles could either be over a long duration of time with a controlled manner 18

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or triggered by outer environment al stimulus such as pH, tem perature, or electric field variance. For sustained release studies, t he drug encapsulated withi n the nanoparticle is released at a sustained rate, controlled by drug molecule diffusion and degradation of the polymer matrix. Therefore, the molecular weight of pol ymer, the functional groups and kinds of polymer are of great importance in the release durati on. The release duration of the drug molecule could vary from days to months. 29 It is preferred that the release of drug molecule in an in vivo environment maintains a constant drug biodist ribution together with an extended blood circulation time. Sustained release of drug molecule could also improve the drug performance and avoid repetitive uptake of drug, which would improve the quality of life of the many who take painful invasive treatments, such as needle injection based insulin delivery for diabetics. For particle materials, PLA and PLGA are two common biopolymers widely used for su stained release. Jacobson and co-workers reported that luciferin encapsulated inside the PLA nanoparticles showed an in vivo sustained release time of over 40 days. 29 In triggered release, polymer molecules are responsive to an environmental stimulus, and alter their polymer chain network morphology or structure accordingly, resulti ng in the drug molecules being released from the nanoparticle. Among many s ensitive biopolymers, chit osan is responsive to pH variance; PLGA-PEG-PLGA triblock polymer to temperatur e variance; and polyperryle (PPy) to electric field variance. Besides the above polymers, more triggered release polymers are being revealed. 30,31 Third, targeted drug delivery could be achieved by coupli ng targeting ligands to the drug encapsulated nanoparticle. Th is coupling allows the specific delivery of therapeutic agents to the tissue of interest bypassing ot her unrelated tissues. Currently, targeted 19

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delivery is one of the most challenging aspects facing drug delivery, and has tremendous significance for the highly toxic or carc inogenic drug delivery, which is commonly prescribed for the treatment of cancer. The majority of anti-cancer drugs are highly biotoxic to most kinds of cells, both canc erous and healthy. Targeted delivery of these drugs directly to the tumor site helps avoid eliminating normal tissue and thus would minimize the destructive side-effects of chemotherapy. A variety of tumor directed targeting ligands have been developed over the la st decade. For instance, chlorotoxin is a short protein ex tracted from the venom of the deathstalker scorpion and it selectively binds with the conductance chloride ionchannel on the glioma cell membranes. 32,33 Another, cyclic Arginine-Glycine-Aspartic (R GD) peptide is a recognition sequence of v3 integrin associated with tumor angiogenesis. 34,35 A number of synthetic and naturally o ccurring polymers have been used in polymeric nanoparticle formulati on. For drug delivery purpos e, there are generally two requirements for the polymer selection: fi rst, the polymer used for drug delivery is biodegradable; second, it needs to show non-biotoxicity and non-immunogenic response when interacts with cells. In the drug deliv ery process, the uptaken nanoparticles need to be eventually degraded and eliminated from the cell. Biopolym ers are usually degraded by enzymes (e.g. lysozyme) and cleared from the body through the citric acid cycle. On the other hand, non-removable nanoparticle vectors, such as carbon nanotubes and silica nanoparticles, will accumulate inside the patient body, which may cause cell damage and long-term damages to th e patient. This accumulation is currently considered as a serious health hazard in drug delivery. 36,37 The design of drug delivery nanoparticles should limit the toxic material in order to reduce the damage to healthy tissues. 20

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Commonly used biopolymers include synthet ic polymers such as polylactide polyglycolide copolymers, polyacrylates, polycaprolatones, and naturally occurring polymers such as chitosan, albumi n, gelatin, alginate, and collagen. 38-42 Among these different polymers, synthetic constructs nor mally show a longer sustained release time (several days to weeks) compared to their natural counterparts. But the application of synthetic polymer nanoparticles is usually lim ited by the harsh synthetic environment required in the particle formation process. Biodegradable Nanofibers in Tissue Engineering Tissue Engineering is defined as an interdisciplinary field that applies the principles of engineering and life sciences toward the dev elopment of biological substitutes that restore, maintain, or improve tiss ue function. by Langer and Vacanti in 1993. 6 The current application of nanotechnology in tissue engineering is to closely imitate nature. In human and animals, natural tissues can be c onsidered to exist as three components: cells, extracellular matrix (ECM) and signaling system. Majority cells are imbedded in the ECM, and the interaction between these ce lls and ECM regulates the cell migration, proliferation, differentiati on, gene expression, and secretion of various hormones and growth factors. Mimicking the natural ECM is of great impor tance for tissue engineering. Natural ECM exists as a network of nanofibers containing proteins and glycosaminoglycans, which comprise a th ree dimensional s patial and temporal environment for cell attachment and growth ECM also dynamically influences the phenotype by providing indirect or direct informational signaling cues. For example, the type I collagen molecule is essential for osteob last cell development into the bone marrow as it provides the integrin binding sites in the ECM. 43 21

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In tissue engineering, tremendous effort has been made to develop nanofibrous scaffold mimicking the natural ECM in chemic al composition, morphology and surface functional groups. The existence of these syn thetic nanofibrous scaffolds works as a temporary ECM for cells attach ment, location and growth. The mixture of cells within the scaffold will be implanted into the target tissue site inside human or animals. Previously formed cell organizations in the scaffold c ontinue growing and eventually merge together with the natural ECM. Thus tissue repar ation and regeneration are possible. The synthetic nanofibrous scaffold will be degraded over a period of time. In order to produce an ideal scaffold, there are two requirements: first, similar to the biopolymer used in drug delivery, the polymer used in tissue engineering should be biodegradable; second, polymers used in tissue engineer ing should also be non-biotox ic and non-immunogenic. After implanting the scaffold in vivo, it must be gradually degraded and replaced by the natural ECM with tissue regeneration. If the scaffold is not biodegradable, it will permanently exist at the implant site and cause an inhomogeneous phase in the newly formed ECM. This phase hinders the biomechani cal properties and functions of the new generated tissue especially musculoskeletal tissue. 44 Nanofibrous scaffolds are designed to be implanted in vivo, so biocompatibility is a key factor to evaluate the performance of the scaffold. Both naturally occurring polymers and synthet ic polymers could be used to form a nanofibrous scaffold. In tissue engineering, natur al polymers are more extensively used due to their similarity to the natural ECM as we ll as their in vivo bi ocompatibility. Popular naturally occurring polymers include collagen hyaluronic acid, gelatin, silk and chitosan. 45 Among these naturally occurring polymers, collagen is considered the most 22

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promising biopolymer in tissue engineering because it is the primary component in the natural ECM. Collagen is a group of structure prot eins which feature a unique triple-helix composed of three polypeptide subunits. In t he natural ECM, there are more than 20 types of collagen. The natural isotypes of collagen differ from each other with respect to their non-helical components and the length of the triple-helix. Among the 20 types of collagen, type I collagen is the most abundant and predominant collagen in the natural ECM; usually found in bone and tendon, skin and other connective tissues. The type I collagen molecule is a triple-helical protein, 300 nm in length and 1.5 nm in diameter. Its triple-helix consists two identical 1 (I) chains and one 2 (I) chain. Under certain conditions, such as pH, temperature, or ionic strength variance, type I collagen could self-assemble into a macromolecular struct ure. Initially, type I collagen molecules will self-assemble into fibrils with a mean diameter of 36 nm. If these conditions persist, type I collagen fibrils will eventually form fibers and fiber bundles. 46-48 Collagen nanofibrous scaffolds have been advant ageous in mimicking the chemical and biological function of the natural ECM. For example, utilizing electrospinning, Rho reported that aortic smooth musc le cells could proliferate and infiltrate into the collagen nanofibrous scaffold a with fiber diameter of 100 nm (Type I collagen) and 250 nm (Type III collagen). 49 Both of the resulting type I and type III collagen nanofibers display a characteristic 67 nm banding structure. Venugopal reported a poly( -caprolactone) (PCL) coated collagen scaffold with similar mechanica l properties to that of skin and showed that human dermal fibroblasts could prolif erate on this scaffold for dermal tissue reparation. 50 23

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On the other hand, organic/inorganic hy brid nanofibrous scaffolds (e.g., nanohydroxyapatite/collagen) are very popular for hard tissue engineering, especially in bone or dental tissue reparati on and regeneration. These hy brid nanofibrous scaffolds show a higher mechanical strength compar ed to the scaffold composed of only biopolymers. Calcium phosphate and calcium ca rbonate are widely used in the formation of hybrid nanofibers because they exist as the major inorganic materials found in the nature tissues. In addition, carbon nanotubes hav e also been studied to incorporate into the polymeric scaffold to construct functional sca ffold due in large part to their mechanical strength as well as electrical conductivity. 51 Methods for Producing Biodegradable Nanoparticles and Nanofibers In the last three decades, extensive effort has been devoted to fabricating biodegradable nanostructure. A considerable number of nanofabrication methods have been developed to formulate polymeric nanoparti cles for drug delivery and nanofibrous scaffolding for tissue engineeri ng. In this section, we present a overview of these conventional nanofabrication methods, payin g close attention to the advantages and disadvantages in order to provide an unbi ased evaluation of our nanopore-injection technique. Nanoparticle Formation Method Nanoprecipitation The nanoprecipitation method (or solvent displacement method) was first introduced by Fessi and co-workers in 1989. 52 This one-step preparation method immediately attracted wide att ention due to its simplicity, s peed, and economic feasibility. Nanoprecipitation allows the preparation of nanoparticles from preformed polymer such as PLA, PLGA, and PMMA, instead of t he respective monom er or oligomer. 24

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Nanoprecipitation systems encompass thre e parts: polymer; polymer solvent; and non-solvent of the polymer. The polymer is initially dissolved in the polymer solvent, and then the mixture solution is added into the non-solvent (usually aqueous solution) with magnetic stirring. Nanoparticles are formed by precipitation simultaneously with the polymer diffused into the non-solvent. The re sulting nanoparticle usual ly has a mean size of around 200 nm. In the nanoprecipitation proc ess, the selection of so lvent and non-solvent has a crucial impact on nanoparticle formation. The so lvent should have a high solubility of the polymer of interest; be misci ble in the non-solvent; and be facile to removable from the product following precipitation. Acetone and ethanol are two frequently used organic solvents in the nanoprecipitation process. If the polymer is not very soluble in organic solvents, multiple solvent blend could be used to improve the polymer solubility. Surfactant is often used in nanoprecipitat ion to stabilize the formed nanoparticles in the aqueous environment. Surfactants used in nanopr ecipitation are usually low molecule weight amphiphilic polymers (a molecule with a hydrophilic group on one end, and a hydrophilic group on the ot her end), which will spontaneously co ated on the surface of the polymeric nanoparticles with the hydrophobic end toward polymer surface and the hydrophilic end toward aqueous solution. Surfac tants improve nanoparticle dispersion in the aqueous phase, prevent parti cle aggregation and minimize surface charge. Common surfactants used in nanoprecipitat ion include Pluronic F68, Dext ran, Poly (vinyl alcohol) (PVA) and Tween 20 or Tween 80. A variety of biomaterials, including peptides and drugs, have been formulated into nanoparticles by nanoprecipitation. For exam ple, Govender prepared PLA and PLGA 25

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nanoparticles by nanoprecipitation and load ed the water-soluble drug procaine hydrochloride into these nanoparticles. 53 Memisoglu reported amphiphilic beta-cyclodextrin nanoparticles which encapsu lated the antifungal drugs bifonazole and clotrimazole. 54 Duclairoir and co-workers reported t he preparation of g liadin nanoparticles by nanoprecipitation. 55 The major weakness of nanoprecipitation is its dependence on materials that need to be hydrophobic to dissolve in the polymer so lvent. Clearly, this requirement limits the application of this method in formulating wate r soluble polymers and drugs. The existence of the organic solvent can cause the active therapeutic compounds such as peptides and nucleic acids to deactivate. Producing hydr ophilic polymers or drug nanoparticles by nanoprecipitation has been extensively probed. One promising solution, reported by Govender and co-workers, is to control the pH of the solvent in order to minimize the ionization of the hydrophilic polymers or drugs in the nanoprec ipitation process. 53 Nano-Emulsion In contrast to the nanoprecipitation met hod, the emulsion based nanofabrication method involves two steps for particle formation (Figure 1-3). The first step is the preparation of emulsified system and the second is the solidification of nanoparticles. The solidification of nanodroplets occurs by either precip itating the polymer (solvent evaporation, self-assembly of a macromolecule, and polymer ization of monomers) or spray drying. In the first step, the em ulsion system is usually formed by emulsifying two immiscible phases (such as water and oil); th e resulting emulsion droplets are stabilized by the surfactants. Several methods have been proposed that decrease the size of the emulsion droplet from microto nanoscale. One common method for decreasing the size 26

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is via a high energy (mechanical) process that forms the nano-emulsion. 56,57 High energy devices, such as rotor/stator devices, ultrasound generators, 58 and high-pressure homogenizers, 57 are introduced into the emulsion pr ocess to create higher interfacial areas resulting in nanoscale emulsion drople ts. Another popular stra tegy to reach the nano-level is a low energy method which gen erates nanoscale emulsion droplets by varying the intrinsic physicochemical properties of the surfactants and co-surfactants (which stabilize the emulsion droplets). For example, when polyethoxylated surfactants are used in the emulsion process, their par titioning coefficient is a function of the environmental temperature. By controlling the temperature, t he emulsion system will express a phase inversion and the bicontin uous system broken up into smaller droplets. 59 The second step in nano-emulsion is so lidifying the emulsion droplets into nanoparticles. There are several methods for accomplish this process: polymerization, solvent removal, or solvent evaporation. If the particle material is monomer, polymerization is the method most commonly utilized. In this polymerization process, monomer droplets are first st abilized by adsorbed surfactants in the nano-emulsion, then initiator molecules are introduced into the monomer matrix by premixi ng or just following nanodroplet formation. Subsequent ly, the radical polymerization reaction occurs within the monomer droplets by triggering initiator molecules with a specific stimulus, such as temperature, pH, UV, ultrasound, or enzyme. 60-63 If the particle materi al is a preformed polymer it is precipitated out by extracti ng the organic solvent from the nano-emulsion system. Several solvent extraction methods ex ist including solvent evaporation, fast diffusion after dilution and salting out. 64-67 27

