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Microfluidic Devices for Isolation of Circulating Tumor Cells

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Title:
Microfluidic Devices for Isolation of Circulating Tumor Cells
Creator:
Sheng, Weian
Place of Publication:
[Gainesville, Fla.]
Florida
Publisher:
University of Florida
Publication Date:
Language:
english
Physical Description:
1 online resource (151 p.)

Thesis/Dissertation Information

Degree:
Doctorate ( Ph.D.)
Degree Grantor:
University of Florida
Degree Disciplines:
Mechanical Engineering
Mechanical and Aerospace Engineering
Committee Chair:
FAN,ZHONGHUI HUGH
Committee Co-Chair:
HAHN,DAVID WORTHINGTON
Committee Members:
ANGELINI,THOMAS ETTOR
TAN,WEIHONG
Graduation Date:
12/13/2013

Subjects

Subjects / Keywords:
Antibodies ( jstor )
Blood ( jstor )
Cancer ( jstor )
Cells ( jstor )
Cytometry ( jstor )
Flow velocity ( jstor )
Fluorescence ( jstor )
Microfluidic devices ( jstor )
Pancreatic cancer ( jstor )
Tumors ( jstor )
Mechanical and Aerospace Engineering -- Dissertations, Academic -- UF
aptamers -- ctcs -- mems -- microfluidics
Genre:
bibliography ( marcgt )
theses ( marcgt )
government publication (state, provincial, terriorial, dependent) ( marcgt )
born-digital ( sobekcm )
Electronic Thesis or Dissertation
Mechanical Engineering thesis, Ph.D.

Notes

Abstract:
Cancer induces high death rate because of the high probability of metastasis. During the progression of metastasis, cancer cells detach from primary tumors or metastatic sites and enter the bloodstream, becoming circulating tumor cells (CTCs). CTCs are thus responsible for the spreads of cancer to distant organs, which lead to cancer-induced death. The level of CTCs can provide valuable information for monitoring cancer status and predicting survival rate of cancer patients. However, CTCs are extraordinarily rare (only a few CTCs in 1 mL blood with billions of blood cells), making their isolation and characterization a formidable technological challenge. Therefore, the objective of this research is to develop microfluidic system-based approaches for efficient isolation of CTCs from blood. Firstly, we developed an aptamer-mediated micropillar-based microfluidic device, for efficiently capturing and enriching rare cancer cells. High-affinity DNA aptamers were used as an alternative capturing agent to antibodies for targeting cancer cells. The device consisted of >59,000 micropillars, which greatly enhanced the interactions between cells and the aptamer-coated surface. With optimized device geometry and flow rate, rare tumor cells were captured from whole blood with high efficiency, purity, throughput, and cell viability. Secondly, we incorporated nanoparticles in microfluidic devices for the enhanced capture of cancer cells. Simultaneous attachment of ~95 DNA aptamers onto each gold nanoparticle surface (forming DNA nanospheres) created an assembly of multivalent binding ligands, with significant enhancement of cell capture efficiency and throughput. The enhanced cell capture also accrues from the increased surface roughness, surface area and ligand density. A high-throughput flat channel device and micromixing device were developed for cancer cell isolation from lysed blood and whole blood, respectively. Finally, we developed a microfluidic geometrically enhanced mixing device for isolation of CTCs from pancreatic cancer patients. We demonstrated the potential utility of the device in monitoring the response to anti-cancer drug treatment in cancer patients. In summary, the microfluidic devices developed in this dissertation provide new means for efficient CTC isolation and accurate CTC enumeration. Since the methods are minimally invasive, the microfluidic devices show great potential for cancer diagnosis, monitoring disease progression and treatment response. ( en )
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.
Thesis:
Thesis (Ph.D.)--University of Florida, 2013.
Local:
Adviser: FAN,ZHONGHUI HUGH.
Local:
Co-adviser: HAHN,DAVID WORTHINGTON.
Electronic Access:
RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2015-12-31
Statement of Responsibility:
by Weian Sheng.

Record Information

Source Institution:
UFRGP
Rights Management:
Applicable rights reserved.
Embargo Date:
12/31/2015
Classification:
LD1780 2013 ( lcc )

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1 MICROFLUIDIC DEVICES FOR ISOLATION OF CIRCULATING TUMOR CELLS By WEIAN SHENG A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTO R OF PHILOSOPHY UNIVERSITY OF FLORIDA 2013

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2 2013 Weian Sheng

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3 To my beloved parents, sisters, and wife

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4 ACKNOWLEDGMENTS I would l ove to express my gratitude to all those who gave me support during my student career, especially my graduate study at the to complete my dissertation without the help from so many people in so many ways First of all, I am deeply thankful to my research advisor Dr. Z. Hugh Fan for al lowing me to join his lab, for his advice s and encouragement for offering me lots of freedom to try my own ideas Without his continuous support and guidance, I w ouldn t be able to finish this dissertation work. I am also sincerely grateful to my committe e members, Dr. David W. Hahn, Dr. Thomas Angelini and Dr. Weihong Tan for their time and effort evaluating my work and their valuable suggestions My thanks and appreciation goes to all the current and former Microfluidics and BioMEMS group members, Dr. R uba Khnouf Dr. Pan Gu, Dr. Ke Liu, Dr. Xin Xu, Dr. Wei Liu, Dr. Xiangjun Zheng, Dr. Jinling Zhang, Dr. Chun wei Wang, Rahul Kamath Imran Sheikh, Ram Mirchandani Kiri Hamaker Shih ming Tsai, Chris Cassano, Shancy Augustine, Kirsten Jackson, Jose Varilla s Kangfu Chen Teodor Georgiev and Sydney Shaouy, for the help I received from them, for the wonderful time I spent with them in the lab I sincerely appreciate all the Dr. Weihong Tan s group members from Department of Chemistry especially to Dr. Kathr yn R. Williams Dr. Meghan B. O'Donoghue Altman, Dr. Qua n Yuan and Dr Xiangling Xiong T heir assistance made my work much smoother. I also owe thanks to Dr. Olorunseun O. Ogunwobi, Dr. Che n Liu and Dr. Thomas J. George from C ollege of Medicine for the col laboration work and help. I also thank the Interdisciplinary Microsystems Group (IMG) members I learned a lot from so many people with different backgrounds I would like to thank all my friends

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5 in the United States and in China especially those friends I met in Gainesville, who made my life wonderful over these years Finally, I would like to specially thank my parents and two elder sisters for their support and unwavering love, which built the past 25 years of my life. I am deeply grat eful to my wife, T ao Chen, for being a fantastic colleague, friend, and lover, who has stood beside me and encouraged me constantly no matter good times or hard times.

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6 TABLE OF CONTENTS page ACKNOWLEDGMENTS ................................ ................................ ................................ .. 4 LIST OF TABLES ................................ ................................ ................................ ............ 9 LIST OF FIGURES ................................ ................................ ................................ ........ 10 ABSTRACT ................................ ................................ ................................ ................... 13 CHAPTER 1 INTRODUCTION ................................ ................................ ................................ .... 15 1.1 Cancer and Circulating Tumor Cells ................................ ................................ 15 1.2 BioMEMS, Lab on a chip and Microfluidics ................................ ...................... 16 1.2.1 Terminology ................................ ................................ ............................. 16 1.2.2 Materials and Fabrication of Microfluidic Devices ................................ .... 17 1.2.3 Advantages of Microfluidic Devices ................................ ......................... 19 1.3 Isolation of CTCs Using Microfluidic Devices ................................ .................... 19 1.4 CTC Targeting Ligands ................................ ................................ ..................... 21 1.4.1 Antibodies ................................ ................................ ................................ 21 1.4.2 Aptamers ................................ ................................ ................................ 21 1.5 Objective and Organization of This Dissertation ................................ ............... 23 2 APTAMER ENABLED EFFICIENT ISOLATION OF CANCER CELLS FROM WHOLE BLOOD USING A MICROFLUIDIC DEVICE ................................ ............ 27 2.1 Background ................................ ................................ ................................ ....... 27 2.2 Device Design, Fabrication and Surface Functionalization ............................... 29 2.2.1 Device Design ................................ ................................ ......................... 29 2.2 .2 Device Fabrication ................................ ................................ ................... 29 2.2.3 Surface Functionalization ................................ ................................ ........ 30 2.2.4 Comparison of Surface Modification Methods ................................ ......... 30 2.3 Cell Isolation and Assay ................................ ................................ .................... 31 2.3.1 Cell Culture ................................ ................................ .............................. 31 2.3.2 Aptamers ................................ ................................ ................................ 32 2.3.3 Cell Capture in a Buffer ................................ ................................ ........... 33 2.3.4 Tumor Cell Isolation from Whole Blood ................................ ................... 34 2.3.5 Instrument Setup ................................ ................................ ..................... 35 2.3.6 Cell Viability ................................ ................................ ............................. 36 2.4 Results and Discussion ................................ ................................ ..................... 37 2.4.1 Cell Aptamer Binding Using Flow Cytometry ................................ ........... 37 2.4.2 Isolation of Lymphocytes in Device ................................ ......................... 38 2.4.3 Ef fects of Channel Depth and Flow Rate ................................ ................. 39 2.4.4 Tumor Cell Isolation from Whole Blood ................................ ................... 41

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7 2.4.5 Cell Viability ................................ ................................ ............................. 42 2.5 Release of Captured Cells ................................ ................................ ................ 43 2.5.1 Motivation ................................ ................................ ................................ 43 2.5.2 Cell Release Using Com plementary DNA ................................ ............... 43 2.5.3 Alternative Cell Release Methods ................................ ........................... 44 2.6 Bonding and Surface Functionalization of Cyclic Olefin Copolymer Microchip for Aptamer Based Cancer Cell Capture ................................ ............. 45 2.6.1 Motivation ................................ ................................ ................................ 45 2.6.2 COC Device Design and Fabrication ................................ ....................... 46 2.6.3 Pressure Free Bonding ................................ ................................ ........... 46 2.6.4 One Step Surface Immobilization of Aptamers on COC .......................... 47 2.6.5 Cancer Cell Capture in COC device ................................ ........................ 48 2.7 Conclusion ................................ ................................ ................................ ........ 48 3 MULTIVALENT DNA NANOSPHERES FOR ENHANCED CAPTURE OF CANCER CELLS IN MICROFLUIDIC DEVICES ................................ .................... 68 3.1 Background ................................ ................................ ................................ ....... 68 3.2 Methods ................................ ................................ ................................ ............ 70 3.2.1 Synthesis and Characterization of Gold Nanoparticle Aptamer Conjugates ................................ ................................ ................................ .... 70 3.2.2 Device Design and Fabrication ................................ ................................ 72 3.2.3 Cell Lines and Buffers ................................ ................................ ............. 73 3.2.4 Flow Cytometric Analysis ................................ ................................ ........ 74 3.2.5 Cell Capture Assay in Microfluidic Devices ................................ ............. 75 3.3 Results and Discussion ................................ ................................ ..................... 76 3.3.1 Synthesis and Characterization of AuNP Aptamer Conjugates ............... 76 3.3.2 Flow Cytometric Analysis Demonstrating High Affinity Binding ............... 77 3.3.3 Enhanced Cancer Cell Capture in a Flat Channel Microdevice ............... 78 3.3.4 Efficient Isolation of Cancer Cells from Whole Blood Using DNA Nanospheres in Micromixer Devices ................................ ............................. 81 3.4 Conclusion ................................ ................................ ................................ ........ 83 4 A MICROFLUIDIC GEOMETRICALLY ENHANCED MIXING CHIP FOR CAPTURE, RELEASE AND CULTURE OF CIRCULATING TUMOR CELLS FROM PANCREATIC CANCER PATIENTS ................................ ........................... 95 4.1 Background ................................ ................................ ................................ ....... 95 4.2 Experimental Section ................................ ................................ ........................ 98 4.2.1 Microfluidic Device Fabrication ................................ ................................ 98 4.2.2 Cell Culture ................................ ................................ .............................. 99 4.2.3 Reagents and Buffers ................................ ................................ .............. 99 4.2.4 Capture of Spiked Tumor Cells in Microfluidic Device ........................... 101 4.2.5 Instrument Setup ................................ ................................ ................... 102 4.2.6 Cell Release and Re culture ................................ ................................ .. 102 4.2. 7 Patient Blood Specimen Collection and Processing .............................. 103 4.3 Results and Discussion ................................ ................................ ................... 104

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8 4.3.1 Target Cell Capture from a Homogenou s Cell Mixture .......................... 104 4.3.2 Micromixer Device Optimization for High Performance Cell Capture .... 105 4.3.3 Tumor Cell Capture from Lyse d Blood and Whole Blood ...................... 106 4.3.4 Cell Release and Cell Viability ................................ ............................... 106 4.3.5 Re culture of Captured Tumor Cells ................................ ...................... 1 07 4.3.6 Isolation of CTCs from Patients with Pancreatic Cancer Using the GEM Chip ................................ ................................ ................................ .... 107 4.3.7 Monitoring Anti cancer Treatment Response Using CTC s .................... 109 4.4 Conclusion ................................ ................................ ................................ ...... 110 5 SUMMARY AND FUTURE WORK ................................ ................................ ....... 128 5.1 Summa ry ................................ ................................ ................................ ........ 128 5.2 F uture Work ................................ ................................ ................................ .... 129 5.2.1 Aptamer Enabled Cancer Cell Isolation ................................ ................. 129 5.2.2 CTC Isolation from Patient Blood ................................ .......................... 130 5.2.3 Other Potential Methods for CTC Detection ................................ .......... 131 APPENDIX A VELOCITY AN D SHEAR STRESS IN MICROCHANNEL ................................ .... 132 B PROTOCOL OF RBC LYSIS FOR WHOLE BLOOD SAMPLES .......................... 134 C BLOOD PROCESSING SAFETY PROTOCOL ................................ ................... 135 LIST OF REFERENCES ................................ ................................ ............................. 137 BIOGRAPHICAL SKETCH ................................ ................................ .......................... 151

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9 LIST OF TABLES Table page 2 1 DNA aptamer sequences, the underlined bases are the toehold sequences ..... 67 3 1 Detailed aptamer sequence information.. ................................ ........................... 94 4 1 Quantification of CTCs per mL of blood among 18 samples from patients with metastatic pancreatic cancer. ................................ ................................ ........... 127 4 2 Quantification of CTCs in healthy donor blood. ................................ ................ 127