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Ionic Gelation Ionic gelation is a simple and mild method to prepare polymer ic nanoparticles via complexation between oppositely charged macromolecules. In ionic gelation, charged polymers are first dispersed in the aqueous so lution and small ions with opposite charges are added into the same solution (Figure 1-4) Then the ionic nanogels are obtained from an aqueous solution by taking ad vantage of the electrostatic interaction between charged polymers and opposite charged ions. Ionic gelati on is an organic solvent free synthesis method, as the entire formation process happens inside the aqueous phase in contrast to that with the nanoprecipitation and nano-emulsion methods. In ionic gelation, polymeric nanoparticles are solidified and stabilized by electr ostatic interactions instead of chemical crosslink, which avoids the potential toxicity and other side effects from crosslinkers. A widely used gelling polymer in ionic gelation is chitosan, a cationic polysaccharide. Chitosan is positively charged in slightly acidic aqueous solutions due to the basic amine group. Chitosan nanogels are usually formed by addition of a polyanion, such as tripolyphosphate (TPP), into the chitosan soluti on. TPP provides electrostatic interactions with the positively charged polysaccharide and forms a spheric al complexation particle. The size of the complexation particle show s a strong dependence on the concentration of the corresponding cationic solu tion. Tokumitosu and co-workers reported the formation of chitosan-gadopentetic acid complex nanopartic les and explored their application in cancer therapy. 68 Polk successfully encapsulated album in into chitosan-alginate complex microcapsules by ionic gelation. 69 Drug encapsulated chitosan nanoparticles prepared by ionic gelation have demonstrated a uni que advantagenamely triggered release responding to either a pH varianc e or an ionic stimulus. This tr iggered release is attributed to the physicochemical property of ch itosan. The amine groups on chitosan are 28

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deprotonated by raising the pH an d neutralizing the solution. As a result, the electrostatic interactions between the oppositely charge d chitosan and TPP become weaken. In addition, the nanostructure of the chitosan nanopar ticle starts dissoci ating and swelling, causing the release of therapeutic agents tr apped within the polymeric matrix. This triggered drug release of thes e chitosan nanoparticle shows promise for the functional delivery of therapeutic agents. Nanofiber Formation Method Extensive research has been focused on developing three dimensional artificial scaffolds at the nanoscale level for tissue engi neering. Nanofibrous scaffolds are very popular due to its high similarity to the natural ECM. There three major nanofabrication methods for preparing nanofibers scaffold: el ectrospinning, self-assembly, and phase separation. Among these three methods, elec trospinning is commonly used to prepare aligned polymer nanofibers; self-assembly can provide insight into the nanofiber formation conditions and can produce very th in nanofiber; phase separation generates mesh nanofibrous scaffolds with a controllable pore size. Electrospinning Electrospinning was first patented in 1902 by Morton. 70 However, it was not until the 1980s that electrospinning became a popular nanof abrication technique, due in large part to the rapid development of nanoscience an d nanotechnology. Electrospinning is the most often employed method to formulate nanofibers from biomaterials. A general electrospinning setup is composed of th ree major components (F igure 1-5): a high voltage power supply, a capillary tube with a pipette or needle of small diameter, and a conductive receiver plate. In most elec trospinning cases, hypodermic needles are used as spinnerets, and aluminum foil as the receiver plate. The pr inciple of electrospinning is 29

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based on the electrostatic interaction between polymeric molecules. Electrospinning uses the electrostatic field to create and accelerate the liquid jet from a ti p of the capillary and obtains nanofibers via post-solidification. In the electrospinning process, the polymer is first dissolved in order to form a polymeric solu tion. This polymeric solution is filled in a capillary tube. Then a high voltage is applie d between the tip of capillary tube and the receiver plate. Charged polymer molecule s produce a repulsion force against each other in a direction opposite to the surface tension. When the applied voltage increases to the critical intensity, the repulsive force will overcome the surface tension and generate a polymer jet ejected from the tip of capillary tube. The so lvent in the polymer jet is evaporated in the air and eventually forms randomly oriented nanofi bers. The resulting nanofibers are collected by a grounded receiver plate. The fiber diameter is mainly controlled by the jet size and the polymer contents within the jet. In most of t he electrospinning experiments, a single jet is formed between the capillary and receiver plate 71-73 and the diameter of the nanofiber is related to the polymer concentration. Normally nanofibers wit h smaller diameter could be formed by using a low concentration polymer solution. 74 In addition, during je t transfers from the capillary to the receiver plate, one jet may be split into multiple sub-jets which reduce the diameter of the nanofibers. 74-76 Various polymers, both naturally occurring and synthetic, have been successfully electro-spun into nanofibers: Schreuder-Gibson and co-workers reported the formation of polycarbonate nanofibers by electrospinning; 77 Wang and Santiago-Aviles successfully synthesized the polyacrylonitrile (PAN) nanofibers by electrospinning; 78 Ding and co-workers obtained polyvinyl alcohol (PVA) nanofibers by electrospinning; 79 Zong 30

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reported the successful preparat ion of the naturally occurring polymer polylactic acid (PLA) nanofibers by electrospinning. 80 Although most of the pol ymers are dissolved in appropriate solvents before electrospinning, there are so me melted and electrospun into nanofibers, instead of in a solution. These polymers include polyeth ylene (PE), polypropylene (PP) and polyethylene teraphthalate (PET ). Electrospinning nanofi bers can create complex and interesting structures, such as beaded, ribbon, porous, and core-shell fibers. 81-83 Electrospinning has many shortcomings. One major problem is the sharkskin effect. 84 It is a phenomenon whereby the polymer jet is extruded during the spinning process, instead of into uniform thin nanofiber s, into spherical or spindle-like bead chains. In the electrospinning process, surface tensi on plays an important ro le in the bead chain formation. When the polymer jet is extruded to form very thin fi bers, the surface tension of the liquid jet tends to form spher ical droplets in order to mini mize its surface energy. This sharkskin effect has proved to be the most difficult obstacle to overcome in electrospinning. Phase Separation Phase separation was first reported by Ma and Zhang in 1999 (termed thermally induced liquid-liquid phase s eparation) for the formation of nanofibrous foam materials. 85 A typical phase separation process includes t he following steps: first dissolve polymer in appropriate solvent and make a homogeneous pol ymer solution with magnetic stirring; second the polymer solution is rapidly transferr ed into a refrigerator or freezer set at a certain temperature. Then phase separation and gelation w ould happen in the polymer solution. After the polymer solution gels comp letely, the polymer gel would be immersed into deionized water for solvent exchange, and water would replace the polymer solvent 31

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within the gel. The solvent ex change process usually takes more than two days. After that, the polymer gel would be lyophilized, and event ually a nanofibrous polymer matrix would be obtained. In this nanofibrous matrix, the resulting nanofiber s are similar to the natural protein nanofibers in the ECM (50-500 nm). In phase separation, several parameters may affect the final nanofiber formation. Gelation is the most important factor in c ontrolling the porosity of the nanofibrous scaffold. Gelation temperature normally affects the diameter of nanofibers. A mild gelation temperature (e.g. 22 C or room temperature) led to fine nano scale fibers. However, high gelation temperatures cause a platelet-like micron-sized gel due to polymer chain nucleation and crystal size growth. Polymer conc entration also has a significant influence on the nanofibers mechanical properties. Increas ing polymer concentration can improve the nanofiber mechanical property but will also decrease the porosity of the scaffold. Other factors that affect mo rphology need to be taken into a ccount; these factors include polymer type, solvent type, and thermal treatment. 86 The phase separation method c an fabricate three dimensi onal porous nanofibrous scaffolds which have potential applications in the tissue engineering. Ma and co-workers revealed that a nanofibrous scaffold formed by phase separation could process porosity as high as 98%. 85 In the three dimensional tissue scaff old, macroporosit y is an important structural parameter that allows cells to localize and proliferate wit hin the matrix and form more organized tissue, instead of growing on ly on scaffold surface. Macroporosity with pore size between 50 m is preferred in the three dimensional tissue scaffold. Macroporosity could be introduced by incor porating porogens such as sugar or salt micron crystals in the polymer solution duri ng the phase separation. Several studies have 32

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shown that these macroporous scaffold s could enhance cell attachment and distribution. 85-87 Compared with electrospinni ng, phase separation requires minimal experimental instrumentation. It could al so simultaneously preserve nanoand macro-sized structures within the scaffold greatly benef iting the development of novel three dimensional tissue scaffolds. The disadvantage of phase separation is its long process duration. It usually takes more than two days to finish one experiment and thus increasing the efficiency of phase separation is important. Self-Assembly Molecular self assembly is another impor tant pathway for polymeric nanofiber formation. Molecules spontaneously organize into stable and ordered nanofibers via non-covalent bands, such as hydrogen bonds, elec trostatic interactions, and/or van der Waals forces. 88 Although these non-covalent bonds ar e relatively weak compared with covalent bonds, the combination of seve ral non-covalent bonds provides a stable structure with fine mechanic strength. Among the various polymers, bi ological molecules, especi ally peptides and proteins, are of particular interest as the new buildi ng blocks in tissue engineering. This technique imitates the natural st ructural proteins self-assemble in to the natural ECM under in vivo environment. Phospholipids are amphiphilic compounds, composed of a cell membrane with other proteins via self-assembly. A num ber of naturally occurring or synthetic polymers have been formulated into nanofibers vi a self-assembly. For example, Malkar and colleagues successfully synthesized the triple-helix struct ure from a peptide amphiphile, which shares many features with the natural ECM. 89 Stupp and co-workers reported peptide nanofibers self-assembled fr om engineered peptide amphiphile under 33

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pH variance. 90 Aggeli demonstrated that beta sheet peptides could be transformed into a nanofiber structure via self-assembly. 91 In the self-assembly process, chirality of the peptides plays an im portant role in the nanofiber formation. For example, a right handed twist of a peptide chain in the beta stranded conformation usually leads to a le ft handed helical ribbon structure. The detailed mechanisms of these self-assembly nanofi bers are still under the investigations. 92 Nanofibers formed by self-assembly also show low polydispersity and good yield, however, self-assembly is a time consuming process. In summary, biocompatible and biodegrada ble nanostructured materials are widely expected to play an important role in the fu ture of medicine and the biological sciences. Nano-sized drug vehicles could improve drug delivery specificity/activity with much reduced side toxicity. For tiss ue engineering, the dev elopment of novel three dimensional artificial nanofibrous scaffolds becomes f easible due to the many new nanofabrication technologies. Professor Charles Martin at University of Florida pioneered a template fabrication technique for preparing nanostructures. A variet y of different nanostructures have been successfully prepared by ut ilizing nanoporous membranes. 93-117 We try to develop, using chemical intuition, a simple and effici ent fabrication technique for synthesizing biodegradable nanopart icles and nanofibers. Along the way, we will explore the biomedical applications of t he obtained nanostructured materi als, paying close attention to drug delivery and tissue engineering. Dissertation Overview The goal of this thesis is to develop a simple and scalable method to produce nanostructured biomaterials for drug deliver y or tissue engineerin g applications. We 34

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present here a novel strategy termed nanopore-injection (N-I ) method, based on the use of a nanoporous membrane that separates the feed and receiver solutions. By pumping one solution into the other, through the membrane, we can generate nanostructured materials at the exits of the membrane nanopores. In Chapter 2, we successfully prepared hydrophobic drug nanoparticles using the N-I method. The resulting nanoparticles we re compared to nanoparticles produced via solution enhanced dispersion and supercritical fl uids system. The influence of flow rate on particle size was studied. The dissolution profile of hydrophobic drug nanoparticles was tested and compared with the untreated drug powder. In Chapter 3, ultrafine or ganic nanoparticles (size < 30 nm) were prepared using the N-I method. Low molecular we ight biopolymer chitosan was selected as the particle material. We also investigated the influence of flow rate and viscosity on size control of the particle formation. The fluorescent dye rhodamine 6G was encapsulated in the chitosan nanoparticle and the encapsulation rate was determined by fluorescent spectra measurements. Chapter 4 described the fabr ication of biodegradable polymeric, PLGA-PEG/MCHB and chitosan/luciferin, nanoparticles (~100 nm) using the N-I method and applications of these polymeric nanoparticles in drug delivery in vitro and in vivo. The capability of sustained release was investigated by fluor escent spectra meas urements. In vitro cytotoxicity of PLGA-PEG/MCHB nanoparticles was tested in the PC-3 cell line by a clonogenic assay. Bioluminescence imaging was performed on mice to evaluate the in vivo cytotoxicity of chitosan/luciferin nanoparticles. 35

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In Chapter 5, the N-I me thod was modified to produce biodegradable nanofibers. Type I collagen, a natural stru cture protein, was selected as the fiber material. The influence of nanopore size on fiber diameter was investigated using a PCTE membrane with different nanopore sizes. A scaffold co mprised of collagen na nofibers was prepared for a tissue engineering study. Biomechanica l strength of the obtained scaffold was characterized by rheology. Growth and pro liferation of cardiac stem cells was investigated on a collagen nanofibrous scaffold. In Chapter 6, organic/i norganic hybrid nanofibers were prepared using the N-I method for hard tissue engineering. Calc ium phosphate doped colla gen nanofibers were successfully produced with the characteristic banding structure found in the natural bone matrix. Synthetic polymer poly(acrylic acid) was also used to form hybrid nanofibers with calcium phosphate and calcium carbonate. Human adipose tissue derived stem cells were tested to proliferate and differ entiate the obtained nanofibrous scaffold. 36

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Figure 1-1. Advantages of biodegradabl e nanoparticles for drug delivery. Figure 1-2. (a) Scheme of nanofi ber network of natural ECM. 118 (b) A typical SEM image of neural interconnect and ECM. 119 Nerves and nerve bundles (yellow), ECM (red), and ganglion cells (blue). 37

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Figure 1-3. Illustration of fabricating biodegr adable nanoparticles through nano-emulsion method. 120 (Reprinted with permission from Ref [120]; Copyright 2008 Elsevier.) Figure 1-4. Illustration of fabricating biodegradabl e nanoparticles through ionic gelation method. 121 (Reprinted with permission from Ref [121]; Copyright 2004 Elsevier.) 38

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Figure 1-5. Illustration of fabricating biodegr adable nanofibers through electrospinning method. 122 (Reprinted with permission from Ref [122]; Copyright 2008 Brill.) 39

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CHAPTER 2 FORMULATING HYDROPHOBIC DRUG NANOPARTICLES Aim The clinic applications of hydrophobic comp ounds are greatly limited by their low bioavailability, as more than 40% of therapeutic compounds are poorly water soluble. 123,124 Considerable effort has been made to improve these hydrophobic drug performance. Among those researches, fo rmulating hydrophobic drugs into nanoparticle form is an efficient strategy to improv e the hydrophobic drug delivery, because drug nanoparticles feature a high surface area and sub-cellul ar size compared with conventional micro-sized drug powder. It has al so been proved that dissolution rate of hydrophobic drugs increased when they were formulated into nanoparticulate form, due to the increased contact area between drug and solvent. 125,126 Meanwhile, other studies suggest that the sub-cellular size of nanoparticl e leads to a better cell uptake at both in vivo and in vitro system. 127,128 Researchers studied t he use of nanoparticle in hydrophobic drug delivery as early as 1970s. 129 Tremendous progress has been made in the last decades during the surgi ng of nanoscience and nanotechnology. 130-133 Several methods have been developed to generate hy drophobic drug nanoparticles, such as nanoprecipitation, 134 nano-emulsion, 135 and ionic gelation. 136 However, drug nanoparticles with uniform small size (<100 nm) and well dispersibility have not yet been produced in a scalable manner. Herein, we developed a simple and effici ent technique for producing hydrophobic drug nanoparticles (NPs) via nanopore-injection (N-I) method (as shown in Figure 2-1). Our strategy is pumping the feed solution dissolved drugs pass through the nanoporous membrane into the receiver solution (in which drugs are insoluble), resulted in 40

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hydrophobic drug nanoparticles formed at the exit of nanopores. Three common hydrophobic therapeutic compounds: silymari n (SM), beta-carotene (BC) and butylated hydroxytoluene (BHT) were selected as model drugs for the demonstration of nanoparticle formation (Figure 2-2). Silymarin is a mixture of flavono lignans exacted from milk thistle, in which silybin is its major chemical constituent with hepatoprotective and anti-cancer clinical effect. 137 Beta-carotene, a terpenoi d compound, serves as a precursor to vitamin A in human and animal metabolism. 138,139 Butylated hydroxytoluene is an antioxidant widely used as food addictive. 140 All of three molecules are poorly soluble in water. Meanwhile, according to liter ature, these molecules show non-toxicity to human body, and are considered safe to be operated in the chemistry laboratory. 137-140 In order to better evaluate our nanopore in jection (N-I) method, we introduced a solution-enhanced dispersion by supercritical fluids system (SEDS), a major commercial formulation method currently used by the pha rmaceutics industry, as a comparison method. 141-143 Both of the hydrodynamic drug nanoparti cles prepared by N-I method and SEDS method were compared in t heir diameters, size distribut ions, and zeta potentials. The particle sizes prepared by N-I method ar e relatively small with a narrow size distribution. And the producing rate could be as high as 2 mg/min. Compared to other nanofabrication techniques, N-I method provides a simpler and more cost-effective mean to produce hydrodynamic drug nanoparticles. We be lieve that these sm all, uniform, and well dispersed drug nanoparticles could exhibit promising applications in the biomedical fields. 41

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Experimental Materials Silymarin (SM) and beta-carotene (BC) were purchased from MP Biomedicals, LLC (Solon, OH), and butylated hydroxytoluene (BHT) was purchased from Acros Organics (Geel, Belgium). Anodized aluminum oxide (AAO) membrane (20 nm in diameter) was obtained from Whatman, INC (Pi scataway, NJ). All other chem icals were purchased from Sigma-Aldrich (St. Louis, MO) in reagent grade and used as received. Formation of Hydrophobic Drug Nanoparticles In N-I experiment, the experi mental setup consists of two half U-tubes and a nanoporous membrane which is sandwiched between the tw o halves (see Figure 2-3). Commercially available anodized aluminum oxide membrane (AAO, Whatman Inc.) was selected as the nanoporous separa te with pore diameter of 20 nm (Figure 2-4). The area of AAO membrane exposed to these solutions was about 2 cm 2 The feed solution contained 25 mg of hydrophobic compound in 10 mL organic solvent (SM and BHT was dissolved in acetone, and BC was dissolved in an acetone/tetrahydrofuran (50/50, v/v) blended solution). The receiver solution was 10 mL phosphate buffered saline (PBS, pH 7.4) solution with 0.5 wt% Plur onic F68. One half of the U-tube was filled with 10 mL of feed solution, the other half was filled with 10 mL receiver solution. Gauge pressure was created by connecting a compressed air outlet with a pressure reduction valve to the feed solution side of the U-tube. Usually 2 psi air pressure was applied in a general nanoparticle formation. In this wa y, the feed solution is pumped in to the receiver solution according to the applied pressure. Vigorous magnetic stirring was used in the receiver solution side to help formed nanoparticle di sperse in the aqueous solution. Obtained 42

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nanoparticles were collected from the receiver solution by filtration, rinsed three times with deionized water, and dried in t he air at room temperature. In SEDS experiment, the exper imental process followed the protocol described in the literature. 141 Briefly, the hydrophobic drug co mpound solution (2.5 mg/mL SM, BC, BHT in DMSO) was injected (1 mL/min) th rough a nozzle with 250 mm internal diameter into a supercritical carbon dioxide (SC-CO2) in a speed of 150 g/min at 40 C and 100 bar. The drug compounds precipitated in the SC-CO2 as an anti-solvent to form nanoparticles. Analysis of Nanoparticles by Electron Microscope The morphologies of the obtained hydr ophobic drug nanoparticl es and untreated drug powder were characterized by a FEI X L30 Sirion SEM. Dry samples on carbon sticky tape were sputter-coated for 90 s at 15 mA with Pd/Au. Average nanopar ticle diameters of SM, BC and BHT were determined by measur ing 50 random nanoparticles in each SEM image. Dynamic Light Scatteri ng (DLS) Measurement Zetasizer Nano ZS (Malvern Instruments, Malvern, PA) was used to measure the hydrodynamic size, polydispersity index (PDI ), and zeta potential of the hydrophobic drug nanoparticles. In DLS measurement, obtained nanoparticles were dispersed in the PBS (pH 7.4) with 0.5 wt% Pl uronic F68 at a concentration of 100 g/mL. X-Ray Diffraction (XRD) Analysis Powder XRD data were recorded on a PANalytical X'Pert PRO X-Ray Diffractionmeter using filtered Cu K radiation ( =1.5406 ) at 45 kV and 20 mA. Data were recorded by step scan with a step size of 0.040 and a step time of 1.0 s. 43

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Dissolution Rate Measurement In dissolution profile experiment, 2 mg SM NPs and untreated silymarin powders were weighed accurately, and immersed in the 30 mL PBS at pH 7.4, respectively. The resulting solutions were rotated at 100 rpm, and the temperature of the PBS (pH 7.4) was maintained at 37 0.5 C. 2 mL of each samp le was withdrawn and centrifuged at 10,000 rpm for 5 min. The centrifuge supernatant solution was collect ed for the ultraviolet (UV) absorbance detection. The UV absorbance int ensity was performed at a wavelength of 325 nm using an Agilent 8453 UV-Visible Spectrophotometer A series of samples were measured at 15 min time interval for 4 h. Results and Discussion As shown in Figure 2-3, an experimental setup comprised a nanoporous membrane and two halves of U-tube containing feed soluti on and receiver solution separately. 2 psi pressured flow was achieved via the connec ted compressed air on the feed solution, which pumped the feed solution flowing into the receiver solution through the AAO membrane. The AAO membrane is 60 m thick and contains 20 nm cylindrical pores at the face of the membrane in contact with the feed solution (Figure 2-4). These pores run parallel to one another for approximately 2 m and then feed much larger (200 nm in diameter) pores that run par allel to one another through the remaining thickness of the membrane. The pore density of the AAO membrane at the entrance, (i.e ., in contact with the feed solution) is around 6 14 /cm 2 144 The receiver solution became turbid immediately after the pressure was appli ed, which indicated the formation of nanodroplets. These nanodroplets formed at the outlet of t he 20 nm nanopores, and were detached from the membrane by the transmemb rane flow and the cont inuous stirring flow. 44

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The dispersed nanodroplets were transformed to nanoparticles immediately due to the dissolution of the feed solvent in aqueous solu tion. No instances of clogging or sticking were found after the experiment. Under a constant air pressure of 2 psi, 40 mg of SM NPs could be obtained within about 20 min (as shown in Figure 2-5). The obtained hydrophobic drug NPs were im aged using a FEI XL30 Sirion scanning electron microscope (SEM), as well as the untreated drug powder. Figures 2-6 a, d and g show the typical SEM images of the SM, BC and BHT NPs obtained via N-I method. These NPs are spherical in diameter of 80-1 00 nm with narrow distribut ion. Figures 2-6 b, e and h show the SEM images of SM, BC, and BHT NPs prepared by SEDS method. Those NPs were in irregular shapes with different sizes. SM, BHT NPs by SEDS have a small diameter at around 20-40 nm and BC NPs by SEDS has a large diameter at around 200-500 nm. As a comparison, Fi gure 2-6 c, f, and i show t he morphology of untreated SM, BC, BHT, which were all in the form of irregular micro-sized powder. Hydrodynamic diameters of drug NP was also characterized by dynamic light scattering (DLS) measurement. The obtained DL S data were summarized in Table 2-1. The hydrodynamic diameters of the SM, BC and BHT NPs obtained by N-I method were 83, 105, and 132 nm, with a polydispersity i ndex (PDI) of 0.180, 0.238, and 0.234, respectively (Figure 2-7). As a comparison, the hydrodynamic diameters of SEDS NPs were 496, 546, and 288 nm with a PDI of 0.387, 0.512, and 0.474, respectively (data not shown). All hydrodynamic sizes of the N-I NP s are smaller than t hose of the SEDS NPs, and their size distributions are narrower. The reason could be attributed to the aggregation of the SEDS NPs, which resulted in the poor dispersibility in aqueous solution. In contrast, N-I NP s have uniform shape and size, and were well dispersed in the 45

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aqueous solution. The hydrodynamic diameters are slightly la rger than the sizes obtained by SEM, which might caused by the interact ion between the particles and surrounding solvent. It is necessary to point out that the hydrodynamic diameter plays an important role in the drug delivery, especially in the in vivo systems. Nanoparticles with hydrodynamic size range of 10-200 nm are generally considered to be optimal for intravenous injection. Previ ous studies in other particle systems has proved that bio-distribution of NPs is str ongly dependent on the NP hydrodynamic diameter. 127,128,145,146 The influence of flow rate of feed so lution on the particle formation process was investigated (shown in Figure 2-8). SM NPs wa s selected as the model NPs in this study, and the hydrodynamic sizes of SM NPs were measured by DLS. From the parallel experiments, it could be found that the flow rate of f eed solution has an impact on the obtained SM NPs size. While the flow rate of feed solution was varied from 0.03 mL min -1 cm -2 to 2 mL min -1 cm -2 by adjusting the pressure fr om the compressed air, the hydrodynamic size of SM NP s decreased from 150 to 83 nm. Considering the overlapped size from their error bar, we do not recognize it as a significant size decrease. It was also found that the smallest particle size (83 nm) was obtained at a flow rate flow rate of 1.5 mL min -1 cm -2 These data might not exhibit a clear and significant size decrease with the increase of feed solution flow rate. We can st ill estimate a slow decreased trend on the increased flow rate from the mean di ameter of the NI nanoparticles. A crystalline structure c hange in SM NPs was observed before and after treated with N-I method. The crysta lline structure of SMNPs wa s examined using powder XRD (Figure 2-9). Before N-I process, the untr eated silymarin powder was a semi-crystalline 46

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material, exhibiting some peaks of medium intensity together wit h a strong underneath scattering phenomenon. As for the crystalline porti on, five characteristic peaks show up at 14.6, 16.5, 19.5, 22.3 and 24.5 of 2 which is also found in other studies. 147-149 After N-I process, SM NP shows no crystalline peaks in the XRD spectra. This is because the resulting SM NPs were transferred into the amorphous phase durin g the N-I process. Our hypothesis is that the total time for sily marin precipitating into nanoparticle is too short to form ordered crystalline st ructure during the N-I experim ent. A calculated flow rate inside the nanopore is over hundreds micromet ers per second. Such a high flow rate resulted in great number of nanodroplets form ed at the exit of nanopore, which were detached due to the wall shear force by t he transmembrane flow and the continuous stirring flow, and subsequently transfo rmed into nanoparticles rapidly. As dissolution rate is a crucial factor in hydrophobic drug delivery, we studied the dissolution rate of the N-I NPs. Silymarin NPs was selected as a model compound in this study, and the resulting dissolution profile is compared with that of untreated SM powder. All the dissolution profiles are characteriz ed using UV adsorption spectrometer. Test conditions, including rotation rate, temperature of the medi um, and method used to obtain samples, were the same for both samples. Figure 2-10 shows the dissolution profile of SM NPs and untreated silymarin powder. The disso lution rate of SM NP s is significantly greater than that of untreated silymarin powder At 30 min, the di ssolve percentage of SM NPs reached more than 80% compared with that of untreated silymarin powder (~15%). As amorphous nanoparticles normally exhibit a faster dissolution, t he enhancement of the dissolution profile is believed to be attributed to the amorphous nature of SM NPs, as well as the increased surface area after N-I process. 47

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Perspective With the proper design and system contro l, we fabricate uniform hydrophobic nanoparticles (SM, BC and BHT NPs) with <1 00 nm size through nanopor e-injection (N-I) method. The obtained nanoparticles exhibit sm aller hydrodynamic diameter and better dispersibility in aqueous solution, compar ed with those made thr ough SEDS method. Due to the rapid precipitation, the obtained N-I NPs were amorphous, which resulted in the faster dissolution rate in PBS. This is crucia l in drug delivery to enhance the efficiency of the desired pharmaceutical compound. The nanopore-injection (N-I) method can be used to fabricate more poorly water-soluble dr ugs and enhance they drug performance than those described here. Moreover, because it is intrinsically a low cost direct technology, N-I method can be used for scalable production when such fabrication become difficult or expensive with traditional nanofabrication methods. 48

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Figure 2-1. Experimental set-up for the hy drophobic drug nanoparticle preparation using nanoporous membrane. M1 Pressure meter. M2 Flow meter. Figure 2-2. Chemical structures of three hydrophobic compounds: silybin, beta-carotene, and butylated hydroxytoluene. 49