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10 LIST OF FIGURES Figure page 2 1 Layout of the microfluidic device.. ................................ ................................ ....... 49 2 2 COMSOL simulation for the micropillars.. ................................ ........................... 50 2 3 Comparison of streamline through different micropillar array s using COMSOL simulation.. ................................ ................................ ................................ ......... 51 2 4 Scheme of surface functionalization and cancer cell capture in the device.. ...... 52 2 5 Confocal fluorescence images of FITC modified aptamers with different conce ntrations immobilized on the surfaces of a microfluidic channel. ............... 52 2 6 Schem e of surface modification of glass or PDMS substrate with streptavidin using silane based method.. ................................ ................................ ............... 53 2 7 Comparison of avidin physical adsorption based method (Adsorption) and silane based method (Silane) toward the immobilization of aptamers. ............... 53 2 8 Schem e of cancer cell capture experiment setup.. ................................ ............. 54 2 9 Counting of cell numbers using ImageJ.. ................................ ............................ 55 2 10 Flow cytometry histograms showing the selective binding of target cells with corresponding aptamers.. ................................ ................................ ................... 56 2 11 Confocal laser scanning microscopy images showing the selective binding of target CEM cells with sgc8 aptamers. ................................ ................................ 57 2 12 Fluorescence image s of cancer cells captured in device.. ................................ .. 58 2 13 Capture efficiency and purity as a funct ion of channel depth and flow rate.. ...... 59 2 14 Tumor cell capture from whole blood using the microfluidic device.. .................. 60 2 15 Fluores cent microscope image shows the viability of captured cells. ................. 61 2 16 Schem e of cell release using complementary DNA sequence. .......................... 61 2 17 Release of captured cells using toehold mediated DNA hybridization.. .............. 62 2 18 Efficient release of captured cells.. ................................ ................................ ..... 63 2 19 Picture of COC microchip and its bonding process.. ................................ .......... 64 2 20 One step surface immobilization of aptamers on COC surface for cell capture 65

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11 2 21 Surface coating of amino aptamers on COC substrate for cell capture.. ............ 66 3 1 Illustration of enhanced cell capture using AuNP aptamer modified surface. ..... 85 3 2 Adsorption and fluorescence spectrum of AuNPs and AuNP aptamer conjugates. ................................ ................................ ................................ ......... 86 3 3 Dynamic light scattering (DLS) analysis of DNA nanospheres.. ......................... 87 3 4 Pictures of the flat channel microdevices. ................................ .......................... 88 3 5 Comparison between a flat channel device and a herringbone groove device for flow mixing.. ................................ ................................ ................................ ... 89 3 6 Flow cytometry shows the strong and specific binding of AuNP sgc8 aptamer conjugates with target CEM cells. ................................ ................................ ....... 90 3 7 Comparison of AuNP aptamer and aptamer alone based CEM cell capture in a flat channel device. ................................ ................................ .......................... 91 3 8 Capture of CEM cells from blood using DNA nanospheres in the flat channel devic e. ................................ ................................ ................................ ................ 92 3 9 Isolation of cancer cells from whole blood using DNA nanospheres in micromixer device. ................................ ................................ .............................. 93 4 1 Picture and design of th e microfluidic geometrically enhanced mixing chip (GEM chip). ................................ ................................ ................................ ...... 112 4 2 Picture of the photomask used for fabrication of the two layer microfluidic mixing device. ................................ ................................ ................................ 113 4 3 Schem e of SU 8 mold fabrication process for the two layer mixing device. ..... 114 4 4 3 D view and SEM image of the herringbone micromixer structure insid e the microfluidic channel. ................................ ................................ ......................... 115 4 5 A high aspect ratio micropillar device tested for cell capture ............................ 116 4 6 Schematic illustratio n of flow and shear stress on cells inside channel. ........... 117 4 7 Flow cytometry test of anti EpCAM binding with different types of pancreatic cancer cells.. ................................ ................................ ................................ ..... 118 4 8 Representative image s of cells before and after capture. ................................ 119 4 9 L3.6pl cell capture effici ency as a function of flow rate ................................ ..... 120

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12 4 10 Comparisons of capture efficiency and purity of L3.6pl cells with different groove width ................................ ................................ ................................ ..... 120 4 11 Regression analysis of the number of the L3.6pl cells capt ured by the microfluidic device versus the number of the cells spiked in 1 mL of lysed or whole blood. ................................ ................................ ................................ ..... 121 4 12 Cell release and cell viability testing. ................................ ................................ 122 4 13 Phase contrast micrograph (10 ) of re cultured cells ................................ ...... 123 4 14 Fluorescence microscope images (40 ) of CTCs captured from patient blood. ................................ ................................ ................................ ................ 123 4 15 CTC number under different treatment cycle correlates with tumor size by CT scans. ................................ ................................ ................................ ............. 124 4 1 6 Fluorescence image s of white blood cells, wit h DAPI positive, cytokeratin negative and CD45 positive. ................................ ................................ ............. 125 4 1 7 Fluorescence image s of circulating tumor cells, with DAPI positive, cytokeratin positive and CD45 negative. ................................ .......................... 126

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13 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 MICROFLUIDIC DEVICES FOR ISOLATION OF CIRCULATING TUMOR CELLS By Weian Sheng December 2013 Chair: Z. Hugh Fan Major: Mechanical Engineering C ancer induce s high death rate because of the high probability of metastasis During the progression of metastasis, cancer cells detach from primary tumor s or metastatic sites and enter the bloodstream, becoming circulating tumor cells ( CTCs ) CTCs are thus responsible for the spread s of cancer to distant organs, which lead to cancer induced death. The level of CTCs can provide valuable information for monitorin g cancer status and predicting survival rate of cancer patients However, CTCs are extraordinar ily rare (only a few CTCs in 1 mL blood with billions of blood cells) making their isolation and characterization a formidable technological challenge. Therefor e, t h e objective of this research is to develop microfluidic system based approaches for efficient isolation of CTCs from blood. First ly we developed an aptamer mediated micropillar based microfluidic device, for efficiently captur ing and enriching rare cancer cells High affinity DNA aptamers we re used as an alternative capturing agent to antibodies for targeting cancer cells. The device consist ed of >59,000 micropillars, which greatly enhance d the interactions between cells and the aptamer coated surfac e. With optimized device geometry and

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14 flow rate rare tumor cells were captured from whole blood with high efficiency, purity, throughput and cell vi ability. Secondly, we incorporated nanoparticles in microfluidic devices for the enhanced capture of canc er cells. Simultaneous attachment of ~ 95 DNA aptamers onto each gold nanoparticle surface (forming DNA nanospheres) created an assembly of multivalent binding ligands with significant enhancement of cell capture efficiency and throughput. The enha nced cel l capture also accrues from the increased surface roughness, surface area and ligand density. A high throughput flat channel device and micro mixing device were developed for cancer cell isolation from lysed blood and whole blood respectively. Finally, we developed a microfluidic geometrically enhanced mixing device for isolation of CTCs from pancreatic cancer patient s. W e demonstrated the potential utility of the device in monitoring the response to anti cancer drug treatment in cancer patients. In summar y, the microfluidic devices developed in this dissertation provide new means for efficient CTC isolation and accurate CTC enumeration. Since the methods are minimal ly invasive, the microfluidic devices show great potential for cancer diagnosis, monitoring disease progression and treatment response.

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15 CHAPTER 1 INTRODUCTION 1.1 Cancer and Circulating Tumor Cells Cancer is a major cause of death worldwide and is a leading public health problem. 1 In the United States, one in four deaths is caused by cancer. It is estimated that > 1.6 million new cases of cancer patients will be diagnosed and ~ 5 8 0 ,000 cancer related deaths (both men and women) will occur in the United States in 201 3 2 The direct medical care costs for all cancers are > $10 0 billion, according to the National Cancer Institute. A primary reason of cancer induced death is metastasis, the spread of cancer cells from the primary tumor to other organs (e.g., lungs, bone s and liver are the most common sites of metastasis). 3 During the progression of metastasis, cancer cells detach from the primary tumor, penetrate the blood vessels and enter into the bloodstream, becom ing circulating tumor cells (CTCs). 4 6 CTCs circulate through the lymphatic system and the bloodstream and migrate to other parts of the body. Eventually, the CTCs extravasate from blood vessels at a distant location and start to proliferate and stimulate angiogenesis, thereby spreading the cancer to other locations and tissues in the body. 7 8 Hence, CTCs hold the key to understanding cancer metastasis and can serve as a potential cancer biomarker. The clinical significance of CTCs towards non hematologic cancer has been widely demonstrated. 9 12 Specifically, CTCs can be used for: 1) early detection of metastasis; 2) monitoring of treatment response; 3) therapeutic design; 4) discovery of biomarkers and drugs ; 5) cancer prognosis. Isolation and enrichment of CTCs will

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16 enable the subsequent cellular and genetic study of CTCs, thus providing a means for discovery of biomarkers and understandin g the biology of metastasis. The high death rate of cancer is associated with the difficulty in cancer diagnosis. Currently, a variety of cancer diagnostic methods have been used in clinics, including medical imaging (e.g., X ray computed tomography, or C T scans), endoscopy, and tumor biopsy. However, most of the se diagnostic methods are invasive and expensive, and some of them are inaccurate. For example, biopsy, the current gold standard of cancer diagnosis, involves removal of tissue or cells from the b ody and examination by experienced surgeons and pathologists. The invasive nature of biopsy prevents patients from being tested in an ongoing or repetitive basis. CTC enumeration and examination, on the other hand, is much less invasive, with only 5 10 mL of patient blood needed; it time monitoring of therapeutic response. However, CTCs are ex traordinar ily rare in nature. Typically, there are only around 13 Thus, the isolation and characterization of CTC s is a major technological challenge. Recently, microfluidic devices provide unique opportunities for rare cell isolation and detection; 14 and they have been used for size based separation, affinity based cell sorting and flow cytometry. 15 17 1.2 BioMEMS, Lab on a chip and Microfluidics 1.2.1 Terminology Microelectromechanical systems (MEMS), also called Microsystem Technology (MST) in Europe or M icromachines in Japan, are miniaturized systems or devices that

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17 combin e electrical and mechanical components with sizes from the sub micrometer to millimeter level. MEMS extend the fabrication technology for semiconductor devices and integrated circuits, with the addition of mechanical components such as beams, springs and d iaphragms. Examples of MEMS are in k ject printer cartridges, accelerometers, gyroscopes, microphones, etc. 18 19 MEMS technology also includes BioMEMS (biological or biomedical MEMS), which are used in biological and chemical analysis. While engineers like to use the term Bi oMEMS to stress its microfabrication technology, chemists prefer to use Lab on a chip and micro total analytical system (TAS) 20 to emphasize that multiple laboratory functions are integrated on a single chip. Microfluidics, however, is an even broader term that desc ribes all fluid handling on the sub millimeter scale, including microchannels, micropumps, microvalves, microdroplets, of systems that process or manipulate small (10 9 to 10 18 litres) amounts of fluids, using 21 Hence, microfluidics is now much more than a subset of MEMS, and is becoming a multidisciplinary field itself, uniquely identified by its materials and fabrication technologies, encompassing the fiel ds of engineering, physics, chemistry and biotechnology. 1.2.2 Materials and Fabrication of Microfluidic Devices Initially, traditional MEMS materials, such as silicon, were used for microfluidic device fabrication. Microfabrication and micromachining tech niques for MEMS were applied for fabrication of such silicon microfluidic devices. 22 23 Photolithography was used for the patterning required in microfabrication; and microstructures were made by subsequent etching. 24 Later, glass was extensively used because of its easy availability and optical transparency. However, because the cost for fabrication of silicon and glass

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18 device was too high, polymer or plastic materials, such as polydimethylsiloxane (PDMS) 25 and thermoplastics, 26 have been playing an increasing ly important role for microfluidic devices, because of their low cost, mass production, optical clarity and biocompatibility. Soft lithography has been rapidly developed for PDMS device fabrication. 27 29 30 which can rapidly prototype devices on various substrates, including planar, curved, flexible and soft substrates, with different kinds of ela as PDMS. Four major steps are generally involved in a soft lithography procedure. 1) Pattern design. The pattern can be designed precisely using a number of computer aided design (CAD) programs (e.g., AutoCAD from Autodesk). 2) Fabrication of the mask. Generally, there are two kinds of masks: transparency mask, a photomask printed on transparency film, and chrome mask, a glass or quartz plate patterned with opaque chrome on its surface. A transparency mask with resolution up to 25,400 d.p.i can be obtained from high resolution printing companies (e.g., CAD/ART Services, Inc.). The smallest feature a transparency mask can produce is ~ 10 m. A chrome mask can produce even smal ler features (~1 m), but with substantially higher cost. Chrome masks are also available to be ordered from commercial suppliers such as Photo Sciences Inc. 3) Fabrication of the master. Conventional photolithography is the primary technique for fabricati ng a master that contains patterned relief structures on the surface, typically with feature sizes larger than 1 m. The fabrication of silicon master starts with spin coating of a thick photoresist (e.g., SU 8) on a wafer, followed by

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19 UV light exposure un der the mask, and the subsequent development. 4) Fabrication of the PDMS device. An elastomeric slab with patterns as relief structures on its surface is the key element of soft lithography. The slab is typically fabricated by casting a liquid PDMS precurs or against the master with the complementary structures patterned on its surface. Sylgard 184 from Dow Corning has been most commonly used commercially available PDMS. Since one silicon master can be used hundreds of times for rapid ly fabricating many PDMS devices, soft lithography results in much lower cost than traditional photolithography in mass production. 1.2.3 Advantages of Microfluidic Devices With their small size, microfluidic devices provide many advantages for biological and chemical analysis, 31 including: 1) small quanti ties of fluids required, resulting in lower cost of reagents, lower sample volumes for diagnostics, and less waste; 2) small dimensions leading to low power consumption and versatility in design; 3) high resolution and detection sensitivity, because of the high surface area to volume ratio, rapid mass transfer and interaction, and short diffusion distances; 4) potential for parallel operation with high throughput and capability to integrate with other miniaturized devices or components (e.g., mixers and det ectors) ; 5) low cost, mass production allowing disposability, and resulting in elimination of cross contamination; and 6) portability, promising for point of care diagnostics. These advantages have propelled microfluidic devices from research laboratories to clinical research and industry in the past 20 years. 1. 3 Isolation of CTCs Using Microfluidic Devices Currently, the most commonly used cell isolation and sorting methods are fluorescence activated cell sorting (FACS) and magnetic activated cell sortin g (MACS).