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Figure 2-3. Photograp h of a typical experimental setup. Figure 2-4. SEM images of nanoporous memb ranes: anodized aluminum oxide (AAO) membrane with (a) 20 nm inlet and (b) 200 nm outlet. 50

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Figure 2-5. Photograph of 40 mg silymarin nanoparticles obt ained within 20 min by using AAO nanoporous membrane. A penny serves as a size marker. Figure 2-6. SEM images of SM BC, and BHT drug nanoparticles. a, d, and g are SM, BC, and BHT NPs via N-I method, respective ly; b, e, and h are SM, BC, and BHT NPs via SEDS, respectively; c, f, and i are untreated SM, BC, and BHT, respectively. The scale bar is 500 nm in all the figures. 51

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Figure 2-7. Hydrodynamic di ameters of (a) SM, (b) BC and (c) BHT drug nanoparticles determined by DLS. Figure 2-8. Effect of flow rate on diameter of the SM NPs obtained. 52

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Figure 2-9. XRD pattern of silymarin nanoparticles and untreated silymarin powder. Figure 2-10. Dissolution profiles for s ilymarin nanoparticles and untreated silymarin powder in PBS (pH 7.4) at 37C. 53

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Table 2-1. Summary of the DLS analysis of hydrophobic nanoparticles. 54

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CHAPTER 3 GENERAL METHOD FOR PRODUCING POLYMERIC NANOPARTICLES USING NANOPOROUS MEMBRANES Aim The spatial and temporal control of the re lease of pharmaceutical s at the site of where they act is a key requirement for the therapeutic use of a drug. 150-152 One method for realizing this objective is to create drug-loaded nanoparti cles made out of biodegradable polymers. 141 Previous work in two laborator ies, one at Stanford University, the other at the University of Florida, has featured the generation of such nanoparticles. 142,143,153-158 We present here an alternative strategy based on the use of a nanoporous membrane that separ ates the two liquids. By pumping one liquid into the other, through the membrane, we can generate nanoparticles at or near the exits of the membrane nanopores. We illustrate this te chnique for the low molecular weight biopolymer chitosan, which is a polysacchari de consisting of 13% units of monomeric N-acetyl-glucosamine and 83% glucosamine units. 159 Low molecular weight chitosan (average MW 20,000 Daltons) is used as a model polymer in our work because it is a naturally biodegradable and biocompatible polysaccharide, which has broad applicatio ns in pharmaceutical and biomedical fields. 160-162 Chitosan is also known as a pH-response polymer, because at low pH, chitosans amines are prot onated and positively charged causing chitosan to be a water-soluble cationic polyelectrolyte. At high pH, these amines become deprotonated, and the polymer loses its charge and becomes insoluble. 163,164 Chitosan serves as a representative material for our process whic h can be adopted for other the production of other organic nanoparticles. 55

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Structure of Chitosan Experimental Materials Track-etched polycarbonate (PCT E, 10 nm in diameter, 6 m in thickness) was purchased from Osmonic Inc (Minnetonka, MN). Anodized aluminum oxide (AAO, 20 nm in diameter, 60 m in thickness) was purchased from Whatman Inc (Piscataway, NJ). Low molecular weight chitosan (CS, Mw 20, 000) was obtained from Sigma-Aldrich (St Louis, MO). Rhodamine 6G (R6G) was purchased from Fisher Scientific Inc (Worcester, MA). All other chemicals were reagent grade and used as received. Purified water, obtained by passing house-distilled water through a Barnstead E-pure water purification system, was used to prepare all solutions. Formation of Ultrafine Chitosan Nanoparticles The U-tube setup consists of two hal f U-tubes and a nanoporous membrane which is sandwiched between the two halves (see Figure 3-1). PCTE membrane with pore diameter of 10 nm, and AAO membrane with por e diameter of 20 nm are used in our experiments. The feed solution contained 25 mg of chitosan in 20 mL of 10 -3 M HCl (pH=3.0). The receiver solution was 10 mL of 10 -3 M NaOH (pH=11). One half of the U-tube was filled with 20 mL of feed solution, the other half was filled with 10 mL receiver solution. In some experiments 250 mbar gauge pressure was created by connecting a compressed air outlet with a pressure reducti on valve to the feed so lution side of the 56

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U-tube. In this way, the feed solution is pump ed into the receiver solution according to the applied pressure. Nanoparticles formed are collected by filtration through PCTE membranes and dried at room temperature. The filter membranes also serve as the substrate for scanning electron microscopy (SEM). Characterization of Chitosan Nanoparticles The size and morphology of formed nanoparticles are observed by scanning electron microscope (SEM) and transmission el ectron microscope (TEM). SEM images were acquired using an FEI XL30 Sirion SE M. Dry samples on carbon sticky tape were sputter-coated for 120 s at 15 mA with Pd/Au. Transmission electron microscopy (TEM) was carried out using a FEI Tecnai G2 F 20 X-TWIN. Samples were deposited on formvar carbon-coated copper grids. Hydrodynamic size and zeta potential of formed nanoparticles was measured by a Zetasizer Nano ZS (Malvern Instruments, Malvern, PA). Chitosan nanoparticles was dispersed in deionized water at a concentration of 100 g/mL, pH=7.0. Encapsulation of Rhodamine 6G in Chitosan Nanoparticles Rhodamine 6G encapsulated chitosan (CS/ R6G) nanoparticles were produced by N-I method. 5.0 wt% R6G is premixed with the feed solution. PCTE and AAO membrane were used to produce CS/R6G NPs. The mo rphology and size of formed nanoparticles were characterized by TEM and DLS. The amount of R6G encapsulated in the chitosan particle was determined by a fluorescent intensity method. Dry CS/R6G particles were dissolved in a phosphate/citrate buffer solution at pH=3.0. The fluorescence of dissolved solution was measured. The resulting fluorescence intensity was ca lculated into a R6G concentration in phosphate/citrate buffer solution using standar d concentration curve. The R6G loading 57

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efficiency of CS/R6G was calculated using the amount of R6G enca psulated divided by the amount of R6G premixed in the feed solution. Results and Discussion The experimental device (Figure 3-1) is composed of a nanoporous membrane, which separates two solutions. The pH of the f eed solution (left in Figure 3-1) is adjusted so that chitosan is soluble in this soluti on. The feed solution is forced under pressure through the pores of the membrane into the re ceiver solution (right in Figure 3-1). The pH of the receiver solution is adjusted such that chitosan is insoluble. When nanodroplets of the soluble chitosan are injected through t he membrane into the receiver solution nanoparticles of chitosan are formed. For the preparation of nanopar ticles with reduced sizes, membranes with uniform and well-defined nanopores are essential. 165-167 In our work, we use commercially available tra ck-etched polycarbonate (P CTE, OSMONIC Inc.) and anodized aluminum oxide (AAO, Whatm an Inc.) nanoporous membranes. The PCTE membrane is 6 m thick and contains track-etched cylindr ical pores with a diameter of 10 nm and pore density of 6 8 /cm 2 (Figure 3-2 a.). The AAO membrane contains is 60 m thick and contains 20 nm cylindrical pores at the face of the membr ane in contact with the feed solution. These pores run paralle l to one another for approximately 2 m and then feed much larger (200 nm in diameter) pores that run parallel to one another through the remaining thickness of the membrane. T he pore density of the AAO membrane at the entrance, (i.e., in contact wit h the feed solution) is around 6 14 /cm 2 (Figure 3-2 b, c). 144 The feed solution contained 25 mg of chitosan in 20 mL of 10 -3 M HCl (pH=3.0). The receiver solution was 10 mL of 10 -3 M NaOH (pH=11). The area of membrane exposed to these solutions, either PCTE or AAO, was 2 cm 2 Gravity flow was achieved via a height 58

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difference between the two solutions, causing t he low pH chitosan feed solution to flow into the high pH receiver solution. Nanodroplets are formed at or near the outlet of the PCTE nanoporous membrane in contact with t he high pH solution, causing precipitation of the chitosan. In the case of the AAO membrane the precip itation likely occurs at near the exits of the 20 nm nanopor es. The chitosan nanoparticles (CSNPs) are carried away from the membrane by the const ant gravity flow. No instances of clogging or sticking were found. Nanoparticles were collected from the rece iver solution by filtration, rinsed three times with deionized water, and dried in air at room temperatur e. We obtained 4.2 g of nanoparticles per hour by PCTE, and 610 g of particles per hour by AAO. These differing values are caused by the large pore density difference between the two kinds of nanoporous membranes. CSNPs were imaged using a TEM-1230 (JEOL) electron microscope, operated at 100 kV. Samples were deposited on carboncoated copper grids and negatively stained with 1% uranyl acetate. Figure 3-3 a shows a typical TEM image of the CSNP obtained using the PCTE membrane having 10 nm nanopores. The nanoparticles were found to have a mean diameter of 5 nm Figure 3-3 b shows that CSNPs obtained using the AAO membrane. These nanoparticles have a mean diameter of 21 nm, which suggests that they are formed near at the exit of the sm aller nanopores (20 nm) in the AAO membrane. Dynamic light scattering (DLS), measur ed with a Zetasizer Nano ZS (Malvern Instruments, Malvern, PA), was used to obt ain hydrodynamic particle diameters. The hydrodynamic diameters of the particles obtained using the PCTE and AAO membranes were 9 nm and 26 nm, respectively (Figure 3-4) The particle size from DLS is slightly 59

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larger than the diameter estimated using electron microscopy because DLS measures the diameter of the particles while still in solution, whereas TEM provides the diameter of the particles after thorough drying. 168 That larger particles are obtained using the AAO membranes reflects the fact that the pore diameter in contact with the receiver solution is 20 nm for this membrane vs. 10 nm for the PCTE membrane. We also investigated the effect of flow rate of chitosan solution on the particle-formation process. CSNPs obtained using the AAO membrane were used in these studies. The flow rate of ch itosan solution was varied from 7.2 L min -1 cm -2 to 32 L min -1 cm -2 by adjusting the height difference bet ween the feed and receiver solutions. DLS measurements were used to obtain the particle diameter s. Particle diameter was found to increase exponentially with flow rate, over the fl ow-rate range investigated (Figure 3-5). At higher flow rates nanowires are formed as found from SEM images (not shown). It was also found that the narrowest particle size distribution was obtained at a flow rate flow rate of 7.2 L min -1 cm -2 The viscosity of the chitosan feed solution also has a profound effect on nanoparticle-formation process. The viscosity of chitosan feed solution was varied by adding glycerol, while maintain ing its pH at 3. Particle sizes initially increased with viscosity but leveled at higher viscosities (Fi gure 3-6). We suggest that this is caused by a change in the diffusion rate, which decreases rapidly as the viscosity increases, causing larger particles to be formed at slower diffu sion rates. When the viscosity of chitosan solution achieves a certain point, particle size stops growing, perhaps owing to the gravity-induced detachment of the nanodroplets from the me mbrane into the sodium hydroxide solution. 60

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For the drug loading and encapsulation st udy, we use rhodamine 6G (R6G) as a model system to mimic a drug molecule. The organic molecule R6G is one of with most often used fluorescent dyes with excitation and emission wavelengths at 525 nm and 555 nm, respectively. 169-171 Using such a fluorescent model compound provides us with a rapid method to evaluate the enc apsulation data, which in turn allows us to optimize the process parameters. In our experiment, 5.0 wt% R6G is premixed with the ch itosan solution. Figure 3-7 shows the TEM images of R6G-loaded chit osan nanoparticles obtained using the PCTE and AAO membranes, respectively, and Figure 3-8 show the corresponding results obtained using dynamic light scattering. The amount of R6G encapsulated in t he chitosan particle was determined by dissolving the dry particles in a phosphate/ci trate buffer solution at pH=3 followed by fluorescence measurements of the released R6 G. When 5.0 wt% of R6 G, referred to the weight of chitosan, was added to the feed solution, and the PCTE membrane was used, the amount of R6G incorporated into the nanoparticles was 2.7 wt% (Table 3-1). The amount incorporated into the particles prepa red using the AAO membrane was 3.3 wt%. Table 2-1 summarized these results and includes the polydispersity index (PDI) values. Perspective In this chapter, we successfully prepar ed ultrafine chitosan nanoparticles by N-I strategy. The obtained nanoparticle has a small size (<30 nm) with narrow size distribution. Particle size increases with flow rate and viscosity of feed solution. Fluorescent dye rhodamine 6G was encapsul ated into chitosan nanoparticles with a encapsulation ratio of about 3 wt%. It provides a general technique for incorporating guest molecules in the host chitosan nanoparti cles. We believe that many other 61

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biodegradable polymer systems can be loaded with different organic compounds, which suggests the practical use of this techniq ue in preparing pharmaceuticals in nanoparticle form for drug delivery. 62

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Figure 3-1. Method for producing chitosan nanoparticles by flow though a nanoporous membrane. Figure 3-2. SEM images of nanoporous memb ranes: (a) track-etched polycarbonate (PCTE) membrane with 10 nm pores; and anodized aluminum oxide (AAO) membrane with (a) 20 nm inlet and (c) 200 nm outlet. 63

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Figure 3-3. Typical TEM images of chitosan nanoparticles (C SNPs) prepared by using (a) the PCTE membrane; and (b) the AAO membrane. In these TEM images, the black area represents t he nanoparticle, and the grey area represents the background. Figure 3-4. Comparison of size distributions of chitos an nanoparticles (CSNPs) prepared by using different nanoporous membranes determined by dynamic light scattering: (a) size of CSNPs obtained by PCTE membrane; and (b) size of CSNPs obtained by AAO membrane. 64

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Figure 3-5. Effect of soluti on flow rate on the diameter of the chitosan nanoparticle obtained. Figure 3-6. Effect of the viscosity of the chitosan feed solution on the diameter of the nanoparticles obtained. 65

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Figure 3-7. Typical TEM images of chit osan-rhodamine 6G nanoparticles prepared by using (a) the PCTE membrane and (b) the AAO membrane. In these TEM images, the black area represents the nanoparticle, and the grey area represents the background. Figure 3-8. Comparison of size distributi ons of chitosan-rhodamine 6G nanoparticles prepared by using different nanoporous membranes determined by dynamic light scattering: (a) PCTE me mbrane; and (b) AAO membrane. 66

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Table 3-1. Statistical size and encapsulat ion efficiency data for rhodamine 6G loaded chitosan nanoparticles. Membrane Particle Diameter TEM (nm) Diameter DLS (nm) PDI Encapsulation Ratio (%) PCTE CS NP 5 2 8 1 0.204 PCTE CS-R6G NP 5 3 9 2 0.108 2.7 AAO CS NP 21 5 26 2 0.228 AAO CS-R6G NP 30 8 30 4 0.333 3.3 67