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20 In FACS or flow cytometry, which is widely used in clinical diagnosis, cells are fluorescently labeled with specific dyes or antibodies and then sorted based on light scattering and fluorescence intensity. 32 However, flow cytometry produces many false negatives, which increase the limit of detect ion, a severe problem when detecting rare cells. Also flow cytometry equipment is large and quite expensive (>$250,000), requiring trained personnel for instrument operation and sample preparation. MACS is also widely used to isolate cells using antibody c oated magnetic beads. Target cells conjugated with magnetic beads are separated using magnetic force. 9 33 35 The FDA approved commercial CellSearch system from Veridex 36 37 is based on this immunomagnetic beads bas ed cell isolation. Other cell isolation methods include sorting based on physical properties such as size and density and micromachine enabled isolation. 38 39 However, these methods have the disadvantage of low sensitivity, low purity or high cost. Microfluidic devices, on the other hand, p rovide an inexpensive means to isolate CTCs with high sensitivity. Considerable research has been performed with microfluidic devices using size based, 40 41 dielectrophoresis based, 42 fluorescence activated 43 44 and magnetic based 45 46 cell sorting. Recently, microfluidic devices with high affinity ligands, primarily antibodies, have emerged as a distinctive method for isolation of rare cells. 47 48 In general, a microchannel was first functionalized with specific antibodies; cell suspension or whole blood was then driven through the device, resulting in target cell capture. Topographic features such as microposts, 49 sinusoid shapes, 48 silicon nanopillars 50 and chaotic mixers 51 have been applied to enhance the interaction between cells and the antibody coated device surface s

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21 1. 4 CTC Targeting Ligands 1.4.1 Antibodies Circulati ng tumor cells express certain receptors which can be used as CTC biomarker s the molecule s that can be used for detect ing and isolat ing CTCs. With the biomarker information, corresponding antibodies can be develop ed for CTC isolation. The most notable CT C biomarker is epithelial cell adhesion molecule ( EpCAM ) since CTCs are essentially epithelial cells detached from primary tumor with epithelial tissue EpCAM is a trans membrane protein expressed on most of normal epithelial cells and is highly expressed in the majority of epithelial cancers, including breast, colorectal, prostate, and pancreatic cancers. 52 53 Thus, monoclonal anti EpCAM antibody has been widely used for CTC isolation. However, it is believed that a significant number of CTCs go through the epithelial mesenchymal transition (EMT) during the progression of metastasis and lo s e epithelial characteristics and become mesenchymal cells thus losing EpCAM receptors on their cell membrane 54 Other CTC biomarker s include h uman e pidermal g rowth f actor r eceptor 2 ( HER2 ) epidermal growth factor receptor (EGFR), and the calcium dependent cadherins 55 For prostate cancer, p rostate specific antigen (PSA) and p rostate specific membrane antigen (PSMA) are highly specific prostate cancer biomarkers and have been extensively used. 51 1.4.2 Aptamers While only a few antibodies have been identified for tumor cells and cancer cell lines, a number of DNA aptamers with high affi nity and excellent selectivity have already been selected for numerous cancers. Aptamers are short (usually 20 100 bases) single stranded DNA or RNA oligonucleotides (or sometimes peptide molecules) that bind to a specific target molecule. Aptamers are gen erated using an in vitro selection process

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22 termed SELEX (Systematic Evolution of Ligands by EXponential enrichment). 56 57 The SELEX procedure usually begins with a random library of 10 13 10 15 DNA or RNA sequences, followed by an iterative process with positive selections and co unter selections for specific amplification of sequences having high binding affinity to the targets. By folding into unique secondary or tertiary structures, aptamers can bind with their target s with high affinity (with dissociation constants from M to p M) and recognize their targets with a specificity that is comparable to antibodies. Aptamers have been selected for a broad range of targets including metal ions, small organic molecules, proteins, biological cells, viruses and bacteria. 58 59 Compared with antibodies, aptamers provide significant advantages, including: 1) rapid and reproducible production by c hemical synthesis; 2) capability of selecting for virtually any target at reasonable cost; 3) long term stability; 4) easy modification with different functional groups or dyes. Because of these advantages, aptamers are becoming the next generation antibod y like molecular probes for diagnostic and clinical application. 60 62 Based on the SELEX process, cell SELEX 63 has been developed in the laboratory of Dr. Weihong Tan, our collaborator, to generate a panel of aptamers targeting different types of cancer cells, including leukemia, liver cancer, small cell lung cancer and colorectal cancer. These aptamers have the additional advantage of selection without preknowledge of the biomarkers on the cancer cell surface, while making it possible to identify the biomarkers after se lection. For example, an aptamer called sgc8 has been selected for CEM cells (human acute lymphoblastic leukemia) with a dissociation constant K d =0.79 0.15 nM. Human protein tyrosine kinase 7 (PTK7)

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23 was later identified as the membrane receptor which bind s with sgc8 aptamer. It was found that CEM cells have a high expression level of PTK7 with 1300 190 receptors per 1 m 2 of cell surface. 64 1. 5 Objective and Organization of This Dissertation Isola tion and enumeration of circulating tumor cells (CTCs) are of great importance for cancer diagnosis and disease monitoring holds the promise to usher in a new era of personalized therapeutic treatments and real time monito ring for cancer patients. But t he extreme paucity of CTCs in blood make s their isolation a formidable technological challenging. The major objective of this research is to study and develop novel microfluidic systems to address the challenges of CTC isolat ion: 1) maximizing the capture of target cells (i.e., CTCs); 2) minimizing the capture of non target cells (i.e., leukocytes and erythrocyte s ) T h e research aims to develop inexpensive microfluidic chips for high performance CTC capture, with accurate CTC counting capability for noninvasive cancer diagnosis and monitoring. This dissertation also aims to use novel DNA aptamer based cancer biomarkers and integrate them with microfluidic system for sensitive cancer cell isolation. The significance of this wor k mainly lies in three aspects. First we developed two generation s of high performance microfluidic devices for efficient tumor cell capture: a unique isotropically etched elliptical micropillar array device with optimized channel geometry, and a geometri cally enhanced mixing microfluidic device. Second, DNA aptamer s with high affinity and excellent selectivity were used for cancer cell capture, while most researche r s are using antibodies; also multivalent DNA nanospheres were used for the first time for e nhanced cancer cell capture. T hird, the geometrically enhanced mixing chip enabled efficient CTC isolation and accurate CTC enumeration

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24 from patients wit h metastatic pancreatic cancer, a disease where biopsies are difficult and the commercial CellSearch sy stem is inefficient. The microfluidic systems developed in this dissertation are highly novel and enabled efficient isolation of CTCs with high efficiency, high purity, high throughput and high cell viability The rest of this dissertation is outlined as f ollows. I n chapter 2, an aptamer mediated, micropillar based microfluidic device is developed, which is able to efficiently isolate tumor cell s from unprocessed whole blood. High affinity aptamers were used as an alternative to antibodies for cancer cell i solation. The microscope slide sized device consists of >59,000 micropillars, which enhanced the probability of the interactions between aptamers and target cancer cells. The device geometry and the flow rate were investigated and optimized by studying the ir effects on the isolation of target leukemia cells from a cell mixture. The device yielded a capture efficiency of >95% and a purity of ~81% at the flow rate of 600 nL/s. Then the device was exploited for isolating colorectal tumor cells from non proces sed whole blood; as few as 10 tumor cells were capt ured from 1 mL of whole blood. The problem of low throughput of a typical microfluidic device was also addressed by processing 1 mL of blood within 28 minutes. In addition, ~93% of the captured cells were found viable, making them suitable for subsequent molecular and cellular studies. Further more captured cells were efficiently released using toehold mediated complementary DNA sequence s which competitively hybridize with cell bound DNA aptamers. C hapter 3 describes the development of a platform combining multivalent DNA aptamer nanospheres with microfluidic device s for efficient isolat ion of cancer cells from blood. Gold nanoparticles (AuNPs) were used as an efficient multivalent platform for

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25 assembling a number of aptamers for high efficiency cell capture Up to 95 aptamers were attached onto each AuNP, resulting in enhanced molecular recognition capability. An 39 fold increase in binding affinity was confirmed by flow cytometry for AuNP aptamer conjugates ( AuNP aptamer ) when compared with aptamer alone. With a laminar flow flat channel microfluidic device, the capture efficiency of human acute leukemia cells from a cell mixture in buffer increased from 49% using aptamer alone to 92% using AuNP aptamer. AuN P aptamer was also utilized in a microfluidic device with herringbone mixing microstructures for isolation of leukemia cells in whole blood. The cell capture efficiency was also significantly increased with the AuNP aptamer over aptamer alone, especially a t high flow rates. The results show that the platform combining DNA nanostructures with microfluidics has a great potential for sensitive isolation of CTC s, and is promising for cancer diagnosis and prognosis. Chapter 4 describes the development of a geom etrically enhanced mixing (GEM) chip for high efficiency and high purity tumor cell capture. The release and culture of the captured tumor cells were also successfully demonstrated. T he high performance microchip is based on geometrically optimized micromi xer structures, which enhance the transverse flow and flow folding, maximizing the interaction between CTCs and antibody coated surfaces. With the optimized channel geometry and flow rate, the capture efficiency reached >9 0 % with a purity of > 84% when capt uring spiked tumor cells in buffer. The system was further validated by isolatin g a wide range of spiked tumor cells (50 50,000) in 1 mL of lysed blood and whole blood. With the combination of trypsinization and high flow rate washing, captured tumor cells were efficiently released. T he released cells were viable and able to proliferate, and showed n egligible difference

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26 compared with intact cells that were not subjected to the capture and release process. Furthermore, the device was applied for detecting CT Cs from metastatic pancreatic cancer patient s ; and CTCs were found in 1 7 out of 1 8 samples (>94%). T he potential utility of the device in monitoring the response to anti cancer drug treatment in pancreatic cancer patients w as also tested, and the CT C numbers correlated with the clinical computed tomograms (CT scans) of tumors. The presented technology shows great promise for accurate CTC enumeration, biological studies of CTCs and cancer metastasis, as well as for cancer diagnosis and treatment monit oring. Finally, the work is summarized in chapter 5 and future work and directions are discussed.

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27 CHAPTER 2 APTAMER ENABLED EFFICIENT ISOLATION OF CANCER CELLS FROM WHOLE BLOOD USING A MICROFLUIDIC DEVICE 2.1 Background Metastases from primary tumors a re the leading causes of death for non hematological cancers. 65 During the progression of metastasis, cancer cells shed from solid tumors and enter the bloodstream, beco ming circulating tumor cells (CTCs), which has a potential to serve as important biomarkers for early diagnosis of cancer metastases. 66 67 While most current methods for cancer diagnosis require invasive biopsy followed by molecular analysis, CTC enumeration is less invasive and provides a means for cancer diagnosis and prognosis, as well as for monito ring the p rogress of treatment. However, CTCs are extremely rare, comprising only a few out of >10 9 hematological cells in 1 mL of blood, making their isolation and characterization a significant technological challenge. Recently, a variety of techniques have been developed for CTC isolation and detection, 68 ranging from the use of immu nomagnetic beads (e.g., CellSearch from Veridex), and size based filtration systems, to microfluidic devices. 40 45 69 71 Among these methods, microfluidic devices with high affinity ligands, primarily antibodies, have provided unique opportunities for detecting CTCs from patient blood. 49 72 However, only a few antibodies have been identified for tumor ce lls and cancer cell lines. In contrast, a number of DNA aptamers with high affinity and excellent selectivity have been selected for numerous cancers. 73 Aptamers are single stranded oligonucleotides that can recogniz e and bind to their target cells by folding into unique secondary or tertiary Part of this cha Aptamer Enabled Efficient Isolation of Cancer Cells from Whole Blood Using a Microfluidic Device, Analytical Chemistry 2012, 84, 4199 4206

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28 structures. They can be easily generated using an in vitro selection process termed cell SELEX (Systematic Evolution of Ligands by EXponential enrichment). Previously, our groups reported a flat channel polydimethylsiloxane (PDMS) device and utilized it to capture target cancer cells from a mixture of target and control cancer cells (1:1 ratio), with ~80% capture efficiency. 74 75 To realize clinical utility, however, a device must be capable of isolating a few tumor cells from milliliters of whole blood (>10 9 cells). As a result, the devic e must possess high capture efficiency (the percentage of tumor cells isolated relative to total tumor cells present), satisfactory cell purity (the percentage of target tumor cells in the cells isolated), and sufficient throughput (the amount of blood pro cessed in a certain period of time). 76 In addition, the captured cells are preferred to remain viable so that they can be further analyzed at the cellular and molecular level 77 (e.g., to study apopt osis of tumor cells). Herein, we report our development of an aptamer functionalized, micropillar based microfluidic device that isolates cancer cells from unprocessed whole blood with the required metrics mentioned above. Aptamers with specific binding t o cancer cells of interest are used as an alternative to antibodies that have been often used for CTC isolation. The micropillars in the microchannel enhanced the probability of the interactions between the cells and the aptamers coated on the channel/pill ar surfaces, resulti ng in high capture efficiency. After optimizing the geometry of the micropillars, efficient isolation of a few tumor cells from whole blood was achieved with sufficient throughput and high cell viability. 78

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29 2.2 Device Design, Fabrication and Surface Functionalization 2.2.1 Device Design The glass micropillar device was designed to be the size of a microscope slide, consisting of eight parallel channels with an array of >59,000 isotropically etch ed, elliptical micropillars as shown in Figure 2 1. The geometric design of the micropillar array was inspired by the deterministic lateral displacement based particle separation, 79 81 in which the flow streamlines are distorted to enhance cell micropillar interactions. The dimension of the elliptical pillars is 30 m (major axis) 15 m (minor axis) 32 m (height), with an interpillar distance of 80 m (center t o center) and an 80 m shift after every 3 rows in the direction of the minor axis. To characterize the design of the micropillar device, computational fluid dynamics simulation was performed using COMSOL Multiphysics (COMSOL, Inc, Burlington, MA) to study the effects of the micropillar geometry and the arrangement of micropillars. The streamlines of the flow across the micropillar array are shown in Figure 2 2, with assumptions of an incompressible fully developed laminar flow, initial velocity at 1 mm/s, and Reynolds number ( Re ) at 0.1. Figure 2 3 shows the comparison of streamlines between the micropillar array with shifts and without shifts. 2.2.2 D evice Fabrication The glass devices were fabricated according to the procedures reported previously. 82 83 In brief, the layout of the device was designed in AutoCAD and then sent to Photo Sciences (Torrance, CA) t o produce a chrome photomask. Glass substrates coated with chromium and photoresist layers were purchased from TELIC (Valencia, CA). The pattern on the photomask was transferred to the glass substrate via photolithography. The glass substrate was then chem ically etched to a channel depth of

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30 24 to 44 m using a mixture of HF, HNO 3 and H 2 O. The channel depth was measured using a Dektak 150 profilometer, and the depth was controlled by the etching time. The glass substrate was then sealed with a 5 mm thick PDM S sheet, fabricated from Sylgard 184 reagents (Dow Corning, Midland, MI) according to the instructions of the manufacturer. Inlet and outlet wells were created at the channel ends by punching holes in the PDMS sheet. 2.2.3 Surface F unctionalization The d evice was functionalized with aptamers through a two step surface modification (Figure 2 4 ): physical adsorption of avidin onto the glass surface (1 5 minute incubation) and immobilization of biotinylated aptamers via biotin avidin interaction (1 5 minute in cubation). Target cancer cells are captured due to the specific binding between cell surface receptors and aptamers. To demonstrate the immobilization of DNA aptamers onto the surface of a microchannel, biotinylated aptamers labeled with fluorescein isothi ocyanate (FITC) was introduced into the channel and confocal microscope images were taken to measure the fluoresc ence intensity on the surface. As shown in Figure 2 5 the fluorescence signal increased with increasing FITC aptamer concentration (after thor ough washing), proving that aptamers were successfully immobilized on the avidin modified surfaces and the amount of aptamers immobilized is dependent on the aptamer concentration. 2.2.4 Comparison of Surface Modification Methods The above mentioned physic al adsorption based surface treatment method wa s compared with a silane based method one commonly used surface modi fication for silicon/glass/PDMS For the silane based method shown in Figure 2 6 the microchannels were first treated with 4% (v/v) 3 merc aptopropyl trimethoxysilane

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31 (MP TM S ) ( Sigma Aldrich St. Louis, MO ) in ethanol at room temperature for 1 hour followed by incubation with 0. 2 mM N y maleimidobutyryloxy succinimide ester (GMBS ) ( P ierce Biotechnology, Rockford, IL ) in ethanol for 30 min at room temperature GMBS serves as the crosslinker for the sulfhydryl group on MPTMS to the amine group on streptavidin or NeutrAvidin. Then, the device was incubated with 10 g/mL streptavidin in PBS for 30 min to attach streptavidin to the GMBS. Afterwards biotinylated aptamers labeled with FITC were introduced into the channel and fluorescence microscopy image was taken to measure the fluorescence intensity. Then we compared the amount of aptamers immobilized between this silane based method and the abov e physical adsorption based method. Results show that silane based method is similar as physical adsorption for glass surface, while physical adsorption gives high er signal than silane based method on PDMS surface as shown in Figure 2 7 T h e silane based surface modification is widely used and proved to be effective However, it involves 2 4 hours of surface treatment with m ultiple incubation and rinsing steps In addition, the plasma treatment of the device surface made the device permanently bonded, and the device was not able to be cleaned and reused. On the other hand, surface modification using avidin physical adsorption wa s facile and robust, with reusable devices, thus it was chosen for future experiments 2.3 Cell Isolation and Assay 2.3.1 Cell Cult ure CCRF CEM cells (CCL 119, T cell line, human acute lymphoblastic leukemia), Ramos cells (CRL 1596, B colorectal adenocarcinoma) and HCT 116 cells (colorectal carcinoma) were purchased from Amer ican Type Culture Collection (ATCC). CEM, Ramos and DLD 1 cells were

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32 cultured in RPMI medium 1640 (ATCC) supplemented with 10% Fetal bovine serum (FBS, heat inactivated; GIBCO) and 100 units/mL penicillin streptomycin (PS, Cellgro, Manassas, VA); HCT 116 c sam e concentration of FBS and PS. 2 atmosphere. The colorectal cancer cell lines were grown as adherent monolayers in 100 mm 20 mm cul ture dishes to 95% confluence. Cells were washed in the dish with Aldrich, St. Louis, MO), dissociated by 0.25% trypsin treatment (2 min) and seeded into culture dishes at a low concentration. 2.3.2 Aptamers DNA aptamers w ere synthesized in house. Aptamer sequences were as follows, ATC TAA CTG CTG CGC CGC CGG GAA AAT ACT GTA CGG TTA GAT TTT TTT TTT AAC ACC GTG GAG GAT AGT TCG GTG GCT GTT CAG GGT CTC CTC CCG GTG TTT TTT TTT T biotin; KDED2a TGC CCG CGA AAA CTG CTA TTA CGT GTG AGA GGA AAG ATC ACG CGG GTT CGT GGA CAC GGT TTT TTT TTT T ATC CAG AGT GAC GCA GCA GGG GAG GCG AGA GCG CAC AAT AAC GAT GGT TGG GAC CCA ACT GTT TGG ACA CGG TGG CTT AGT TTT TTT TTT T biotin. For flow cytometry and fluorescence (FITC) or carboxytetrame thylrhodamine (TAMRA). All aptamers were synthesized using an ABI3400 DNA/RNA synthesizer (Applied Biosystems, Car lsbad, CA) with reagents purchased from Glen Research (Sterling, VA). DNA purification was performed with a ProStar HPLC (Varian, Walnut Creek, CA) using a C18 column (Econosil, 5U,

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33 250 4.6 mm) from Alltech Associates (Deerfield, IL). DNA concentration w as determined by UV Vis measurements using a Cary Bio 300 UV spectrometer (Varian). The specific binding of aptamers and target cells was verified using confocal fluorescence microscopy and flow cytometry. For confocal fluorescence microscopy, cells were f irst incubated with 250 nM TAMRA labeled aptamers in the binding buffer. After washing three times with the washing buffer (DPBS with 4.5 g/L glucose and 5 mM MgCl 2 ), fluorescence microscope images were take n using a confocal microscope. Flow cytometry was performed with a FACScan cytometer (BD Immunocyt ometry Systems, San Jose, CA). Briefly, 200,000 cells were incubated with FITC labeled DNA aptamers at 250 nM for 15 min in 200 L of the binding buffer (or the capturing buffer as specified later), and 10,0 00 counts were measured in the flow cytometer. 2.3.3 Cell Capture in a Buffer Immediately before experiments, cells were rinsed with the washing buffer and resuspended at 10 6 cells/mL. were treated wi th Vybrant DiI or Vybrant DiD cell labeling solutions (Invitrogen, Carlsbad, CA) for 5 min at 37 C, then rinsed with the washing buffer, and resuspended at 10 6 ce lls/mL in the capturing buffer. Labeled cells were stored on ice and further diluted to the d esired conc entrations before experiments. Target CEM cells were spiked into the control Ramos cells to form a final concentration of 10 ,0 00 cells/mL for CEM cells and 10 6 cells/mL for Ramos cells. To initiate cell capture experiments, one channel volume o f 1 mg/mL avidin (Invitrogen, Carlsbad, CA) in phosphate buffered saline (PBS) was first introduced into the device, followed by incubation for 1 5 min and then three rinses with the binding buffer [washing buffer supplemented with yeast tRNA (0.1 mg/mL; Si gma Aldrich),

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34 bovine s erum albumin (BSA) (1 mg/mL)]. Then, one channel volume of 30 M biotinylated sgc8 aptamer with a poly thymine (10 T) linker was introduced into the device and incubated for 1 5 min, followed by three times of rinsing with the binding b uffer. Finally, 1 mL of a mixture of CEM cells (target) and Ramos cells (control) in the capturing buffer was pumped into the channel at a flow rate of 600 nL/s (or other flow rates specified in the text). To prevent cells from settling in the cell suspen sion while continuously pumping, a capturing buffer was prepared by mixing 1:1 volume ratio of the binding buffer and Histopaque 1119 (Sigma Aldrich). The density of the capturing buffer was approximately 1.06 g/ml, which was close to the density of blood and cells. Histopaque 1119 increased the viscosity of the capturing buffer to that of whole blood to mimic the situation of isolating cells from whole blood. 84 The BSA in the capturing buffer can passivate the surfaces to reduce the no nspecific adsorption of cells in the channel. At the end of the experiment, the microchannel was washed three times with the binding buffer, followed by taking fluorescent images for the determinati on of the cell concentrations. For the study of capturing Ramos cells using TD05 aptamer, CEM cel ls were used as control cells. For the study of capturing DLD 1 cells and HCT 116 cells using aptamers KDED2a 3 and KCHA10, Ramos cells were ag ain used as the control cells. The concentration of all target cells was a t 10 0 00 cells/mL, and the concentration of all control cells was at 10 6 cells/mL for all cell capturing experiments in the buffer. 2.3.4 Tumor Cell Isolation from Whole Blood The single donor human whole blood was obtained from Innovative Research (Novi, MI), with anticoagulant of Ethylenediaminetetraacetic acid (EDTA). Colorectal cancer cells DLD 1 and HCT 116 were then spiked into the blood. The protocol of avidin adsorption and aptamer immobilization was performed as described in the previous

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35 section. For capture study, to avoid damage by trypsin to the proteins on the cell surfaces, non enzymatic cell dissociation reagent (1) (MP Biomedicals, Solon, OH) was used (instead of trypsin) to detach cells from the culture dishes. Immediately before introduci ng into the device, cells were filtered using a 40 m BD Falcon cell strainer (Becton, Dickinson and Company, Franklin Lakes, NJ) and then spiked into whole blood at a predetermined concentration of 10, 100, 1,000, 10,000 cells/mL 2.3.5 Instrument Setup The cell suspension (or whole blood) was introduced into the device by pumping. As shown in Figure 2 8 a Micro4 syringe pump (World Precision Instruments, Sarasota, FL) with a 1 mL syringe was connected to the inlet of the device via polymer tubing and a female luer to barb adapter (IDEX Heal th & Science, Oak Harbor, WA). The outlet of the device was c onnected to a waste collector. For tumor cell isolation from whole blood, no buffer additive (Histopaque 1119) was used to avoid the p roperty change of whol e blood. Instead, we used a tiny magnetic stirring bar inside the 1 mL syringe, with a stir plate beneath the sy ringe, to avoid cell settling. The magnetic stirring bar kept cells in suspension while blood was b eing pumped through the device. The device wa s placed on the stage of an Olympus FV500 IX81 confocal microscope (Olympus America, Melville, NY) for detecting cell s capture d To determine cell concentrations, a set of three images corresponding to the red fluorescent cells, blue fluorescent cells, an d transmission images was acquired at ei ght positions in each channel. As shown in Figure 2 9, i mages were then imported into ImageJ (NIH), and cell counts were obtained using the Analyze Particles function after set ting an appropriate threshold. Cell numb ers were further verified by comparing fluorescent images with transmission images; only those with appropriate cell

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36 morphology in the transm ission images will be counted. Capture efficiency was determined by dividing the number of the target cells capture d by the number of total target cell s introduced into the channel. The purity of cells captured was determined by dividing the target cells captured by the total cells captured. 2.3.6 Cell Viability To determine the viability of cancer cells captured in t he device, we performed two assays on these cells: 1) propidium iodide (PI) and acridine orange (AO) staining (Invitrogen) and 2) MTS assay (CellTiter 96 AQueous Non Radioactive Cell Proliferation Assay, Promega, Madison, WI). PI is a membrane impermeant stain, thus it labels only the dead cells with red fluorescence by penetrating the membranes of dead cells and binding to their DNA. AO is a membrane permeable dye that binds to nucleic acids of all cells, re sulting in green fluorescence. By PI/AO staining nonviable cells and viable cells can be differentiated by their difference in fluoresc ent images under a microscope. We followed the instructions of the manufa cturer to carry out the assay. In brief, 200 L of PI/AO working solution was prepared to conta in 2 M PI and 2 M AO in PBS. After cell capture in the microfluidic device, one channel volume of the PI/AO solution was introduced into the d evice and incubated for 10 min. The cells were then examined under the confocal microscope and fluorescent image s were taken to evaluate the viability of captured cells. MTS assay is a colorimetric method for determining the number of viable cells in proliferation assays. To implement the assay, cells were first released from the device by high flow rate washing an d introducing air inside channel followed by ri nsing with the washing buffer. These cells were then collected and quantified. Typically, 20,000 captured cells in 100 L of fresh cell culture medium were seeded in each well of a 96

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37 well plate, and 3 repeat s were simultaneously carried out. For comparison, cells without going through the capture experiment were seeded at the same concentration into other wells (3 re peats) of the same microplate. under 5% CO 2 for 48 h, 20 L of the MTS assay reagent was added to each well and incubated for another 3 h. A plate reader was used to measure the absorbance at 490 nm to evaluate the cell viability. 2.4 Results and Discussion 2.4.1 Cell Aptamer Binding U sing Flow Cytometry To ascertain if a poly(T) modified aptamer preserves its binding affinity and specificity to its target cells, flow cytometry analysis was carried out. Figure 2 10 A shows the histogram of CEM c ells from the flow cytometer. Compared to cells only, a large shift in the fluorescence signal was observed for those cells conjugated with sgc8 aptamers. The result suggests that poly(T) appended sgc8 aptamers still have specific binding with CEM cells. A random single strand DNA library or TD05 aptamer did not have specific binding with CEM cells, showing a tiny shift in the fluorescence signal compared to cells only. Similarly, Figure 2 10 B shows that poly(T) modified TD05 aptamers bound selectively wit h Ramos cells while sgc8 aptamer or a DNA library did not have specific binding. Figure 2 10 C shows that the comparison of the flow cytometry between the cell aptamer binding in the binding buffer and that in the capturing buffer. As detailed in the previo us section, the capturing buffer was prepared by adding Histopque 1119 to the binding buffer for matching the density of blood. The flow cytometry results indicate that the addition of Histopque 1119 to the binding buffer did not have any adverse effect on the binding of sgc8 aptamer with CEM cells.