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CHAPTER 4 BIODEGRADABLE POLYMERIC NANOPARTICLES AS DRUG DELIVERY VEHICLE Aim The applications of nanostructured biomaterials in drug delivery increase the need of developing specific and active drug delivery systems, which should distinctly advantage in improved bioavailability, enhanc ed drug performances, targeted delivery and sustained release. Biodegradable polyme ric nanoparticles have been extensively investigated for sustained release or targeted delivery of various therapeutics, including small molecular drugs and macromolecules. Tailo ring of the functional nanocarriers, such as the selection of matrix material and encapsu lation efficiency, has a profound influence on release profile, delivery e fficacy and drug performance. In this chapter, we use nanopore-injection (N-I) method for preparing two different biodegradable polymeric nanoparticl es: poly(lactide-co-glycolide)-co-poly(ethylene glycol) (PLGA-PEG) and chitosan nanoparticles. PLGA-PEG is a synthetic amphiphilic polymer composed of two biodegradable pol ymers widely applied in the drug delivery realm, while chitosan is a naturally occurring biopolymer used in many areas such as biomedical, pharmaceutical and biotechnological fields as well as in the food industry. Both PLGA-PEG and chitosan are Food and Drug Administration (FDA) approved safe biopolymer for drug delivery research. Ther apeutic agents were physically dispersed in the biodegradable polymer matrix (9-Amino6-chloro-5H-benzo(a) phenoxazine-5-one (MCHB) in PLGA-PEG NPs; Luciferin in chitosan NPs). MCHB is a toxic drug converted from a novel non-toxic prodrug 6-chloro-9-n itro-5-oxo-5H-benzo(a) phenoxazine (CNOB) using an specific enzyme E. coli nitroreductase ChrR6 (discovered by Professor A. C. Matin at Department of Microbiology and Imm unology in Stanford University). Utilization 68

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of prodrug in drug delivery, instead of direct ly delivery of toxic drug, could greatly minimize the damage to the normal tissue, thus enhance the effectiveness in treating cancer diseases. Hydrophilic compound luciferin is sele cted as another model therapeutic agent. Luciferin is a substrate for the enzyme lucifera se that produces a bioluminescent signal at approximately 610 nm at 37 C which can be measured in vivo. In our experiments, luciferin was encapsulated with chitosan to form chitosan/luciferin (CS/Luc) nanoparticles. Our drug encapsulated biodegr adable nanoparticles exhibited the capability of sustained release, which is favored in drug delivery. In vitro cytotoxicity of PLGA-PEG/MCHB nanoparticles tested by clonogen ic assay revealed an improved drug performance in eliminating PC-3 prostate ca ncer cells. CS/Luc nanoparticles expressed in vivo biocompatibility demonstrated by bi oluminescence imaging. We believe that biodegradable polymeric nanoparticles produced by N-I method have a promising future as novel drug delivery vehicle. 69

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Experimental Materials 9-Amino-6-chloro-5H-benzo(a) phenoxazine-5-one (MCHB) was provided by Prof. A. C. Matin (Department of Microbiology and Im munology, Stanford University). Luciferin was obtained from Polysciences Inc. (Warrington, PA) and used as received. Heterobifunctional PEG (amine-PEG-carboxylat e) (MW =3,400 g/mol) was obtained from Nektar Therapeutics (San Carlos, CA). Poly (D, L-lactide-co-gl ycolide) (PLGA) was obtained from Lactel Absorbable Polymers (Pelham, AL) with terminal carboxylate groups. All other chemicals we re purchased from Sigma-Aldr ich (St Louis, MO) and used without further purification. Synthesis and Characterization of PLGA-PEG Diblock Copolymer Synthesis of PLGA-PEG diblock copolymer is using the prot ocol reported by Gu and co-workers, 172 covalent bind PLGA-COOH with NH2-PEG-COOH using EDC/NHS chemistry: Reactant solutions were prepared as follo ws: a total of 10 g of PLGA-carboxylate (0.56 mmol) was weighed accurately, and disso lved in 20 mL dichloromethane (DCM); 270 mg N-hydroxysuccinimide (NHS, 2.2 mmol) and 460 mg 1-ethyl-3-(3-dimethylami nopropyl)-carbodiimide (EDC, 2.4 mmol) were weighed accurately, and dissolved in 4 mL DCM separat ely. EDC/NHS solution was slowly added into the PLGA-carboxylate solution. The re sulting PLGA-NHS wa s precipitated with 20 mL ethyl ether/methanol (50/50, v/v) blended solution. The PLGA-NHS was separated by 70

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centrifugation at 4,000 rpm for 20 min. The precipitation wa s washed and centrifuged for three times, and dried under vacuum over nigh t. A total of 2 g PLGA-NHS (0.118 mmol) resulting from the last st ep was weighed accurately and dissolved in 8 mL DCM, then added with 500 mg amine-PEG-carboxylate (0.148 mmol) and 56 mg N,N-Diisopropylethylamine (DI EA, 0.44 mmol), and stirred fo r 2 h. The resulting PLGA PEG diblock copolymer was precipitated with ether/methanol (50/50, v/v) blended solution and washed with the same solution for three times, to remove the unreacted PEG. PLGA-PEG diblock copolymer wa s dried under vacuum over night, and stored in -20 C for further experiment. NMR characterization of PLGA-PEG dibl ock copolymer: the PLGA-PEG diblock copolymer was characterized by a Mercur y 400 nuclear magnetic resonance (NMR, 400 MHz 1 H). For the preparation of sample solution: 5 mg PLGA-PEG was dissolved in 1 mL of deuterated chloroform (CDCl 3 ). Formation of Drug Encapsulated Nanoparticles PLGA-PEG/MCHB NPs Preparation: N-I method was used to prepared the nanoparticles. The detailed process was described as in Chapter 2. Briefly, in a typical U-tube setup, 50 mg PLGA-PEG and 2.5 mg MCHB were completely dissolved in 40 mL acetone as the feed solution, and PBS (pH 7.4) with 0.5 wt% Pluronic F68 was used as the receiver solution. 20 nm AAO membr ane was selected as the nanoporous membrane. 2 psi gauge pressure was applie d on the feed solution side. Obtained nanoparticles are collected by filtration and rinsed with deionized water for three times, and dried at room temperature. Chitosan/luciferin NPs Pr eparation: N-I method wa s used to prepared the nanoparticles. Feed solution contained 1.25 mg/m L chitosan and 62.5 g/mL luciferin in 71

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phosphate buffered saline (PBS, pH 3.0). Receiver solution contained PBS (pH 11.0) with 0.5 wt% Pluronic F68. All ot her parameters remained the same as described in the PLGA-PEG/MCHB NPs Preparation. Characterization of Drug Encapsulated Nanoparticles Scanning electron microscope (SEM) images were obtained using a FEI XL30 Sirion SEM. Dry samples on carbon sticky tape we re sputter-coated for 90 s at 15 mA with Pd/Au. Zetasizer Nano ZS (Malvern Instruments, Malvern, PA) was used to measure the hydrodynamic size, polydispersity index (PDI), and zeta potent ial of the obtained nanoparticles. In DLS measurement, nanoparticles were dispersed in the PBS (pH 7.4) with 0.5 wt% Pluronic F68 at a concentration of 100 g/mL. Sustained Release Study of Drug Encapsulated Nanoparticles A total of 2 mg obtained nanoparticles are accurately weighed and immersed in the 30 mL PBS (pH 7.4). The resulting solution was rotated at 100 rpm, and the temperature of the PBS (pH 7.4) was maintained at 37 0.5 C. 500 L of each sample was withdrawn and centrifuged at 10,000 rpm for 5 min in or der to remove the nanoparticles from the solution. The centrifuge supernatant solution wa s collect for the fluorescence absorbance measurement. The fluorescence absorbance intensity was performed using a Spectra Max Gemini EM Fluorescence Microplate Reader A series of samples were measured within total time duration up to 2 weeks. Fluorescent Microscopy Imaging For cell uptake studies, PC-3 prostate cancer cell line was used in this experiment. PC-3 cells were seeded on 35 mm glass-bottom ed culture dishes at a density of 8.5 4 cells/dish. On day 2 of culture, the ce lls were rinsed with PBS prior to addition of 72

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PLGA-PEG/MCHB NPs dropwise (200 L) with incubation at 4 C for 1 h. Unbound NPs were rinsed off with a gentle PBS wash, the incubation buffer was replaced, and the cells were warmed to 37 C for various times up to 4 h in the absence or presence of treatments. After incubation, the cells were rinsed four times with PBS and imaged with a Zeiss LSM 510 Meta NLO imaging system equipped with a Coherent Chameleon multiphoton laser mounted on a vibration-free table. In Vitro Cytotoxicity Study by Clonogenic Assay In the clonogenic assay, 1000 PC-3 prostate cancer cells were incubated with the following three samples: free MCHB (9.66 g), PLGA-PEG Null na noparticles (200 g), and PLGA-PEG/MCHB nanoparticles (200 g, MCHB: 9.66 g) for 1 h at 37 C, trypsinized and plated for clonogenic assay. The cultures of PC-3 cell lines were incubated at 37 C in a humid ified incubator with 7.5% CO 2 After two weeks, the number of colonies formed in each dish was c ounted with an Omnicon FAS II counter (Bausch & Lomb, Rochester, NY). Drug effects on the PC-3 cells are assessed as a reduction in PC-3 cell colony growth in the treated cultur es compared to the untr eated controls. In our experiment, control sample wa s the untreated PC-3 cells. In Vivo Cytotoxicity Stud y by Bioluminescence Imaging We use transgenic mice that express fire fly luciferase (FVB-luc+) to evaluate delivery of luciferin from the chitosan nanop articles. Four mice were intravenously injected with CS/Luc NPs (two mice) and chit osan null NPs (two mice); two mice were subcutaneously injected with CS/Luc NPs (right area in the mice) and chitosan null NPs (left area in the mice). For all injected nanoparticle solutions, they contain 2.0 mg nanoparticles dispersed in 250 mL of 0.5 wt % Pluronic F68 in PBS. Bioluminescence imaging was performed on a Xenogen IVIS 200 using a cooled charge-coupled device 73

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camera. Data was analyzed with LivingImage software (Xenogen) and expressed in photons per steradian per second for each region of interest such that the data are not dependent on camera settings, chamber geometry, or integration time. Results and Discussion PLGA-PEG/MCHB Nanoparticle In order to synthesize PLGA-PEG copolymer, we covalent bind PLGA-COOH with NH2-PEG-COOH using EDC/NHS chemistry: The terminal carboxyl group on the hydropho bic PLGA reacted with the terminal amine group on the hydrophilic PEG, and form an amphiphilic diblock polymer. The 1H NMR characterization of PLGA-PEG is shown in the Figure 4-1. The presence of PEG on the PLGA-PEG was visualized by the c haracterized peaks between 3.4 and 3.8 ppm, which is in accordance with literature. 172 It confirmed that PEG was covalently bonded with PLGA and formed diblock copolymer. PLGA-PEG/MCHB NPs were formulated by N-I method, and a typical SEM image is shown in Figure 4-2. From this image, it could be found that PLGA-PEG/MCHB NPs exhibit a smooth spherical morphology with a relatively narrow size distribution. The mean diameter of these NPs is around 90 nm calculated by the size measurement using ImageJ. Dynamic light scattering (DLS) measuremen t was performed on a Zetasizer (Nano ZS) to analyze the hydrodynamic diameter and the zeta potential. As shown in Figure 4-3, PLGA-PEG/MCHB NPs show a hydrodynamic diameter of 96 7 nm with a low 74

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polydispersity index (PDI) of 0.143. It revealed that PLGA -PEG/MCHB NPs formed by N-I method have a uniform size with in 100 nm, and could be well dispersed in the aqueous environment without forming aggregations. The zeta potential of these NPs is .5 mV in pH=7.4 PBS. The negative charge is from the carboxyl groups at the terminal end of the hydrophilic PEG block. The outer sphere composed of PEG is believed to help nanoparticle escape from reticuloendothelial syst em elimination, resulted in enhanced biocompatibility and circulation ha lf-life in vivo. It also minimi zes electrostatic interactions with opposite charged biomolecules in the in vivo environment, preventing from aggregation. Meanwhile, the functi onal PEG chains will facilitate surface modification of the nanoparticles, such as antibody conjugation for targeted delivery. MCHB as a fluorescent model compound (e x: 575 nm, em: 625 nm) allows us to evaluate the encapsulation ratio with a rapid method, i.e. fluorescence spectrum. In our experiment, the amount of MCHB encapsulated in PLGAPEG particle was determined by dissolving the cleaned dry nanoparticles in acetone followed by fluorescence measurements. When 5.0 wt% of MCHB, refe rred to the weight of PLGA-PEG, was added to the feed solution, the amount of M CHB incorporated into the nanoparticles was 4.8 wt%. The encapsulation efficiency (EE) of MCHB is calculated to be 97%, using the formula EE = Amount of drug bound/Total amount of drug used for nanoparticle production. The in vitro sustained release of PL GA-PEG/MCHB NP was studied under the intimating degradation condition in human body, dispersing them in PBS (pH 7.4) at 37 C and rotated at a low speed. The release prof ile of PLGA-PEG/MCHB NPs is shown in Figure 4-4. In this figure, the profile demons trates a rapid increase of cumulative drug 75