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38 Confocal microscopy images also shows the strong binding of CEM cells with fluorescent sgc8 aptamer (Figure 2 11A), while there is no binding between Ramos cells and sgc8 aptamer (Figure 2 11B), demonstrating t he strong and specific binding of aptamers with their target cells. 2.4.2 Isolation of Lymphocytes in Device The performance of the microfluidic device was demonstrated first by sorting leukemia cells: CCRF CEM cells (human acute lymphoblastic leukemia) that function as cells. Biotinylated sgc8 aptamers have specific binding with CCRF CEM cells, and they were immobilized onto the micropillars/microchanne ls. A cell mixt ure containing 1 0 000 CEM cells and 10 6 Ramos cells in 1 mL of the capturing buffer was used as a sample. To differentiate these two types of cells during imaging, CEM and Ramos cells were pre stained with Vybrant DiI (red) and DiD (blue), respectively. Figure 2 12 A shows an image of the cancer cell mixture prior to sorting in the device and it is essent ially all Ramos cells in blue. Figure 2 12 B shows an image of cells captured after processing 1 mL of the cell mixture, and the majority of cells ar e now target CEM cells in red. These images show qualitatively that significant enrichment of the cancer cells was obtained through the microfluidic device. Figure 2 12 C shows a single cancer cell captured on a micropillar in the device. To quantify the enrichme nt factor, we counted c ells before and after sorting. Cell labeling and confocal fluorescence detection enabled the counting of the number of the target cells introduced into the device ( Ti ), the number of the target cells captured ( Tc ), and the number of the control cells captured ( Cc ). The cell capture efficiency ( E ) can be calculated by E = Tc / Ti and the cell purity ( P ) in the captured cells can be calculated by P = Tc /( Tc + Cc ).

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39 As high as 98% capture efficiency was obtained from the cell mixture when the flow rate was at 300 nL/s as discussed in detail below. The high capture efficiency partially arose from the specific binding of sgc8 aptamers with CEM cells, with a dissociation constant of Kd = 0.8 0.09 nM. In addition, a poly T linker at the end o f the biotinylated aptamer sequence should minimize the steric effects of the device surface on the aptamers, The specific and strong binding of sgc8 aptamer s to CEM cells was verified by con f ocal fluorescence microscopy. Figure 2 12 D shows the fluorescent micrograph of TAMRA labeled sgc8 aptamer specifically bound to unstained CEM cells, but these aptamers did not bind to Ramos cells ( Figure 2 11B ). Fluorescence on cell surfaces not only demon strated the binding of the aptamers to the target cells, but also showed a possible way to identify unstained CTCs captured from a sample via a fluorescently labeled aptamer. 2.4.3 Effects of Channel Depth and Flow Rate To optimize the performance of th e microfluidic device, we investigated the effects of the channel depth and flow rate on capture efficiency and cell purity. By changing channel depth (pillar height), the size of micropillar sizes and the interpillar gap were altered simultaneously due to isotropic etching. 85 As a result, the geometry of the micropillar device can be studied by varying the channel depth. Figure 2 13 A shows that the capture efficiency reduced slightly with the increasing channel depth from 24 m to 44 m, whereas the cell purity in Figure 2 13 B improved significantly with the same change in the channel depth. The decreased capture efficiency with the increasing channel depth is likely due to the reduction in the probability of cell encounters with the top a nd bottom surfaces in a deeper channel as well as cells

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40 interaction with aptamers on pillars with wider inter pillar gap. Nevertheless, the results show that our design of the pillar row shift and the curved surfaces of micropillars enabled sufficient inte raction opportunities of cells with the surfaces, essentially maintained the capture efficiency when the channel depth was increased. However, the increased channel depth drastically reduced the non specific binding, particularly geometric trapping of cont rol cells, therefore significantly improvin g the cell purity. Based on the data in Figures 2 13 A and B, we chose a depth of 40 m as the best trade off between the capture efficiency and cell purity. Figures 2 13 C and D show the effects of the flow rate on the capture efficiency and the cell purity. The capture efficiency reduced with the increasing flow rate because of a larger shear force at a higher flow rate and the reduced interaction ti me between cells and surfaces. The cell purity improved with the increasing flow rate due to the fact that non specifically bound cells were washed away with a stronger shear force at a higher flow rate. Based on these results, we chose 600 nL/s as the best compromise between the capture efficiency and cell purity; and this flow rate results in sufficient throughput. Using a device with the optimal channel depth of 40 m and at a flow rate of 600 nL/s, we obtained cell purity of (81 3)% with ca pture efficiency of (95 2)%. These experimental conditions were used f o r all subsequent experiments. To compare this device with the previous efforts using a flat channel device, 75 we plotted both results in Figure 2 14 A. It show s that the capture efficiency of the device in this work is significantly better the flat channel in t he previous work. The results verify the design in which 1) the increased surface area via micropillars enhanced the loading capacity of

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41 aptamers, and 2) the row shift of micropillars and channel geometry significantly increased the probability of cell enc ounters with aptamers on the surfaces. Using the optimal channel geometry and flow rate, we also studied 3 other types of cancer cells in the device as shown in Figure 2 14 B. They were captured in a cell mixture us ing their respective aptamers. To study t he isolation of Ramos cells, an aptamer called TD05 was used as it has specific binding with Ramos cells (with a dissociation constant of Kd = 74.7 8.7 nM 73 ). A cell mixture containing 1 0, 000 Ramos cells and 10 6 CE M cells in 1 mL buffer was used and the captur e efficiency was (93 2)%. and DLD 1 cells (colorectal carcinoma ) were also studie d HCT 116 cells have strong affinity with KCHA10 aptamer (Kd = 21.3 1.7 nM) while DLD 1 cells show specific binding with KDED2a 3 aptamer (Kd = 29.2 6.4 nM). 86 87 Using a cell mixture containing 10,000 of each type of carcinoma cell and 10 6 Ramos cells (as the control), we obtained capture efficiency of (97 3)% for HCT 116 cells and (91 1)% for DLD 1 cells, respectively. 2.4.4 Tumor Cell Isolation from Whole Blood To mimic the isolation of CTCs from patient blood, we spiked colorectal carcinoma cells, HCT 116 cells, into whole blood that was used as received. One mL of unprocessed whole blood, spiked with 100 HCT 116 cells, was introduced into the microf luidic devi ce at a flow rate of 600 nL/s. Using the experimental procedures described in the previous s ection, we obtained capture efficiency of (96 8)% as shown in Figure 2 14 C. A similar experiment using another carcinoma cells, DLD 1 cells, resulted i n capture efficiency of (92 6)% (note that a diff erent aptamer was used).

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42 To illustrate the potential of the device for clinical applications, we evaluated the isolation of HCT 116 cells from whole blood at concentrations of 10,000, 1,000, 100 and 10 cel ls/mL. Capture efficiencies of >95% were achieved in all cases, and a calibration curve between the number of the cells spiked and the number of the cells captured is shown in Figure 2 14 D. Comparable results were obtained for capturing HCT 116 cells from the capturing buffer. The results show that the device has a potential to detect CTCs in clinical samples since the number of CTCs in 1 mL of peripheral blood of cancer patients is often in the range of 1 100. 88 In addition, we addressed the problem of low throughput of a typical microfluidic device by connecting 8 microchannels t hr ough bifurcation (Figure 2 1). The width of each channel is 2 mm. With the optimal flow rate of 600 nL/s, the time required to process 1 mL of whole blood in the device is 28 minutes, which is favorable compared with hours of operation required in the benc hmark instrument. 2.4.5 Cell Viability Isolation and enumeration of tumor cells in peripheral blood of cancer patients is important for medi cal diagnostics and prognosis. However keeping the cells viable during the isolation process is important for subse quent molecular and cellular studies, so that potential therapeutic treatment can be derived after understanding the metastasis mechanisms. 89 The viability of the c ancer cells captured was examined with PI/AO assay and MTS assay as describe d in the Experimental Section. Figure 2 1 5 shows fluorescent microscope image of all cells captured from the mixture of CEM and Ramos cells using PI/AO cell viability assay. The ma jority of cell colors are in green, indicating that the most of the ca ptured cells are still viable. We obtained cell viability of (94 2)% for CEM

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43 cells when they were processed through the device under the optima l flow rate and channel depth. In additio n, we used cell proliferation (MTS assay) as an alternative, and we found cell viability of (93 3)%, which is statistically t he same as in the PI/AO assay. These results indicate that the cells captured using our device are suitable for subsequent cell c ulture and molecular analysis. 2.5 Release of Captured Cells 2.5.1 Motivation After tumor cells captured in the microfluidic device, a primary challenge is to detach the captured cells. For ligand based surface capture methods, cells are firmly captured wi th antibodies or aptamers. Release of the cells of interest is very important for subsequent cell culture and further cellular and molecular analysis. In addition, be cause that the detection of captured cells relies on the time consuming task of scanning t he entirety of a microfluidic device using a microscope to generate statistically valid data 90 e fficient release of captured cells will enable rapid detection method s such as flow cytometry. Several groups have reported the use of trypsin base d, DNase facilitated and temperature mediated methods for cell release. 48 91 93 However, the efficiency of thes e approaches is not satisfactory. In addition, some of them are even detrimental to cells. Therefore, efficient cell release with negligible cell damage is of great significance. Herein, we have developed methods for efficient release of captured cells whi le kept cells viable 2.5.2 Cell Release Using Complementary DNA For ligand based cell capture, DNA aptamers provide many advantages over antibodies for cell release. Here, we developed a release method using complementary DNA sequence s which can competit ively hybridize with the cell bound DNA aptamers.

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44 After hybridization, aptamers were released from the cells leading to cell detachment shown in Figure 2 1 6 R esults indicate that the complementary DNA can release ~6 0 % of captur ed cells on the channel su rface Flow cytometry results (Figure 2 1 7 B ) also show that part of the aptamers were released from cell surface after incubating with complementary sgc8 (c sgc8) for 3 0 min with subsequent washing (with decreased fluorescence intensity). Aptamer usually h as a hairpin structure (with a loop and a stem) after binding with its target proteins on the cell surface, making the hybridization difficult to initiate. To enhance DNA hybridization, a toehold sequence was incorporated to the end of the aptamer and the complementary DNA. The DNA aptamer sequences used for cell capture and release are listed in Table 2 1. As shown in Figure 2 1 7 A, the toehold sequences displaced the base pair in the stem part, promoted the opening of a typical hairpin structure of aptamer thus enhancing the DNA hybridization reaction. Flow cytometry analysis also show ed enhanced DNA hybridization (Figure 2 1 7 B ) with the toehold mediated DNA strand displacement This DNA engineering method increased the release efficiency of the target cel ls from 6 0 % to 82% (Figure 2 1 7 C) Figure 2 18 shows representative cell image s before and after release, demonstrating the effective release of captured CEM cells from the device using toehold mediated DNA hybridization 2.5.3 Alternative Cell Release Met hods Besides the complementary DNA based method, the following are potential methods for aptamer mediated cell release. One method for cell release will involve the synthesis of a disulfide bond on the aptamer. After cell capture, cleavage of the disulfide bond can be induced by a biocompatible reducing agent (sodium 2

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45 sulfanylethanesulfonate, or Glutathione), which cuts the aptamer and releases the cells. Since the reducing agent is a small molecule, it has very low steric effect, allowing much more intera ction between the reducing agent and the apt amers compared with DN ase Another method for cell release will involve us ing a photocleavable (PC) biotin or PC linker conjugated aptamer. The cell release is also feasible for PC biotin conjugated antibodies. P C biotin or PC linker modified aptamer will be used for cell capture and the aptamer can be cleaved upon UV light irradiation. After capture, cells can be released by UV irradiation for ~10 min. With a photomask, this photocontrollable method can also be used for cell micropatterning. 2. 6 Bonding and Surface Functionalization of Cyclic Olefin Copolymer Microchip for Aptamer B ased Cancer Cell Capture 2.6.1 Motivation While silicon and glass have been widely used for lab on a chip applications, polymers hav e been increasingly used in diagnosis because of their inexpensive fabrication. 94 Among them, polydimethylsiloxan e (PDMS) ha s enjoyed popularity because of its fast prototyping using soft lithography, and easy surface modification similar to silicon and glass. 25 However, PDMS is poorly suited to mass production, which limits its disposability. In addition, th e elastomeric PDMS suffers from mechanical softness and gas permeabi lity, which limits its application for many diagnostic purposes. Recently, thermoplastics, 26 most notably, p olymethyl methacrylate ( PMMA ) and cyclic olefin copolymer (COC) have played an increasingly important role for microfluidic and lab on a chip device fabrication because of their low cost, mass production capability and disposability. Thus thermoplastic microfluidic s is invaluable for in vitro diagnosis, such as circul ating tumor cell (CTC) detection

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46 Cyclic olefin copolymers (COCs) or Cyclic ol efin polymers (COPs) are increasingly popular because of their excellent mechanical, therm al chemical, and optical properties. Compared with other thermoplastics, COC device s ha ve excellent optical transparency, high chemical resistance and wide range of glass transition temperature ( T g ), which makes COC ideal for lab on a chip applications 95 However, COC has not been widely used because of its inert surface for chemical modification and biomolecule immobilization. Herein, we developed a facile surface modification method for COC to immobilize biomolecules, such as aptamers. A ntibodies can also be immobilized on COC surface with the si milar protocol. We also developed a pressure free bonding method for fa bricating robust COC device. Then we employed the fabricated COC microchi p for efficient capture of cancer cells using aptamers 2.6.2 COC Device Design and Fabrication The micropillar device design is the same as the device reported in section 2 .2 A glass device was first fabricated using the same protocol reported in section 2.2 Afterwards, the glass device was sent to a vendor (NiCoForm, Inc Rochester, NY ) for fabrication of a metal mold using electroplating. W ith the metal mold (also called E form) a COC device can be easily fabricated using compression molding. 96 Figure 2 19 A shows a picture of the fabricated COC device, with micropillar array shown in the SEM images (Figure 2 19 C &D) 2.6.3 Pressure F ree Bonding Thermop lastic devices were often fabricated by bonding a cover film with a substrate containing microchannels or other microfeatures. 97 Typical bonding techniques includ e thermal fusion, solvent bonding, surface treatment and a dhesives. 98

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47 Most bonding methods involve pressure (e.g., lamination), which leads to collapse of wide channels (e.g., the inlet and outlet area of our device shown i n Figure 2 19A) Here, a COC substrate with microfeatures was bonded with another COC film by spincoating a layer of PDMS membrane (1500 rpm, 30 seconds) on the COC film without applying any pressure Both the COC substrate ( Zeonor 1020R Zeon Chemicals L .P., Louisville, KY ) and PDMS coated COC film (TOPAS 8007 TOPAS Advanced Polymers, Inc. Florence, KY ) we re treated with UV/Ozone before being attached together for the bonding. A scheme of the bonding process is shown in Figure 2 19B. 2.6.4 One Step Surf ace Immobilization of Aptamers on COC T he s urface functionalization of COC is difficult because of the saturated hydrocarbon structure of COC which has only C C and C H bonds. Methods of CO C surface functionalization include direct oxygen plasma treatment, photografting, silanization, etc. Among these methods, silanization provides a robust immobilization of biomolecules by forming covalent bonds, but it usually involves using cross linkers such as 1 ethyl 3 (3 dimethylaminopropyl) carbodiimide (EDC) and N hydroxysuccinimide (NHS), 1,4 Phenylene diisothiocyanate (PDITC), toluene 2,4 diisocyanate (TDI), and succinimidyl 4 (N maleimidomethyl) cyclohexane 1 carboxylate (Sulfo SMCC) for immobilization of biomolecules. 99 The cross li nking step makes the surface modification complicated and less efficient. Here in we developed a method of COC functionalization using 3 Isocyanatopropyl triethoxysilane (IPTES), which avoided the time consuming cross linking. The N=C=O group on IPTES end can directly react with NH 2 group on amin o aptamers without any catalyst, making our surface modification efficient and facile. Figure 2 20 shows the process of surface modification of COC device for immobiliz ing aptamers and captur ing cancer cells. FITC modified amino aptamer was

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48 used to determine the immobilization efficiency with c onfocal laser scanning microscope for the detection. Results show ed that a strong fluorescent signal was detected on IPTES treated COC surface after washing (Figure 2 21A) which indicate d that fl u orescent DNA aptamer s were immobilized on the surface of COC. Control experiments without using IPTES show ed very weak fluo rescence as shown in Figure 2 21B & C 2.6.5 Cancer Cell Capture in COC device For the IPTES treated COC chan nel, amin e modified aptamer was used for immobiliz at ion on the device surface. Then, target cancer cells were introduced into the channel through pumping at a flow rate of 300 n L /s. The micropillar based COC device achieved ~9 2 % capture of CEM cells using amino sgc8 aptamer, and ~90% capture of Ramos cells using amino TD05 aptamer as shown in Figure 2 21D 2. 7 Conclusion In this chapter, a DNA aptamer enabled, micropillar based microfluidic device was demonstrated for the isolation of cancer cells in unpr ocessed peripheral blood. The unique geometry of the micropillar array in the device resulted in the hi gh performance cell isolation. High affinity aptamers were used as an alternative to antibodies for cancer cell isolation. This microfluidic device enabl ed the isolation of as few as 10 tumor cells from 1 mL of untreated whole blood with >95% capture efficiency within 28 minutes. A cell release method was developed and a thermoplastic device was fabricated for cell capture. The advantages of such a device over the benchmark methods include rapid analysis, no pre treatment of blood samples, and low detection limit. As a result, the device has a potential to be used for clinical applications such as cancer diagnosis, prognosis, and monitoring the progress.

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49 Figure 2 1. Layout of the microfluidic device. A) Picture of the device consisting of 1 inlet, 1 outlet, and 8 channels connected through bifurcation. The size of the device is 3 in. 1 in., the same size of a microscope slide. B) Optical micrograph ( 10X) of a portion of m icropillar array in a channel. C) Scanning electron microscope (SEM) image of isotropically etched elliptical micropillars in the glass substrate.

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50 Figure 2 2 COMSOL simulation for the micropillars S treamline and velocity m agnitude of flow across through the micropillar array : A) a t the top of the micropillars; B) a t the center of the micropillars. Since the micropillars were isotropically etched, the pillar size varies from the top to the bottom. Distorted streamlines indic ated enhanced interactions between cells and device surfaces.

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51 Figure 2 3. Comparison of streamline s through different micropillar array s using COMSOL simulation. A) The micropillar array with a shift after every three rows with distorted streamlines f or enhancing cell micropillar interactions ; B) The microp illar array without any shifts limited streamline distortion is observed.

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52 Figure 2 4 Scheme of surface functionalization and cancer cell capture in the device. Avidin is immobilized on the surfa ce of the microchannels/micropillars via physical adsorption, followed by immobilization of biotinylated aptamers through biotin avidin chemistry. Target cancer cells are then captured via the specific interaction between the aptamers and the receptors on cell surfaces. Figure 2 5 Confocal fluorescence images of FITC modified aptamers with different concentrations immobilized on the surfaces of a microfluidic channel. 1 mg/mL of avidin was introduced into the channel and incubated for 1 5 min, followed by three times of washing using the binding buffer. FITC labeled sgc8 poly(T) biotin aptamers were then introduced into the channel, incubated for 1 5 min, and washed three times with the binding buffer. The c oncentrations of aptamers are: A) 2.5 M, B) 25 M, C) 50 M, and D) 100 M.

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53 Figure 2 6 Schem e of surface modification of glass or PDMS substrate with streptavidin using silane based method With streptavidin modified surface, biotinylated aptamer was then immobilized on the surface, followed by cell capture d Figure 2 7. Comparison of avidin physical adsorption based method (Adsorption) and silane based method (Silane) toward the immobilization of aptamers on PDMS and glass surface, respectively.

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54 Figure 2 8 Schem e of cancer cell capture experi ment setup. 1 mL whole blood spiked pumping, CTCs were captured among the aptamer functionalized micropillars. The bottom picture shows the scanning electron microscope (SEM) image of the micropillar array.

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55 Figure 2 9. Counting of cell numbers using ImageJ. A) The intensity threshold, cell size and circularity can be appropriated selected for accurate cell counting B) With display of the overlay out line, each cell count can be confirmed; adjustment can be made for cluster of cells.

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56 Figure 2 10 Flow cytometry histograms showing the selective binding of target cell s with corresponding aptamers. A) CEM cells select ively bind with sgc8 aptamers. B) Ramos cells select ively bind wi th TD05 aptamers. C) Comparison of cell aptamer binding in the binding buffer (BB) and in the capturing buffer (CB).

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57 Figure 2 11 Confocal laser scanning microscop y images showing the selective binding of target CEM cell s with sgc8 aptamers A) Fluorescence image of CEM cells select ively bind with TAMRA labeled sgc8 aptamers (Red fluorescence). B) Ramos cells do not bind with sgc8 aptamers. Left panels are TAMRA fluorescence pseudo co lo red red, and right panels are the overlay of TAMRA fluor escence and the bright field image (scale bar = 20 m)

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58 Figure 2 12 Fluorescence image s of cancer cells captured in device. A) Representative image of low abundant target CEM cells (stained with a red fluorescent dye) among high abundant control Ramos cells (blue) before so rting. B) Representative image of CEM cells (red) among Ramos cells (blue) after sorting (1 mL of the cell mixture was enriched thr ough the microfluidic device). Scale bar = 50 m. C) Image of a CEM cell captured on the wall of an el liptical micropillar in the device (scal e bar = 20 m). D) Microscopy image of unstained CEM cell s bound with fluorescently labeled aptamers, with color only on the surface of target cells (scal e bar = 20 m)

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59 Figure 2 13 Capture efficiency and puri ty as a function of channel depth and flow rate. A) The capture efficiency as a function of the channel depth. B) The purity of cells captured as a function of the channel depth. The flow rate is 6 00 nL/s for both A) and B). C) The capture efficiency as a function of the flow rate. D) The purity of cells captured as a function of the flow rate. The channel depth is 40 m for C) and D). In all experiments, CEM cells were used the target cells and Ram os Cells as the control cells. The error bars represent on e standard deviation of 3 repeat ed experiments.

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60 Figure 2 14 Tumor cell capture from whole blood using the microfluidic device. A) Comparison of the capture efficiency between a flat channel device reported previously and the mic ropillar device in thi s work. B) Capture efficiencies of 4 types of cancer cells in the microflu idic device with micropillars. A different aptamer with specific binding with cells of interest was used for each t ype of cancer cells. C) Capture efficiencies of DLD 1 cells and HCT 116 cells in whole blood. D) Regression analysis of the number of the cells captured by the microfluidic device versus the number of the cell s spiked into 1 mL of samples. HCT 116 cells at different concentrations were spiked either into the capturing buf fer with Ramos cells as the control or into whole blood. Two calibration curves overlap with each other, reflecting no significant difference bet ween buffer and blood samples. The error bars represent one standard deviation of 6 repeats for 10 cell sample s and 3 repeats for other cell numbers.

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61 Figure 2 1 5 Fluorescent microscope image shows the viability of captured cells. Image of CEM cells captured in the microfluidic device after PI/AO staining. The red c olor indicates nonviable cells ( PI staining) w hile the gree n only color indicates viable cells ( AO staining) scale bar = 50 m. Figure 2 1 6 Schem e of cell release using complementary DNA sequence. A) After cells captured inside microchannel complementary DNA was added to the channel. B) Complem entary DNA competitively hybridize d with the aptamer s thus releasing the captured cells.

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62 Figure 2 1 7 Release of captured cells using toehold mediated DNA hybridization. A) Schematic of toehold mediated DNA hybridization by DNA strand displacement T he complementary DNA competitively hybridizes with sgc8 aptamer, the toehold facilitates the opening up of the hairpin structure, thus enhancing the DNA hybridization. B) Flow cytometry assay shows the binding of fluorescent sgc8 with CEM cells, with a h igh fluorescence signal observed (green and cyan histogram). After adding complementary DNA, the aptamer is released by competitive hybridization, and the signal shifts back (blue histogram). W ith the toehold sequence, the signal shifts back further (purpl e histogram). C) Comparison of releasing cells using hybridization alone and toehold mediated hybridization.

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63 Figure 2 1 8 Efficient release of captured cells. A ) Representative microscopy image of the captured cells before release. Target CEM cells (re d) were efficiently captured with a few control Ramos cells (blue) nonspecifically captured. B ) Representative microscopy image of the captured cells after treatment with Toehold based complementary DNA for 30 min and washing; most of the cells were releas ed. Scale bar = 50 m.

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64 Figure 2 1 9 Picture of COC microchip and its bonding process A ) Picture shows the free bonding; B) The bonding process of COC device; C) 100 SEM image of the COC micropillar array; D) 600 SEM image of the COC micropillars.

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65 Figure 2 20 One step surface immobilization of aptamers on COC surface for cell capture UV/ O zone treated COC was first incubated with IPTES, followed by washing, then amino modified aptamer was incubated for 2h for immobilization, finally, target cancer cells were specifically captured on the COC device.

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66 Figure 2 21 Surface coating of amino aptamer s on COC substrate for cell capture A ) With IPTES coating, fluorescent amino aptamers were efficiently immobiliz ed on COC surface; B) Without IPTES coating, few aptamer s w ere immobilized; C) Comparison of the fluorescence intensity for A and B. D) Cell capture efficiency of CEM cells and Ramos cells using NH 2 sgc8 aptamers and NH 2 TD05 aptamers, respectively.

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67 Table 2 1. DNA aptamer sequences, the underlined bases are the toehold sequences Name Sequences sgc8 ( poly T biotin) ATC TAA CTG CTG CGC CGC CGG GAA AAT ACT GTA CGG TTA GAT TTT TTT TTT biotin complementary sgc8 TCT AAC CGT ACA GTA TTT TCC CGG CGG CGC AGC AGT TAG AT toehold sgc8 ( poly T biotin) GAG TGA GGT TTT T AT CTA ACT GCT GCG CCG CCG GGA AAA TAC TGT ACG GTT AGA TTT TTT TTT T biotin complementary toehold sgc8 TCT AAC CGT ACA GTA TTT TCC CGG CGG CG C AGC AGT TAG AT A AAA ACC TCA CTC

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68 CHAPTER 3 MULTIVALENT DNA NANOSPHERES FOR ENHANCED CAPTURE OF CANCER CELLS IN MICROFLUIDIC DEVICES 3.1 Background The isolation of rare cells in peripheral blood such as circulating tumor cells (CTCs) is high ly important but challenging. 13 35 100 CTCs are cancer cells shed from either primary tumors or metastatic sites and are highly related to the initiation of metastasis and the spread of cancer to distant organs T hus CTCs hold the key for understanding metastasis, diagnosing cancer and monitoring treatment response. 10 12 67 However, the extraordinary rarity of CTCs makes their isolation and characterization technically challenging. Traditionally, methods based on flow cytometry have been used in clinics, but with a considerable number of false negatives and low detection sensitivity. 32 101 The only FDA approved CTC enumeration method is CellSearch Assay, which uses a ntibody coated magnetic beads for CTC isolation. However, it suffers from low CTC capture efficiency. 37 10 2 Recently, microfluidic devices with monovalent capture ligands, including antibodies 17 47 51 103 and nucleic acid aptamers, 74 75 78 have been extensively used for immunocapture of rare tumor cells. Ho wever, most efforts for increasing the sensitivity of cell capture are based on engineering complicated structures inside the microfluidic devices, such as microposts, sinusoidal channel s, and silicon nanopillar s etc., for enhancing ligand cell interactio ns. 48 50 72 104 These structures make the device fabricati on time consuming and induce significant nonspecific cell capture, causing low specificity. Multiv alent D NA Nanospheres f or Enhanced Capture o f Cancer Cells i n Microfluidic Devices ACS Nano 2013, 7, 7067

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69 Herein, we have investigated use of nanotechnology based multivalent binding to enhance cell capture in microfluidic devices. Multivalent binding, the simultaneous interaction of multiple ligands on one entity with the complementary receptors on another, has been widely used for achieving high affinity molecular recognition in biological processes. 105 109 The multivalency enhanced binding between the ligands and targets in those biological systems has been extensiv ely investigated. 110 112 To achieve multivalent binding, scaffolds from numerous nanoscale structures, suc h as dendrimers, 113 114 nanorods, 115 nanoparticles, 116 polymers 117 and proteins, have been used by researchers for assembl ing multiple ligands. And dendrimer med iated multivalent binding have been used for enhanced surface capture of cells. 114 Recently, nuclei c acid aptamers have been selected for targeting numerous cancers 60 63 and nanomaterial aptamer conjugat es 118 119 have been extensively used for enhanced molecular recognition but none of them have been used for enhanc ing captur e of cancer cells. 120 124 Here, we hypothesize the nanoparticle aptamer conjugates could greatly improve the efficiency of capturing cancer cells. We chose gold nanoparticles (AuNPs) as the multivalent ligand scaffolds to assemble multiple DNA aptamers (DNA nanospheres) owing to their easy synthesis and conjugation with DNA. 125 126 Our flow cytometri c analysis demonstrated the multivalent binding between aptamers and cells through AuNP conjugation. Then we developed a flat channel microfluidic device which is able to capture cancer cells from buffer or lysed blood with high efficiency and high through put using the AuNP aptamer conjugates (AuNP aptamer). The enhanced binding affinity afforded by the AuNP aptamer modified surface significantly increased the capture efficiency of target cancer cells. And the AuNP aptamer maintained high