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release in the first 12 h, ov er 50 wt% MCHB was released. It may suggest that the MCHB encapsulated at or near the surface of the particle released much faster than those encapsulated within the center. After that, th e release speed of MCHB slowed down, and the nanoparticles gradually released the rest MCHB over a total time of 7 days (approached 95%). Such a controlled release prof ile facilitates the NPs for the delivery of anticancer therapeutics. The efficacy of the MCHB encapsulated NPs to defeat cancer cells is reflected by their cytotoxicity for the cancer cells. Then, we tested the in vitro cellu lar cytotoxicity of the obtained PLGA-PEG/MCHB NPs by clonogenic assa y, which is a microbiology technique for studying the effectiveness of specific agents on the survival and proliferation of cells. The in vitro differential cytotoxicity of MCHB free drug, PLGA -PEG null NPs, and PLGA-PEG/MCHB NPs, as well as the blank c ontrol, were tested using PC-3 prostate cancer cell line. The total number of colonies in blank control sample was counted and serviced as a background. Figure 4-5 shows t he quantitative analysis re sults. In the free drug samples, around 50 % of PC-3 cells were eliminated. No killing effect was observed in the PLGA-PEG null NPs samples, which s uggests that PLGA-PEG nanoparticle, as a drug delivery vector, is non-toxic to PC-3 cells. When PLGA-PEG/MCHB NPs were cultured with PC-3 cells, all the PC-3 cells were eliminated after 1 h incubation. It suggests that PLGA-PEG/MCHB NPs are signi ficantly more cytotoxic than free MCHB. The observed toxicity in the PLGA-PEG/MCHB may due to the cell uptake and subsequent MCHB release. These cell viabilit y data confirmed that using PLGA-PEG NP as a vector improved the MCHB deliver y efficiency, and enhanced MCHB in vitro performance. 76

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The fluorescent image of PC-3 cells a fter cultured with PLGA-PEG/MCHB NPs was shown in Figure 4-6 a. Fluorescence signals were from the internalized PLGA-PEG/MCHB NPs by PC-3 cells. The image is blurred because PC-3 cells were vibrating under the microscope due to the cytotoxicity of MCHB. A much clearer fluorescent image was observed by incubating PC-3 cell with non-toxic PLGA-PEG/CONB NPs, which is shown in Figure 4-6 b. Chitosan/luciferin Nanoparticles (CS/Luc NPs) As reported in the Chapter 3, we have successfully prepared ultrafine chitosan nanoparticles (< 30 nm) by N-I method. In this chapter, we prepared chitosan nanoparticles (130 nm in diamete r) for drug delivery through the similar procedure. Figure 4-7 shows a typical SEM image of the CS/Luc NPs. The nanoparticles were found to have a mean diameter of about 130 nm. Dynamic li ght scattering (DLS) was used to determine the hydrodynamic diameter of the CS/Luc NPs (Figure 4-8) The hydrodynamic diameter of the particles obtained using the AAO me mbranes was 137 13 nm with a PDI of 0.274. The zeta-potential of CS/Luc NP is 7.0 mV in pH=7.4 PBS. Th is positive surface charge of obtained nanoparticle attributes to free amine groups on chitosan polymer chain. Those free amine groups acquire prot ons from aqueous solution at near neutral pH and lead to a slightly positive charge existing on the surface of the nanoparticles. The drug loading and encapsulation of CS/Luc NP was examined following the same method in PLGA-PEG/NPs. Luciferin is natural fluorescent compound with the excitation wavelength at 330 nm and the emi ssion wavelength at 525 nm. When 5.0 wt% of luciferin, referred to the weight of chitosan, was added to the feed solution, and the AAO membrane was used, the amount of luci ferin incorporated into the nanoparticles was 1.8 wt%. The encapsulation efficiency (EE) of luciferin is calculated to be 36%. EE of 77

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luciferin in CS NPs (36%) is much lower than that of MCHB in PLGA-PEG NPs (97%) because both NPs were eventually precipitat ed in the aqueous phase, and luciferin (salt form) is very hydrophilic and difficult to be trapped in the polymer matrix compared with hydrophobic MCHB. The in vitro sustained release of CS/Luc NPs was shown in Figure 4-9. CS/Luc NPs was dispersed in PBS (pH 7.4) at 37 C and rotated at a low speed. The release half life of CS/Luc NPs, which means 50 wt% of luciferi n encapsulated in the CS NPs is released into the solution, is about 3 days. The total release time of CS/Luc NPs is about 12 days. An early rapid release of luciferin in the release profile was observed, which is similar as the one in PLGA-PEG/MCHB NP release profile. We also studied the in vivo toxicity of obtained CS/Luc NPs. Xenograft model was used by intravenous and subcutaneous injecti on of CS/Luc NPs within luciferase transgened mice. All mice used in our ex periment were transgene engineered mice expressing luciferases, which could conver t existing luciferin in mouse body into bioluminescence signals. The results are shown in Figure 4-10. In Figures 4-10 a and b, two pairs of mice were intravenously injected with CS/Luc NPs and CS Null NPs. The first pair of mice showed whole body fluoresc ent signals, which suggested CS/Luc NPs distributed over the mice body with blood circul ation. No biolumines cence signals were observed in the second pair of mice which were injected with CS Null NPs, confirming that the bioluminescence signals obs erved in the mice were from the luciferin released from CS/Luc NPs. In Figure 4-10 c, a pair of mi ce was subcutaneously in jected with both CS Null NPs (Left side) and CS/Luc NPs (Right side) at areas of inte rest. Bioluminescence signals were observed at the injection site of CS/Luc NPs due to the release of luciferin, 78

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whereas non fluorescence in the left area. After injections, all three pairs of mice used in our experiment survived and behaved normal ac tivities. No biotoxic effects were observed with those mice, which suggested t hat our CS/Luc NPs are biocompatible and safe for further in vivo drug delivery study. Perspective We have successfully prepared nanoparticles composed of synthetic or naturally occurring materials by N-I method. Hydrophobic/hydrophilic drugs were encapsulated in the polymeric nanoparticles in high efficiency. These nanoparticles exhibit the ability of sustained release of encapsulated drugs: 7 da ys for PLGA-PEG/MCHB NPs; 11 days for CS/Luc NPs. In vitro cytoto xicity experiment proved that PLGA-PEG nanoparticle vector could enhance MCHB delivery efficiency. CS /Luc nanoparticles could be successfully detected in the in vivo bioluminescence imaging. Many multif unctional drug delivery nanoparticles could be produced by N-I method in the future, combining sustained release with pathway tracking. 79

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Figure 4-1. NMR characterization of (a) PLGA, (b) PLGA-PEG diblock copolymer. Figure 4-2. Typical SEM image of PLGA-PEG/MCHB nanoparticles. 80

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Figure 4-3. Hydrodynamic di ameter of PLGA-PEG/MCHB NPs determined by DLS. Figure 4-4. In vitro sustained releas e profile of PLGA-PEG/MCHB NPs. 81

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Figure 4-5. In vitro cytotoxicity study of PLGA-PEG/MCHB NPs. Figure 4-6. Fluorescent im age of PC-3 cell incubated wit h (a) PLGA-PEG/MCHB NPs, and (b) PLGA-PEG/CNOB NPs. 82

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Figure 4-7. Typical SEM image of CS/Luc nanoparticles. Figure 4-8. Hydrodynamic diameter of CS/Luc NPs determined by DLS. 83

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Figure 4-9. In vitro sustained re lease profile of CS/Luc NPs. Figure 4-10. In vivo biotoxicity study of CS/Luc NPs by bioluminescence imaging. 84

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Table 4-1. DLS data for PLGA-PEG/MCHB and CS/Luc nanoparticles. 85

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CHAPTER 5 FORMATION OF BIODEGRADABLE NANO FIBERS BY NANOPOROUS MEMBRANE Aim In human and animals, majority cells are imbedded in the natural extracellular matrix (ECM), which is a network of nanof ibers containing collagens, glycosaminoglycans, proteoglycans, and glycoproteinsglycosaminogl ycans. The interaction between these cells and ECM regulates the cell migration, prolifer ation, differentiati on, gene expression, and secretion of various hormones and growth factors. 43 So mimicking natural ECM is of great importance in t he tissue engineering. Collagen is widely considered as the mo st promising biopolymer in tissue engineering because it is the primary component in the natural ECM. Collagen is a group of structure proteins feat uring a unique triple-helix wh ich is composed of three polypeptide subunits. There are more t han 20 types of collagen in human body. The differences between the isotypes of collagen are the nature of the non-helical parts and the length of their triple-helix. Type I collagen is the most abundant and predominant collagen, which is a triple-hel ical protein with 300 nm in l ength and 1.5 nm in diameter. Under certain circumstance, such as pH, tem perature, or ionic stre ngth variance, type I collagen molecules will self assembly into macr omolecular structure (fibrils and bundle of fibrils). Collagen nanofibers are widely used as novel building block in engineered tissue scaffold. 46-48 In this chapter, we successfully prepared collagen nanofibers by a modified nanopore-injection (N-I) method. The obtained collagen nanofiber s exhibit a uniform size distribution with a mean diam eter of about 100 nm. Fiber diameter increases with nanopore size of PCTE membrane. Collage n nanofibrous scaffolds were prepared by 86

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filtration of obtained nanofibers, and were used for cardiac stem cells (CSCs) growth and proliferation study. The nanof ibrous scaffold prepared by N-I method may provide a solution to the challenge of three dimensiona l artificial scaffold in tissue engineering. Experimental Materials Type I tropo-collagen extracted via acid -solubilization of rat tail tendon was purchased from BD Biosciences (Bredford, MA). Stock solutions were 3.44 mg/ml in 0.1 M acetic acid. All chemicals were purchased from Sigma Aldrich (St. Louis, MO) and were used without further purification. Formation of Collagen Nanofibers The U-tube setup consists of two hal f U-tubes and a nanoporous membrane which is sandwiched between the two halves (see Figure 5-1). Tra ck etched polycarbonate (PCTE) nanoporous membranes with pore diameters between 50 nm and 1 m were used in our experiments. One half of the U-tube was filled with 6 mL of feed solution containing 1 mg/mL collagen and 1 mM HCl (pH 3); the other half was filled with 4 mL receiver solution containing 1 mM NaOH (pH 11). A 4 psi gauge pressure was created by connecting a compressed air outlet with a pressu re reduction valve to the feed solution side of the U-tube. In this wa y, the feed solution is pumped into the receiver solution at a constant flowrate according to the applied pressure. Nanofibers formed are collected by filtration through PCTE membranes and dried at room temperatur e. PCTE filter membranes also serve as the substrate fo r scanning electron micr oscopy (SEM). In control experiments, different kinds of membranes (PCTE wit h different pore sizes) were used in order to investigate the influence of filtration upon arti fact formation. The nanofiber always had the same appearance. 87

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Characterization of Collagen Nanofibers Scanning Electron Microscope (SEM): SEM images were acquired using an FEI XL30 Sirion SEM. Dry samples on carbon sti cky tape were sputter-coated for 120 s at 15 mA with Pd/Au. The diameters of the fibers were evaluated with the software ImageJ. Transmission Electron Microscopy (TEM): TEM analysis was carried out using a FEI Tecnai G2 F20 X-TWIN. For TEM, samples were deposited on formvar carbon-coated copper grids, without prior filtration. Rheology analysis: The rheol ogical experiments were carried out using a TA AR-G2, equipped with an 8 mm parallel plate geometry. While the plate geometry was oscillated at a frequency, we measured the torque (stress) that was required to arrive at a certain deformation (strain). Frequency sweep tests have been carried out with a strain of = 1 %, strain sweep experiments were held at a constant frequency of = 1 rad/s. The nanofibrous scaffold used in rheology study was prepared by filtration of collagen nanofibers on a PCTE membrane, ri nsed with deionized water for three times and dried at room temperature. Isolation and Culture of Card iac Stem Cells (CSCs) Animal protocols were approved by the St anford University Animal Care and Use Committee. The L2G85 transgenic mice of F VB background with beta-actin promoter driving Fluc-eGFP was used as stem cell don or. CSCs were isolated from 6to 12-week-old L2G85 mice, The myocardial tiss ue was cut into a 1 to 2mm pieces, washed with Hanks' balanced salt solution (Invitrogen, Carlsbad, California), and incubated with 0.1% collagenase II for 30 min at 37 C with frequent shaking. Cells were then filtered through 100 m mesh. The cells obtained were cultured in Iscove's modified Dulbecco's medium supplemented with 10% fetal bovine serum (FBS) (Hyclone, Logan, Utah), 0.1 88

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mmol/L nonessential amino acid s, 100 U/mL penicillin G, 100 g/mL streptomycin, 2 mmol/L glutamine, and 0.1 mmol/L beta-mercapt oethanol. 2 to 3 weeks post seeding, a population of phase-bright ce lls appeared over the adhered fi broblast-like cells. These phase-bright cells were collected by 2 wa shes with phosphate buffered saline (PBS), and 1 wash with cell dissolution buffer (Gibco, Grand Island, New York) at room temperature under microscopic monitoring, and sub-cu ltured in poly-lysine-coated plates (BD Pharmingen, San Jose, California) with the same medium (Online Videos 1 and 2). In vitro cell growth and differentiation: an a liquot of CSCs was maintained in culture to monitor cell growth and stability of Luciferas e (Luc) expression during the course of the experiment. To determine Luc ex pression, every 15 days cells were trypsinized, counted and an aliquot containing 2 5 cells was re-suspended in r eporter lysis buffer (RLB) 1X (Promega) and frozen at 80 C. At the end of the experim ent the collected samples were used to determine Luc activity and DNA concentration Scaffold Seeding Small numbers (up to 200,000) of CSCs ce lls were used to seed the nanofibrous scaffolds. For seeding, each scaffold was immersed in an aliquot (50 L) of cells in DMEM (containing 10% FBS, Pen/St rep antibiotics. The construc ts were maintained under standard cell culture conditions fo r the rest of the experiment. At that time, 1mL culture medium was added per construct. After 24 h of incubation, the scaffolds were noninvasively imaged, washed three times with 1mL of PBS (to elimi nate the luciferin), and stored at 20 C until the in vitro luciferase and fluorescence assays were performed. For comparison purposes, a series of the same number of CSCs cells in suspension in 50 L PBS were prepared in parallel and treated in the same way as scaffolds seeded with cells. 89