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70 capture efficiency with increased flow rate, which considerably improves the sample throughput of the microfluidic device. 127 The scheme of the AuNP aptamer mediated cell capture is shown in Figure 3 1. The microfluidic device surface is first coated with avidin by physical adsorption. 75 78 Then, biotinylated aptamer conjugated AuNPs are immobilized onto the channel through biotin avidin interaction. When a sample containing t arget cancer cells passes through the channel, cells are captured via the specific interaction between the aptamers and the target cell receptors. Since each AuNP is conjugated with ~95 aptamers, we hypothesize that the AuNP aptamer b inds to cell surface markers in a cooperative manner, leading to multivalent effect and resulting in enhanced cell capture efficiency Besides the multivalent binding, t he AuNP aptamer modified surface increases the surface roughness 104 and allows enhanced local topographic interactions betwe en the AuNP aptamer s and nanoscale receptors on the cell surface, 50 128 129 contributing to the increased cell capture 3.2 Methods 3.2.1 Synthesis and Characterization of Gold Nanoparticle A ptamer Conjugates Hydrogen tetrachloroaurate (III) (HAuCl4), t risod ium citrate dihydrate, t ris (2 carboxyethyl) phosphine hydrochloride (TCEP), t ris (hydroxymethyl) aminomethane (Tris) and sodium acetate were obtained from Sigma Aldrich (St. Louis, MO). Acetate buffer (500 mM, pH 5.2) was prepared using a mixture of sodi um acetate and acetic acid. Tris acetate buffer (500 mM, pH 8.2) was prepared using Tris and acetic acid. AuNPs were prepared using the protocols reported previously. 130 Briefly, 100 mL of 1 mM HAuCl 4 solution was heated till reflux. Then, 10 mL of 38.8 mM sodium citrate was added and reflux w as continued for another 20 min. The diameter of such prepared

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71 AuNPs was ~13 nm, measured by transmi ssion electron microscop y (TEM). The concentration of the AuNPs was ~13 nM, determined by UV Vis measurement at 520 nm using a Cary Bio 300 UV spectrometer (Varian) (Figure 3 2A ) DNA aptamers were synthesized in house. Thiol modified s gc8 a ptamer sequence was: 5 thiol PEG ATC TAA CTG CTG CGC CGC CGG GAA AAT ACT GTA CGG TTA GA biotin 3 The sequences of all aptamers used are listed in Table 3 1. For flow cytometric analysis, a fluorescein isothiocyanate (FITC) modifier was used to replace the biotin linker. All DNA aptamers were purified using a ProStar HPLC (Varian, Walnut Creek, CA) with a C18 column (Econosil, 5U, 250 4.6 mm) from Alltech Associates ( Deerfield, IL), with t riethylammonium acetate a cetonitrile as eluent. DNA concentration was determined by UV Vis measurement at 260 nm. Thiol modified aptamers were conjugated on AuNPs using the reported protocols. 131 130 132 Aptamers (9 L, 1 mM) were added with acetate buffer (1 L, 500 mM) and TCEP (1.5 L, 10 mM) and incubated for 1 h at room temperature to activate the thiol group. Then the TCEP treated aptamer was added to 3 mL of as prepared AuNPs and incubated for 16 h. Finally, Tris acetate buffer (30 L, 500 mM) and NaCl (300 L, 1M) were added and the mixt ure was incubated for 24 h. Unconjugated aptamers was then removed by centrifugation at 14,000 rpm for 15 min. The aptamer concentration in the supernatant was measured, and the final conjugated aptamer concentration in the AuNPs was determined by subtrac ting the supernatant concentration from the previous aptamer concentration. The final AuNP concentration was 12.7 nM with an aptamer concentration was 1.2 M giving an average of approximately 95 aptamers on each AuNP. Dynamic light scattering (DLS)

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72 measu rement was performed to evaluate the hydrodynamic diameter of the AuNPs before and after conjugation with aptamers using Zetasizer Nano ZS, (Malvern, Worcestershire, United Kingdom) (Figure 3 3 ). Zeta potential measurements were performed using the same in strument. Fluorescence spectroscopy (Figure 3 2B ) also demonstrated the successful conjugation of aptamer on the AuNP. T he fluorescence signal of each AuNP aptamer conjugate is much higher than that of individual aptamer. 3.2.2 Device Design and Fabricati on A single flat channel device was initially used for proof of concept studies, and then eight flat channels were parallelized to form a high throughput device. As shown in Figure 3 4 A the single flat channel device was designed with a length of 50 mm, w idth of 2 mm, depth of 100 m, and with single inlet and outlet. Three independent devices can be incorporated within one microscope slide size (3 in. 1 in.). To increase the throughput, eight channels were connected through parallelization, and uniform flow was maintained in the eight channels. The size of the high throughput device is also 3 in. 1 in., as shown in Figure 3 4 B Both of the two devices were made of polydimethylsiloxane (PDMS), and bonded to a 3 in. 1 in. glass slide. PDMS devices wer e fabricated according to the procedures reported by 30 T he layout of the device was designed in AutoCAD and then sent to CAD/Art Services, Inc. (Bandon, OR) to produce a high resolution transparency photomask. Silicon wafers (Silicon Inc., Boise, ID) were first spin coated with SU 8 2035 photoresist (MicroChem, Newton, MA) using a spin coater (Laurell Tech., North Wales, PA). Then the pattern on the photomask was transferred to the silicon substrate via UV exposure. After development, a silicon master patterned with the complementary structures was obtained. PDMS devices were fabricated by casting a liquid PDMS

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73 precursor against the master using Sylgard 184 reagents (Dow Corning, Midland, MI) according to the instructions of the manufacturer. To prevent the cured PDMS from sticking to the silicon master, TFOCS (Tridecafluoro 1,1,2,2 tetrahydrooctyl 1 trichlorosilane) (Sigma Aldrich) was vacuum vaporized to the surface of the master. The channel depth which was controlled by the spin speed of the SU 8 was measured using a Dektak 150 pr ofilometer. The PDMS substrate was then sealed with a glass microscope slide, and inlet and outlet wells were created at the channel ends by punching holes in the PDMS sheet. The design of herringbone mixer device was inspired by several works in the liter ature, 51 133 and the dimensions were chosen for optimal cell capture, as shown in Figure 3 9 A The mixer d evice was fabricated as described above but using a two layer SU 8 fabrication technique, with two coating and exposure steps and a single developing step. 134 The silicon mold consists of a first layer as the main channel and the second layer as the herringbone ridges, which become grooves after tr ansfer to the PDMS substrate. As shown in Figure 3 5, enhanced mixing occurs with the herringbone groove structure inside the PDMS microchannel, which generates transverse flow and microvortex. 3.2.3 Cell Lines and Buffers T cell human acute lymphoblastic leukemia cells ( CCRF CEM cells CCL 119) and B cell cells ( Ramos cells CRL 1596 ) were purchased from American Type Culture Collection (ATCC). CEM and Ramos cells were cultured in RPMI medium 1640 (ATCC) supplemented with 10% feta l bovine serum (FBS; heat inactivated; GIBCO) and 100 units/mL penicillin streptomycin (Cellgro, Manassas, VA). Both cultures were incubated at 37C under 5% CO 2

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74 phosphate buffered saline w ith c alcium and m agnesium (PBS) (Fisher Scie ntific, Hampton, NH) was used to wash cells. A solution of 50 mg/mL (5%) bovine serum albumin (BSA) (Fisher) and 0. 1 % Tween 20 (Fisher ) in PBS was used for rinsing the unbound molecules on the surface and resuspending cells for the cell capture. BSA and T ween 20 in PBS can fully passivate the surfaces to reduce nonspecific adsorption of cells in the channel. 3.2.4 Flow Cytometric Analysis Flow cytometry was used to evaluate the targeting capabilities of AuNP aptamer conjugatess toward specific cells. Fluor escence measurements were made with a FACScan cytometer (BD Immunocytometry Systems, San Jose, CA). Briefly, 200 000 cells were incubated with FITC labeled free aptamer or AuNP aptamer conjugates in of PBS (containing 0.1% BSA) for 30 min on ice. Af ter incubation, the cells were PBS and 10 000 counts were measured in the flow cytometer to determine the fluorescence. Varying concentrations of free sgc8 and AuNP sgc8 aptamers were used to determine thei r binding affinities. The FITC labeled random DNA library was used as a negative control to determine nonspecific binding. All of the experiments for the binding assay were repeated three times. The mean fluorescence intensity of target cells labeled by ap tamers was used to calculate the specific binding by subtracting the mean fluorescence intensity of nonspecific binding from random DNA library. 63 The equilibrium dissociation constants ( K d ) of the aptamer cell interaction were obtained by fitting the dependence of fluorescence intensity of specific binding on the concentration of the aptamers to the equation Y = B max X /( K d + X ) using SigmaPlot (Jandel, San Rafael, CA), where Y is the fluorescence intensity and X is the concentration of aptamers.

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75 3.2.5 Cell Capture Assay in Microfluidic Device s Immediately before cell capture experiments, cells were washed with PBS and resuspended at 10 6 cells/mL. By following the ma CEM and Ramos cells were stained with Vybrant DiI (red) and DiD (blue) cell labeling solutions (Invitrogen, Carlsbad, CA) respectively then washed with PBS and resuspended at 10 7 cells/mL in the PBS containing BSA and Tween 20 Labeled cells were stored on ice and further diluted to the desired concentrations before cell capture The single donor human whole blood was obtained from Innovative Research (Novi, MI), with anticoagulant of e thylenediaminetetraacetic acid (EDTA). Ly sed blood was obtained by treating whole blood with r ed blood cell lysing buffer (Sigma Aldrich ) (containing NH 4 Cl) according to manufacturer s instructions. Different concentrations of CEM cells were then spiked in whole blood or lysed blood. To start ce ll capture experiments, one device volume (~100 L ) of 1 mg/mL avidin (Invitrogen) in PBS was first introduced into the device, followed by incubation for 1 5 min and then three rinses with PBS Then, 100 L of sgc8 aptamer or AuNP sgc8 aptamer was introduc ed into the device and incubated for 1 5 min, followed by three rinses with the PBS containing BSA and Tween 20 Finally, 1 mL of cell mixture or blood sample spiked with cancer cells was continuously pumped into the device at a flow rate of 1.2 L/s (or ot her flow rates specified in the text). For cell capture using antibody, anti PTK7 biotin ( Miltenyi Biotec Auburn, CA) was used instead of sgc8 or AuNP sgc8 aptamer. Afterwards the device was washed three times with PBS to remove nonspecifically captured cells followed by acquiring fluorescent images to determine the cell number s. To test the purity of captured cells from lysed blood or whole blood, DAPI ( Invitrogen ) was introduced into the device to label the

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76 nonspecifically captured white blood cells. B y following the manufacturer s instructions, 300 nM DAPI was incubated with cells for 10 min, followed by rinsing with PBS. The cell suspension s or blood samples were introduced into the device by pumping. 78 A Mi cro4 syringe pump (World Precision Instruments, Sarasota, FL) with a 1 mL syringe was connected to the inlet of the device via FEP ( Fluorinated ethylene propylene ) tubing and a female luer to barb adapter (IDEX Health & Science, Oak Harbor, WA). To avoid c ell settling, a tiny magnetic stirring bar was placed inside the 1 mL syringe, with a stir plate beneath the syringe. The magnetic stirring bar kept cells in suspension while cell mixture or blood was being pumped through the device. The device was placed on the stage of an Olympus IX71 fluorescence microscope (Olympus America, Melville, NY) for detecting capture d cell s To determine cell numbers set s of images corresponding to the red fluorescent cells, blue fluorescent cells, and transmission images w ere acquired at different positions in each channel. Images were then imported into ImageJ (NIH), and cell counts were obtained using the Analyze Particles function after setting an appropriate threshold. Cell count s were further confirmed by comparing fluore scent images with transmission images; only those with appropriate cell morphology in the tr ansmission images were counted. 3.3 Results a nd Discussion 3.3.1 Synthesis and Characterization of AuNP A ptamer Conjugates AuNPs were prepared following the methods detailed in the experimental section. Figure 3 1 C shows the transmission electron microscopy (TEM) image of the AuNPs, with an average diameter of 13.6 nm. The as prepared AuNPs were then functionalized with thiol modified DNA aptamers, and the TEM image is shown in Figure 3 1 D with average size of 13.7 nm. A 24 unit p olyethylene glycol (PEG) spacer

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77 between AuNP surface and aptamers was added to minimize the steric effects of the particle surface on aptamers and to increase the loading of DNA on AuNPs 131 Figure 3 1 C & D show that the properties of AuNPs remained unc hanged after conjugation with aptamers, without any aggregation. Dynamic light scattering (DLS) measurements showed that the hydrodynamic diameter of AuNPs was 17.4 nm. After conjugation with aptamers, the hydrodynamic diameter increased to 61.8 nm, demons trating the successful conjugation of aptamers onto AuNPs (Figure 3 3 ). Zeta potential measurements indicated that the AuNP s ha d a zeta potential of 12.5 mV. After modification with aptamers, the zeta potential became 23.2 mV, which is attributed to the negative charges carried by DNA aptamers The comparison of properties between AuNPs and AuNP aptamers is made in Figure 3 1 E 3.3.2 Flow Cytometric Analysis Demonstrating High Affinity Binding To investigate the AuNP aptamer mediated multivalent binding, we directly measured the binding behaviors of AuNP sgc8 aptamer conjugates (AuNP sgc8) and free sgc8 aptamer (sgc8) using flow cytometry. Sgc8 is an aptamer that has specific binding with CEM cells (human acute lymphoblastic leukemia) with a nanomolar (n M) dissociation constant (K d ). 63 Ramos cells (human Burkitt s lymphoma) that do not bind with sgc8 aptamer were used as control cells here. Figure 3 6 A shows a no ticeable increase in fluorescence signal for both AuNP sgc8 and free sgc8 aptamer compared to the random DNA library (Lib) and AuNP Lib proving that both have strong binding with their target cells. Besides, AuNP sgc8 produces a higher fluorescence signal than free sgc8, even with 10 times lower concentration. As shown in Figure 3 6 B n either free sgc8 nor AuNP sgc8 shows a signal increase when incubated with control Ramos cells, demonstrating the specificity of both free aptamers and AuNP aptamers Furthe rmore,