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Microscopy Fluorescent Imaging CSCs cell seeding and attachment was visualized by phase contrast and fluorescent (excitation f ilter 470 20 nm, emission f ilter 525 25 nm, Chroma Technology Corp., Rockingham, VT) microscopy equipment (CK40 microscope, Olympus, Melville, NY) with digital camera (S pot RT color, Diagnostic Instruments Inc., Sterling Heights, MI). For fl uorescent imaging, a 530 25 nm wavelength filter (Chroma Technology Corp.) was attached to the CCD camera. Samples were placed in the imaging cabinet and high resolution (1 pixe l binning) images were acquired at a camera exposure time of 1 s. Fluoresc ent signal was expressed in cts/pix/s. Bioluminescence Imaging (BLI) BLI of cultured CSCs was performed to charac terize the cell proliferation condition, when reaching cell confluence. For BLI, medium was removed from the plates, cells were rinsed twice with PBS 1 and 300 L or 1 mL of luciferin reagent stock (Promega) was added to each well, and imaged immediately. For imaging, a plate was placed in the detection (BLI) was carried out using an IV IS 100 imaging system (Xenogen, Alameda, CA. For in vitro imaging, CSCs cell seeded on 6 well plates or seeded with the collagen nanofiber scaffolds were loaded in individual wells of 24-well plates, immersed with 200 L of D-luciferin solution (300 g/mL in PBS), and imaged after 5 min using the medium binning setting of the instrument for 1min. To moni tor cell proliferation on the nanofiber scaffolds in vitro, the cells were imaged on the day of seeding (day 0) as well as days 3, 7, and14 post seeding. Light emission was quantified using Living Image software (version 2.50.2; Xenogen). Each frame depicted the biolumine scent signal in pseudocolor superimposed on the respective gray-scale photographic image. Data from a region of interest surrounding each well were manually selected on the frames and were reported 90

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as the total photon flux defined by the net phot on count emitted per se cond per centimeter squared per steradian (p/s/cm 2 /sr). Statistical Analysis Each in vitro experiment was performed twice in triplicate. Numerical results were reported as meanstandard error of the mean. Results and Discussion In this chapter, we successfully prepared collagen nanofibers based on the modified nanopore-injection (N-I) setup (as shown in Fi gure 5-1). Modified N-I method features two major differences in comparison with nanoparticle formation process. First, magnetic stirring is removed from the system because wall shear force created by magnetic stirring may cause obtained nanofibers break into fragments or particles. Second, a higher pressure flow is applied in the N-I experiment (f rom 2 psi to 4 psi). The resulting high flux of feed solution tends to form fibers instead of particles at the exit of the nanopore. The morphology of obtained nanofibers was in vestigated by SEM and TEM. Figures 5-2 a, b show the typical SEM images of collagen nanofibers at low and high magnifications. Collagen nanofibers are in relatively unifo rm size with smooth surface. The mean diameter of nanofiber s measured from the image is 102 13 nm. The length of nanofibers is in the range of few micrometers to hundred microm eters. From Figure 5-2 b, we could see that some collagen nanofibers fu rther self assembly into bundles. Figure 5-3 shows typical TEM images of obtained nanofibers. The aver age diameter of nanofibers analyzed from TEM im age is about 90 nm, which is similar to the result obtained from SEM image. The inse t of Figure 5-3 b shows the electron diffraction pattern on a single collagen nanofiber and it reveals that the collagen nanofibers are in 91

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amorphous phase. A nanofibrous scaffold (~ 3 mg) could be obtained within 2 h by filtration (as shown in Figure 5-4). The influence of nanoporous membrane on nanofiber formation was also investigated. A comparative experiment was carried out under other fixed conditions without nanoporous membrane. Re sulting products were collected by filtration and characterized by SEM. Compared with the sa mple produced with N-I method (Figure 5-5 a), no nanofibers were formed in the comparative experiment as shown in Figure 5-5 b. Without nanoporous membrane, feed solution is di rectly mixed with the receiver solution, and resulted product was a smooth coll agen film. It suggested that nanoporous membrane played a crucial role in collage n nanofiber formation. Because the self assembly of collagen molecule is believed happening at the exit of the nanopore. The uniform diameter and spacing between individual nanopores help forming well dispersed nanofibers with a narrow diam eter distribution. Collagen nanofiber formation with four diffe rent pore size (50 nm, 200 nm, 400 nm and 1000 nm) PCTEM membrane was investigated (Figure 5-6). In order to reduce the difference in flow speed caused by the differenc e in flow area, a pressure of 4 psi over atmospheric pressure was applie d at the side of the U-tube containing the feed solution. Nanofibers grown in 50 nm pores had a diameter of (50 15) nm, (120 50) nm for 200 nm pores, (270 120) nm for 400 nm pores, and (760 240) nm for 1000 nm pores. Smaller pores prevented the collagen from passing the membrane due to congestion of the pores. We also studied the biomechanical char acteristics of the collagen nanofibrous scaffold measured by rheology measur ement. Small-deformation oscillatory 92

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measurements were performed to evaluate the viscoelastic behavior of the nanofibrous scaffold. Figure 5-7 shows the mechanical spec tra of the collagen nanofibrous scaffold that was prepared by filtration of collagen nanofibers. In th is rheology measurement, the storage modulus (G) and the loss modulus (G) were monitored as a function of different frequency. The value of G represents energy stored elastically in the system, whereas the value of G represents energy dissipated through viscous effects. As shown in Figure 5-7, the value of the storage modulus G exceeds that of the loss modulus G by almost one order of magnitude, indicating that obt ained scaffold is a strong and rigid gel. Collagen nanofibrous scaffold was tested for the ability of supporting cell growth and proliferation using cardiac st em cells (CSCs). CSCs are normally found in heart tissue, capable of differentiating into cardiac cells associated with contracting myocardium and vascular structures. The aim of using nanofib rous scaffold to support CSC growth is eventually replacement of dead tissue in the diseased human heart with new generated tissues seeded in our biodegradable scaffold. In our experiment, collage nanofibrous scaffold and blank control were incubated with CSCs. After 1 week incubation, CSC proliferation was evaluated by fluorescent imaging and bioluminescence imaging, as shown in Figure 5-8. Figure 58 a shows fluorescent image of CSCs growth with collagen scaffold, compared with the blank control sa mple. These fluorescent cell images showed that CSCs on collagen scaffold could grow a nd proliferate normally as in the control sample. In the high magnification image, it is clearly shown some stretched CSCs, which suggests that scaffold interacts with cells re sulting in cell flattening and spreading on the scaffold surface. 93

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Bioluminescence imaging assay was also us ed to evaluate the activity of CSCs on collagen scaffold. Basically, luciferin was add ed to the CSCs on scaffold, and expressed as bioluminescence signals through luciferase Bioluminescence intensity was measured and represented the activity of luciferase in CSCs indicating the viability of CSCs. From Figure 5-8 b, it could be observed that bioluminescence intensity of CSCs on scaffold is similar to that of control sample, which conf irmed that CSCs proliferated in a similar way as cell grew in blank sample. It could be drawn that collagen nanofibrous scaffold shows non-biotoxicity, and could be used to support cardiac stem cell proliferation. Perspective In conclusion, we present a novel and e fficient method for the fabrication of biodegradable type I collagen nanofibers. Uniform colla gen nanofibers were produced with PCTE membrane. Furthermore, diameter of collagen nanofiber could be controlled by the pore size of nanoporous membrane. Us ing the membrane with pore size ranging from 50-1000 nm, nanofibers with diameter ranging from 50760 nm could be obtained. The nanofibrous scaffold exhibited distinct biomechanical strength. In vitro experiment shows the nanofibrous scaffold supports cardiac st em cell growth and pr oliferation, as an early indication of scaffold biocompatibility. We believe that many other biopolymers could be formulated into nanofibers by N-I method for tissue engineering purposes. 94

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Figure 5-1. Method for produc ing collagen nanofibers by fl owing though a nanoporous membrane. Figure 5-2. Typical SEM images of coll agen nanofibers prepared by using the PCTE membrane at (a) high magnificati on and (b) low magnification. 95

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Figure 5-3. Typical TEM images of (a) a bundle of collagen nanofibers (b) a single collagen nanofiber. Inset is the related selected area el ectron diffraction pattern (SAED pattern). Figure 5-4. Photograph of co llagen nanofibrous scaffold prepar ed by N-I method in 2 h. A penny is used as a size marker. 96

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Figure 5-5. SEM images of (a ) collagen nanofibers prepared by N-I method (b) collagen film prepared without nanoporous membrane. Figure 5-6. Effect of nanopore size on t he diameter of the collagen nanofibers. 97

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Figure 5-7. Rheology study of collagen nanofibrous scaffold. Figure 5-8. (a) Fluorescent mi croscope imaging of CSCs in (A) and (C) blank control (low and high magnification); (B) and (D) co llagen nanofibrous scaffold (low and high magnification). In fluorescent micr oscope images, bright area represents CSCs, and black area represents backgroun d. (b) bioluminescence image of CSCs proliferation on blank contro l and collagen nanofibrous scaffold. 98

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CHAPTER 6 ORGANIC/INORGANIC HYBRID NANO FIBERS FOR TISSUE ENGINEERING Aim Organisms bring forth a wide va riety of organic/inorganic hybrid materials, called biominerals. The most common biomineral s are the phosphate and carbonate salts of calcium that are used in conj unction with organic polymers such as collagen and chitin to give structural support to bones and shells. Bi omineralization has inspired chemists to seek new synthetic strategies for creating inorganic materials with complex forms e.g. by pattern recognition of self -organized organic assemblies. 173,174 Next to the advancement of our understanding of biological processes, the main goal of these studies is to find new materials for bone grafting, tissue engineerin g, or other medical applications. One of the most promi nent features of biominerals is that they seldom exhibit typical crystalline morphologies, including sharp edges and angled corners. In fact, in many cases biominerals seem to be molded into a specific shape, just like polymeric plastics that are extruded in industrial processes. 175,176 Several non-classical crystallization pathways have been proposed for biomineralization. 177-180 In the last several years, evidence for the importance of an amorphous precursor phase rapidly accumulated and quickly became the dominant view in the field. 176,181,182 In 2003 Gower and co-workers 175,181 proposed the polymer induced liquid precursor (PILP) process for the crystallization of calcium carbonate (CaCO 3 ). Using acidic hydrophilic polymers as additives, they were able to observe a transient phase of liquid calcium carbonate droplets. These droplets can be extruded into constrained vo lumes where they coalesce and transform into a more stable mineral phase. 183,184 As has been established by several 99

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studies, 185-187 the role of the acidic polymer is twofold: it suppre sses bulk crystallization of CaCO 3 and stabilizes the amorphous phase. In this chapter, we present a new approach for preparing nanofibers that was inspired by these biomineraliz ation processes. Our strategy is based on the use of a nanoporous track etched polycarbonate (PCTE) membrane that separates two liquids, a feed solution and a receiver solution. N anofibers are formed by pumping the feeder solution through the membrane into the receiver solution. In prior chapt ers, this strategy has been successfully used for the fabricati on of biodegradable nanoparticles. The feed solution contains an inorganic cation, such as Ca 2+ and a long-chain polymer, such as collagen, which serves as the scaffolding for the nanofiber. The receiver solution contains an anion, such as phosphate or carbonate, which induces precipitation of the respective inorganic salt along the long-chain polymer. Using this method, for the first time, it wa s possible to incorporate calcium phosphate into collagen nanofi bers without any additional polymers or proteins. 188-190 This was possible due to the simult aneous formation of collagen nanofibers and amorphous calcium phosphate at the exit of the pores of the PCTE membrane. Therefore, we are able to present a bottom-up approach fo r the artificial fo rmation of the basic building blocks of bones. In further experiments, our method was also tested with a synthetic polymer/biomineral composite system. Th e feed solution was an aqueous solution of Ca 2+ and poly(acrylic acid) (PAA), while the rece iver solution was an aqueous solution of carbonate (CO 3 2). By pumping the feed solution th rough the nanoporous membrane into the receiver solution, we were able to generate nanofibers that consist of amorphous 100

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calcium carbonate and PAA. Similar experiment s featuring phosphate anions instead of carbonate, gave rise to nanofibers that cons ist of a composite of amorphous calcium phosphate and PAA. In order to demonstrate t he clinical usefulness of the fibers generated with this approach in a tissue-engineering context, human adipose derived stem cells (hADSCs) have been grown on the collagen scaffold c onsisting of aggregates of nanofibers. Experimental Materials All chemicals were purchased from Sigm a Aldrich (St. Louis, MO) and were used without further purification. Calcium chloride (CaCl 2 ), sodium bicarbonate (Na 2 CO 3 ) solutions and sodium biphosphate (Na 2 HPO 4 ) were prepared fresh on a daily basis using Millipore water. Poly(acrylic acid) (MW 8,000 g/mol, 45% in water) was added to the CaCl 2 prior to the experiments. T he PAA concentration in the CaCl 2 solution was 150 mg/mL unless otherwise noted. The samples were vortexed for 2 min in order to insure proper dissolving of the polymer Type I tropo-collagen from ra t tails was purchased from BD Biosciences (Bredford, MA). Stock solu tions were 3 mg/mL tropo-collagen in 10 mM HCl. 10X PBS buffer was obtained from Invitrogen (Carlsbad, CA). Formation of Hybrid Nanofibers The U-tube setup consists of two hal f U-tubes and a nanoporous membrane which is sandwiched between the two halves (see Figure 6-1). Tra ck etched polycarbonate (PCTE) nanoporous membrane with pore diameter of 200 nm was used in our experiments. One half of the U-tube was filled with 6 mL feed solution containing 1 mg/mL collagen, 1 mM CaCl 2 and 1mM HCl (pH 3), the other half was filled with 4 mL receiver solution containing 0.66 mM Na 2 HPO 4 and 1 mM NaOH (pH 11). In some experiments 4 101

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psi gauge pressure was created by connecting a compressed air outlet with a pressure reduction valve to the feed solution side of th e U-tube. In this way, the feed solution is pumped into the receiver solution according to the applied pressure. Nanofibers formed are collected by filtration through PCTE membranes and dried at room temperature. PCTE filter membranes also serve as the substrate for scanning electron microscopy (SEM). In control experiments, different kinds of membranes (AAO, PCTE with different pore sizes) were used in order to investigat e the influence of filtration upon artifact formation. The nanofiber a lways had the same appearance. PAA/CaCO 3 and PAA/Calcium Phosphate nanofibers were produced in the similar way as described above. For PAA/CaCO 3 formation, feed solution contained 20 mM CaCl 2 and 150 mg/ml PAA; receiver solution contained 20 mM Na 2 CO 3 For PAA/Calcium Phosphate formation, feed so lution contained 20 mM CaCl 2 and 150 mg/mL PAA; receiver solution contained 15 mM Na 2 HPO 4 Other conditions rema in the same as in collagen/Calcium Phosphate experiments. Analysis of Nanofibers by Electron Microscope SEM images were acquired using an FEI XL30 Sirion SEM. Dry samples on carbon sticky tape were sputter-coated for 120 s at 15 mA with Pd/Au. The diam eters of the fibers were evaluated with the software ImageJ. Transmission electron microscopy (TEM) was carried out using a FEI Tecnai G2 F20 X-TWIN. For TEM, samples were deposit ed on formvar carbon-coated copper grids, without prior filtration. Stem Cell Preparation Human adipose derived stem cells (hADSCs) were isolated from donors and expanded in culture. Cells we re cultured in Dulbecos Modified Eagles Medium (DMEM) 102

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supplemented with 10% fetal bovine serum, 1% penicillin/str eptomycin and 0.05% fibroblast growth factor. Two-dimensional sheets of nanofibers pr epared on PCTE filter membranes were placed at the base of a 96 well plate (n=3). Trypsin was added to the hADSCs to remove them from cell culture flasks and 8x103 cells we re seeded per well in 200l of media further supplemented with -glycerolphosphate, ascorbic-2-phosphate, dexamethasone and sodium pyrinate. Cells were cultured for 16 days and media was refreshed every second day. In Vitro Cytotoxicity Study by Cell Titer Assay Cell Titer 96 (Promega) assay was perfo rmed to quantify cell pr oliferation at days 5, 11 and 16. Cell media was removed and Cell Titer 96 AQueous One Solution was added to the cells. Quantification was perform ed with a microplate reader according to the manufactures protocol. Fluorescent Microscopy Imaging Following cell culture, cells were fixed with 4% paraformaldehyde for 15 minutes and washed extensively with phosphate buffered saline solution (PBS). Fluorescein isothiocyanate (FITC) phalloidin (Santa Cruz Biotechnology) was used to stain the actin filaments and samples were mounted with VEC TASHIELD HardSet Mounting Media containing DAPI. Statistical Analysis Statistics was performed using MiniT ab. A Tukeys comparison was used to determine differences between timepoints and groups (p<0.05 was considered as statistically different). Data is presented as mean standard deviation. 103

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Results and Discussion Triple-helical single tr opo-collagen molecules spontaneous ly self-assemble into fibers under the right conditions. 191-194 In collagen nanofibers, each triple helix is shifted relative to its molecular neighbor by a multiple of 67 nm in the direction of the helix (See the scheme in Figure 6-1). Later ally, the helices are arranged in a hexagonal pattern in respect to each other within the fiber. This arrangement leads to the characteristic band pattern of collagen fibers. Collagen fibers are most stable at moderately basic pH (9-11) and high ion (especially phosphate) concentrations. In order to obtain minera lized collagen nanofibers, CaCl 2 was added to the feed solution while phosphate was given to the re ceiver solution. The other experimental conditions for the growth of mineralized co llagen fibers were simila r to the experiment fabricating collagen na nofibers (Figure 6-1). The forma tion of mineralized fibers is especially sensitive to the calcium concentration. With lower calcium concentrations (1 mM CaCl 2 in the feed solution), only the inside of the fibers was mineralized, as can be clearly seen by the visible enhancement of the band pattern of the collagen fibers (compare Figures 6-2 a and c). With higher calcium concentrations (up to 5 mM CaCl 2 ), the fibers exhibit mineralized shells (Figure 62 e, f). The shells ap pear to be segmented with each segment having a diameter of r oughly 67 nm, which equals the distance found in the band pattern of collagen fibers. With CaCl 2 concentrations as high as 20 mM, plate-like hydroxyapatite crystal s precipitate in large bundles that are interconnected by collagen. Selected area electron diffraction (SAED) and energy dispersive spectroscopy (EDS) were performed for all experimental conditions. While EDS showed amounts of calcium and phosphate that were roughl y proportional to the amount of CaCl 2 in the feed solution, the mineral phase was always am orphous (see inserts in Figure 6-2). 104

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It is important to note that in these ex periments, collagen does not induce the PILP phase. 188 For this mechanism an acidic polymer would be necessary. In our experiments, collagen fibers form at the same time as the amorphous calcium phosphate. The prevalence of the amorphous phase is a result of the rapid fl ow of the feed solution that results in the creation of a highly supersatu rated phase at the exit of the pores. A rough estimation of the flow rate of the feed solu tion through the membrane gives a flow speed of the order of 100 m/s. In comparison, derived from the diffusion coefficient of collagen 195 and the resulting diffusion length, collagen molecules in dilute solutions move at a speed of about 5 m/s. As a result, t he nanofiber formation only takes around 2 h. For the formation of a crystalline phase of calc ium phosphate, reaction times of at least four days are typical. 188 This fast and coincident fo rmation of nanofibers and particles yields a collagen/calcium phosphate composit e material without the addition of acidic polymers or natural noncollageneous proteins. In a further series of exper iments, synthetic polymer, in stead of naturally occurring biopolymer, was used in the formation of nanofibers. The advantage of using synthetic polymer is that they are much less expensiv e than those natural proteins, which allows the scale-up production for tissue engineering applic ations. Poly(acrylic acid), an acidic and hydrophilic polymer, is selected as the m odel synthetic polymer in our experiments. Poly(acrylic acid) has been successfully used as a mimic to acidic proteins and polysaccharides for facilitating biomineralizat ion. The formation of nanofibers made of calcium carbonate and PAA was investigated. These nanofibers can be easily observed by scanning electron microscopy (SEM). Nanofibers grown in 200 nm pores had a diameter of 110 60 nm for 200 nm pores. The l ength is poorly controlled. It is very likely 105

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that the nanofibers break as a result of t he filtration process or at an unspecified time during the extrusion. While some nanofibers we re as long as 100 m, the length of most fibers is on the order of 20 m. The nanofiber s are evenly distributed across the filter substrate with the occasional occurr ence of bundles (Figure 6-3 a). The structure of the nanofi bers was examined by selected area electron diffraction (SAED), which was coupled to TEM (Figure 6-3 b). The resulting diffractogram shows that the nanofibers were amorphous. Several control experiments were performe d by exchanging one of the reaction components by water to prove that the fi bers indeed consist of calcium carbonate and PAA. Without PAA, uncontrolled prec ipitation of mainly vaterite crystals occurred. Without calcium chloride or without sodium bica rbonate, no nanofiber formation could be observed. One control experiment was performed by reversing the flow direction of the PAA-rich phase. In this experiment, PAA was dissolved in the sodium bicarbonate solution instead of the calcium chloride solu tion. No nanofibers were formed in this experiment. Instead, discs around 10 m in si ze were deposited on the filter substrate (Figure 6-3 c). Those were most likely PILP droplets that had been fla ttened as a result of vacuum filtration. In additional experiments, the Na 2 CO 3 receiver solution was exchanged by Na 2 HPO 4 of the same concentration. Here, too, the formation of nanofibers could be confirmed (Figure 6-3 d). The nanofibers containing calcium phosphate are shorter and appear to be more rounded and softer t han those with calcium carbonate. 106

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As is evident by the above presentation of the results involving PAA as the polymeric component, the synthesis of fibers is governed by the form ation of the PILP phase at the exit of the pores. 181 Upon entering the pores, the PAA chains will likely become axially aligned along the pores as a result of the strong elongational velocity gradients in the entry region of the pores. This alignmen t will persist within the pores because of the presence of shear gradients. The negatively charged carboxylic polymers are surrounded by a cloud of positively charged calcium ions; near the exit of the pores, carbonate ions are added to the diffuse ion cloud around the polymers, giving rise to the PILP phase. While being extruded from the pores, the acidity of the polymer, in concert with the concentrated ions, sustains the metastable state of the amorphous calcium carbonate phase. Unlike the collagen fibers, the nanofibers are a homogeneous phase of an amorphous mineral/polymer com posite without a special pattern as it arises in the self assembly process of collagen. In general, t he PILP phase will pr eferentially form in contact with an additional interface, because of the reduction of the nucleation energy threshold at interfaces. In colloid chemical terms, again, the three-phase contact is energetically favorable in the presence of a finite contact angle. The fibers were tested for thei r ability to support cell gr owth in-vitro using human adipose derived stem cells (hAD SCs) as a model cell line for tissue engineering. This cell type is found in abundance within our body capable of differentiating down the mesenchymal linage and an excellent c andidate for future tissue engineering applications. The Cell Titer 96 proliferation assay dem onstrated increasing proliferation in all groups at all time points with the exception of co llagen nanofibers at da y 16 (Figure 6-4). 107

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There is also a trend indicating that the inclusion of calcium phosphate enhances cell proliferation. Interestingly, only the calcium phosphate c ontaining groups exhibited a statistical increase in alkaline phosphatase (ALP ) activity which is an early indicator of bone cell differentiation (Figure 6-5). T he introduction of calcium phosphate had a profound effect on fiber morphology and mechanical stiffness, which occurs on a scale at which cells can interact. Within our cellular analysis, the increase in CaCl 2 concentration increased proliferation and ALP production. Xie et al. have shown that calcium phosphate can induce osteoblasts differentiation 196 while Sere et al. have shown that by combining calcium phosphate with collagen, ce lls up-regulate matrix production. 197 Cell actin staining indicated intimate contact with the underlying surface and out-stretched cells with connecting filopodia were witnessed (Figure 66). Cells rapidly covered the nanofibrous surface and began to grow in multilayers. Perspective In conclusion, we present a straigh tforward method for t he fabrication of organic/inorganic hybrid nanofibers that was strongly inspired by processes found in biomineralization. Naturally occurring type I collagen and synthetic PAA were selected as the polymeric component Using PAA as the pol ymeric component, nanofibers could be produced that consist of an amorphous mixtur e of calcium phosphate and collagen, or PAA. That this method is generally suited for the fabrication of nanof ibers of different compositions has been demons trated by exchanging the Na 2 HPO 4 solution by Na 2 CO 3 of the same concentration in the receiver solution with PAA as polymeric component. The collagen nanofibers could be mineralized by the formation of amorphous calcium phosphate simultaneous to fiber formation. It was also shown that the fibers supported stem cell growth and proliferation, as an early indication of scaffold biocompatibility. It 108

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should also be noted that the inclusion of calcium phosphat e within the fibers up-regulated stem cell alkali ne phosphatase production. Thus, we believe that this new approach holds much promise in future studies about the production of new nano-structured materials as well as advances in the field of biomineralization. 109

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Figure 6-1. Experimental se tup and proposed model for t he formation of mineralized collagen fibers. 110

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Figure 6-2. (a, b) Unmineralized collagen fibers (c, d) Mineralized collagen fibers (1 mM CaCl 2 ), (e, f) Mineralized collagen fibers (2.5 mM CaCl 2 ) and (g, h) Mineralized collagen fibers (5 mM CaCl 2 ). Inset images in (b, d, f, h) are SAED patterns. 111

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Figure 6-3. (a) Bundle of PAA/CaCO 3 nanofibers (b) TEM micrograph and SAED pattern of a PAA/CaCO 3 (c) Flattened PILP droplets, (d) PAA/Calcium Phosphate nanofibers. Figure 6-4. Proliferat ion of hADSC's on nanofibers. indicate statistical difference between groups at the same timepoint. 112

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Figure 6-5. Alkaline phosphatase production from hADSC's cultured on nanofibers. indicates statistical difference at the same timepoint Figure 6-6. Fluorescent images of hADSCs cultured on nanofibers. The green indicates actin filaments while blue indicates cell nuclei. 113

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CHAPTER 7 CONCLUSIONS We have developed a simple and effi cient method to produce biodegradable nanoparticles and nanofibers for drug delivery and tissue engine ering applications. This novel nanopore-injection (N-I) method is based on the use of nanoporous membranes. Chapter 1 introduced and reviewed all rela tive background information for these studies, including the current progress in nanostructured biomaterials and their applications in drug delivery and tissue engineering, as well as current nanofabrication methods. In Chapter 2, we fabricated unifo rm hydrophobic nanoparticles (silymarin, beta-carotene, and butylated hy droxytoluene) with < 100 nm size through N-I method. The obtained nanoparticles exhibit smaller hydrodynam ic diameter and better dispersibility in aqueous solution, compared with those made through SEDS method. The obtained N-I NPs were amorphous, and showed improved dissolution profile in PBS. In Chapter 3, we successfully prepared ul trafine chitosan nanoparticles by N-I strategy. The obtained nanoparticle has a small size (<30 nm) with narrow size distribution. Particle size increases with flow rate and viscosity of feed solution. Fluorescent dye rhodamine 6G was encapsul ated into chitosan nanoparticles with a encapsulation ratio of about 3 wt%. In Chapter 4, we have successfully prepared MCHBE encaps ulated PLGA-PEG and luciferin encapsulated chitosan nanoparticles by N-I method. These nanoparticles exhibit the ability of sustained release of encapsulated drugs: 7 days for PLGA-PEG/MCHB NPs; 11 days fo r CS/Luc NPs. In vitro cyt otoxicity experiment proved 114

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that PLGA-PEG nanoparticle vector could enhance MCHB delivery efficiency. CS/Luc nanoparticles could be successfully detected in the in vivo bioluminescence imaging. In Chapter 5, biodegradabl e collagen nanofibers are prepared by modified N-I method using PCTE membrane. Diameter of collagen nanofiber could be controlled by the pore size of nanoporous me mbrane. Using the membrane wit h pore size ranging from 50-1000 nm, nanofibers with diam eter ranging from 50-740 nm could be obtained. The nanofibrous scaffold exhibited a strong biom echanical strength. In vitro experiment shows the nanofibrous scaffold supports cardiac stem cell growth and proliferation. In Chapter 6, organic/inor ganic hybrid nanofibers were produced by N-I method, including collagen/calcium phosphate nanofiber, and poly(acrylic acid)/calcium carbonate or calcium phosphate nanofiber. These fibers ar e amorphous. It was also shown that the fibers supported stem cell grow th and proliferation, an early indication of scaffold biocompatibility. The work presented here has demonstrated that N-I method could be used to produce biodegradable nanoparticl es and nanofibers. These obtained nanostructured materials portray a promising future in biomedical applications. Although further studies will be needed to explore the functionalization of those products, the research presented in this dissertation may hopefully provide some insight for related nanomedical studies. 115

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BIOGRAPHICAL SKETCH Peng Guo was born in Liaoyuan, China in 1983. He entered Jilin University at Changchun, China in 2001 as undergraduate student majoring in chemistry. He obtained a Bachelor of Science degree in July 2005. In August 2006, Peng Guo joined Dr. Charles R. Martins research group in the Department of Chemistry at University of Florida. He started his research in the fields of biomedical nanomaterials, especially on the fabrication of biodegradable na nostructure for drug deliver y and tissue engineering. He completed his research in May 2011, obtai ning a Doctor of Philosophy degree in chemistry. 127