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78 the binding affinity of sgc8 and AuNP sgc8 to CEM cells was measured quantitatively by studying the ir binding with varying concentrations of sgc8 and AuNP sgc8 aptamers As demonstrated in Figure 3 6 C & D AuNP sgc8 show s a 39 times higher binding a ffinity (K d = 0.10 0.02 nM) than that of free sgc8 (K d = 3.9 0. 5 nM). T he lower dissociation constant of AuNP sgc8 suggests a multivalent mediated enhancement in binding affinity when multiple aptamers on the AuNP surface bind to multiple receptors on the cell membrane. Note that the concentration of AuNP is used instead of the concentration of aptamer when measuring the dissociation constant of AuNP aptamer. 3.3.3 Enhanced Cancer Cell Capture in a Flat Channel Microdevice To study the cancer cell captu re using AuNP aptamer we first developed a microfluidic laminar flow device with flat channels (Figure 3 4B ), which allow ed us to directly compare the capture performance between AuNP aptamer and aptamer alone. After coating surfaces with AuNP sgc8 aptame r, a cell mixture containing 10 5 target CEM cells and 10 6 control Ramos cells (1:10 ratio) in 1 mL of phosphate buffered saline (PBS) w as introduced into the channel. Note that the cell solution was continuously pumped into the device without any interrupt ion. CEM and Ramos cells were pre stained with Vybrant DiI (red) and DiD (blue), respectively. Figure 3 7 A shows a representative image of cells captured using AuNP aptamer, a high percentage of target CEM cells (red) was captured, while most control Ramos cells (blue) were washed away In another set of experiments with the same conditions, sgc8 alone was used instead of AuNP sgc8. Figure 3 7 B shows a typical image of cells captured after washing using aptamer alone (without the nanoparticle conjugation). The results in Figure 3 7 A & B clearly indicate that much more target CEM cells were captured using AuNP aptamer than with aptamer alone, demonstrating that enhanced cell capture was

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79 achieved by the AuNP conjugat ion T he capture efficiency using AuNP aptam er and aptamer alone was also studied at different flow rate conditions (with different shear stresses). W e found that AuNP aptamer exhibited more enhancement in the capture efficiency at higher flow rates, as shown in Figure 3 7 C At a flow rate of 1.2 L /s, AuNP aptamer maintain ed a capture efficiency of (9 2 4 )%, while aptamer alone yield ed a capture efficiency of only ( 49 6 )%. The capture efficiency was defined as the ratio of the number of the target cells captured to the number of the target cells initially seeded. The AuNP aptamer enables significant increase in capture efficiency for the target cells. We also studied the purity of the captured cells and found that the capture purity is not affected by the AuNP conjugation. The purity was defined a s the ratio of the number of the target cells captured to the number of total cells captured. As shown in Figure 3 7 D similar purity was obtained for AuNP aptamer and aptamer alone when the same flow rate wa s used; this suggest s that AuNP aptamer does not introduce more nonspecific binding relative to aptamer alone, which is consistent with flow cytometry results on Figure 3 6 B However, the AuNP aptamer allows us to use higher flow rate s to maintain the capture efficiency, higher purity can thus be obtain ed because nonspecifically adsorbed cells are more easily washed away with a stronger shear force at a higher flow rate. 78 In addition to th e DNA nanosphere mediated multivalent binding the enhanced cell capture also accrues from the nano sphere modified surface. The increas es in the surface roughness and total surface area compared with plain surface, allowed enhanced local topographic interactions between the aptamer coated nano particle and nanoscale components on the cell surface. 128 Moreover, the nanoparticle surfaces

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80 packed the aptamers in a highly dense manner, accommodating more aptamers to be immobilized than a plain surface, which is an additional advantage of using AuNP aptamer. T he increased ligand density also contribute s to the enhanced interaction between cells and aptamers. Furthermore, t he enhanced binding strength afforded by the multivalency effect lower s the detachment ratio of immobilized cells, thus increasing the capture efficiency compared to aptamer alone To evaluate the versatility of our system, we also applied the system for capturing Ramos cells using AuNP TD05 aptamer conjugates. TD05 is an aptamer with specific binding to Ramos cell s. 73 A capture efficiency of 90% was obtained with AuNP TD05, while TD05 aptamer alone yield ed only 41% capture, showing significant enhancement in capture efficiency as a result of using DNA nanosphere. The reduced capture efficiency at higher flow rates (shown in Figure 3 7 C ) is due to increased flow induced shear stress and the decreased interaction time between cells and aptamers on surfaces. We further characterized the distribution of captured cells at different locations of the 50 mm long microchannel with different flow rates. As shown in Figure 3 8 A at flow rate of 1.2 L/s (with a shear stress of 0.4 dyn/cm 2 ), 65% of the cells were captured in the first 25% of the channel coated with AuNP aptamer. W ith an increased flow rate of 2.4 L/s (Figure 3 8 B ), the cells captured were dis tributed along the channel because cells needed longer flow distance (travel length ) to have an opportunity to interact with aptamers coated on the surfaces, and the attached cells experience d proportionally increased shear stresses. T he cell surface inter action is due to the ligand receptor binding as well as gravitational force. W ith the AuNP conjugation, the PEG spacer extends the aptamer strands into the 3D space of flow, increasing the

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81 accessibility and frequency of interactions between aptamers and ce lls to permit more efficient cell capture under higher flow rates. To explore the clinical utility of the system, we assessed the isolation of CEM cells from lysed blood (blood with red blood cells lysed) at concentrations ranging from 10 5 to 100 cells/mL As shown in Figure 3 8 C as few as 100 cells were efficiently isolated from 1 mL of lysed blood within 14 min. However, when we tried to capture cancer cells from unprocessed whole blood directly, the capture efficiency wa s significantly low er (even at a low flow rate), as shown in Figure 3 8 D T he relatively low capture wa s primarily due to the reduced interaction chances between target cells and AuNP aptamer, which is caused by abundant red blood cell blockage. 3.3.4 Efficient Isolation of Cancer Cells from Whole Blood U sing DNA Nanospheres in Micromixer Devices Although the laminar flow flat channel device achieved high efficiency when capturing cells in PBS or lysed blood, it showed a low capture efficiency ( < 60%) when capturing cells from whole bloo d. To enable the efficient capture of CTCs from whole blood, we integrated the AuNP aptamer system into a herringbone groove based micromixer device ( Figure 3 9 A ). T he staggered herringbone mixer generate s microvortex and chaotic mixing inside the microcha nnel, which significantly enhance s the cell surface interactions leading to higher capture efficiency 51 72 133 We first evaluated the isolation of 10 4 CEM cells (pre stained by DiI, red) spiked in 1 mL of whole blood at a flow rate of 1 L/s. After cell capture and rinsing, 4,6 diamidino 2 phenylindole (DAP I) was introduced into the device to test the purity of the target cells. DAPI stained all the cancer cell s and leukocytes with blue color and verif ied that captured cells retain intact nuclei. As shown in Figure 3 9 B cells positive to both DAPI

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82 and DiI w e re target CEM cells (blue merged with red), while cells positive to DAPI only were white blood cells (blue only) A purity of (70 6)% was obtained when capturing CEM cells from whole blood, with a capture efficiency of (9 5 3 ) %. T his capture purity fro m whole blood is much higher than those reported in literature (~50% & 14%). 47 51 Further, we tested the ca pture efficiency over a wide range of flow rates from 0.5 L/s to 3 L/s. Control experiments using identical device and conditions with aptamer alone (no AuNP conjugation) were then conducted. Much higher capture efficiencies w ere obtained using AuNP apta mer than aptamer alone, especially at high flow rate s (Figure 3 9 C ) The combined effect of high affinity binding from AuNP aptamer with the passive mixing provided by the herringbone structures enabled high capture efficiency from whole blood (93%) at hig h flow rate (1.5 L/s) To compare the AuNP aptamer based cell capture with traditional antibody based cell capture, anti protein tyrosine kinase 7 (PTK7) antibody was used for capturing CEM cells with identical device and conditions. For the binding betwe en CEM cells and sgc8 aptamer, our previous study identified PTK7 as the marker for CEM cells. 135 As shown in Figure 3 9 C the capture efficiency of CEM cells using anti PTK7 is comparable with aptamer alone, but significantly less than AuNP aptamer. T o test the limit of detection for the AuNP aptamer based cell capture system cell spike number s from 10 5 to 100 w ere explored, and >90% capture efficiency were obtained for all cases at the flow rate of 1.5 L/s Regardless of whether the red blood cells are intact or lysed, high capture efficiency is always obtained by the integration of AuNP aptamer with a herring bone mixer (Figure 3 9 D ) In addition, with the flow rate of 1.5 L/s (5.4 mL/h), 1 mL of blood sample can be processed in 11 minutes, which gives sufficient throughput for clinical applications. T he system gives

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83 more benefit at higher flow rates, maintain s a target cell capture efficiency of >75% for all flow rates up to 3 L/s With this flow rate, only 42 min is needed for processing 7.5 mL blood, the amount of blood needed to detect clinical relevant CTC number. Compared with reported work, t his AuNP ap tamer modified mixer device enables >90% capture at a flow rate 5.4 mL/h, 2 to 4 fold higher than reported aptamer alone based micropillar device (2. 2 mL/h) 78 and antibody coated herringbone device (1.2 mL/h). 51 The results show that the AuNP aptamer modified herringbone device has a great potential for clinical CTC isolation and enumeration 3.4 Conclusion In this chapter we demonstrated the use of gold nanoparticles as an efficient high affinity vehicle for molecular assembly of aptamers for target cancer cell capture in microfluidic devices. Up to 95 aptamers were attached onto each AuNP, resulting in enhanced aptamer mo lecular recognition capability. Flow cytometry results demonstrated the high affinity binding effect using AuNP aptamer conjugates. The capture efficiency for target cancer cells was significantly increased using the AuNP aptamer conjugates because of the cooperative, multiple ligand receptor interactions as well as the increased surface roughness and ligand density W ith the AuNP aptamer surface immobilization a flat channel microfluidic device wa s able to capture 100 cancer cells from 1 mL of lysed bloo d with ~90% capture efficiency within 14 min (4.3 mL blood/h) Using the integration of the AuNP aptamer with a herringbone mixer design, efficient capture of rare cancer cells from whole blood was achieved, with a throughput of processing 1 mL of blood in 11 min The high efficiency, throughput and purity make the system suitable for clinical isolation of CTCs from patient blood

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84 The use of leukemia cell targeting aptamers allows the platform to be suitable for minimal residual disease (MRD) detection. MR D is the small amount of leukemia cells remaining in patient blood during or after treatment when the patient is at remission, which is the major cause for cancer relapse. 136 137 Our system capable of efficient isolation of rare cells is suitable for sensitive detection of MRD, which will be promising for monitoring treatment response and predicting cancer re lapse. H owever, aptamers are currently not as widely used as antibodies, and limited numbers of aptamers have been developed for targeting CTCs in patient bloods. Our future efforts will include incorporating AuNPs with CTC marker binding aptamers [e.g., a nti EpCAM aptamer ( epithelial cell adhesion molecule ) 138 anti PSMA aptamer ( prostate s pecific membra ne antige n )] 139 for capturing CTCs from cancer patients, as well as exploring release and culture of ca ptured CTCs. Spherical DNA nanostructures have been well developed and widely used for cancer cell detection; however, to our knowledge, this is the first use of aptamer nanospheres for enhanc ing cancer cell capture Our results show that the combination o f nanotechnology with a microfluidic device 140 has a great potential for sensitive isolation of cancer cells from patient blood, and is promising for cancer diagnosis and monitoring treatment response.

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85 Figure 3 1 Illustra tion of enhanced cell capture using AuNP aptamer modified surface. A) With AuNP conjugation multiple aptamers on the AuNP surfaces bind with multiple receptors on the cell membrane, leading to cooper ative, multivalent interactions. B) Without AuNP, aptame r alone binds with receptors via monovalent interaction, with much less interactions C ) Transmission electron microscopy (TEM) image of AuNPs. D ) TEM image of AuNPs conjugated with aptamers, scale bar = 100 nm. E ) Comparison between AuNP and AuNP aptamer in terms of particle diameters from TEM images, hydrodynamic diameters from dynamic light scattering (DLS) measurements, and zeta potential measurements.

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86 Figure 3 2. Adsorption and fluorescence spectrum of AuNPs and AuNP aptamer conjugates. A ) Adsorpt ion spectrum of AuNPs, ( max = 5 20 nm ), using a molar absorptivity of 2.7 10 8 L mol 1 cm 1 the concentration of the AuNP is ~13 nM. B) Fluorescence spectrum of fluorescein labeled aptamers at (a) 10 nM and (b) 1 M ; (c) the fluorescence of AuNP aptamer con jugates at 10 nM. Around 95 fluorescein labeled aptamers were conjugated to each AuNP. Thus, the fluorescence signal of each AuNP aptamer is much higher than individual aptamer, as shown when comparing (a) and (c).

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87 Figure 3 3 Dynamic l ight s cattering (DLS) analysis of DNA nanospheres. A) DLS of AuNP s. B ) DLS of AuNP sgc8 aptamer conjugates. T he hydrodynamic diameter of AuNP increased from 17.4 nm to 61.8 nm after conjugation with aptamers. A B

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88 Figure 3 4 Pictures of the flat channel microd evice s A ) T he single flat channel device ; B ) T he parallelized flat channel device with 8 channels connected. The size for both devices is 3 in. 1 in. T he single fat channel device was used for proof of concept studies; data from this device is not shown All the data presented in this chapter are from eight channel devices.

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89 Figure 3 5 Comparison between a flat channel device and a herringbone groove device for flow mixing A ) With a flat channel device, the flow is laminar, with minimal mixing betw een the green dye and red dye, and mixing is only caused by diffusion; B ) With a herringbone groove structures inside the microchannel, transverse flow and chaotic mixing occurred, with enhanced mixing between the red dye and green dye

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90 Figure 3 6 Flow cytometry show s the strong and specific binding of AuNP sgc8 aptamer conjugates with target CEM cells. A ) CEM cells selectively bind with free sgc8 and AuNP sgc8 aptamers; negligible signal change was observed for cells incubated with random DNA library (L ib) or AuNP Lib conjugates (NP lib) compared with cells only. B ) Control R amos cells d id not bind with either AuNP sgc8 or sgc8 alone (with no signal shift for either case), demonstrating the specificity of free sgc8 and AuNP sgc8 aptamers to CEM cells. C D ) Flow cytometry analysis determine s the binding affinity of AuNP sgc8 ( C ) and sgc8 alone ( D ) to CEM cells.

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91 Figure 3 7 Comparison of AuNP aptamer and aptamer alone based CEM cell capture in a flat channel device A B ) Representative image of the t arget CEM cells (red) and control Ramos cells (blue) captured in the flat channel device using ( A ) AuNP sgc8 aptamer conjugates; ( B ) sgc8 aptamer alone. Cell suspensions were continuously pumped into device without interruption. Scale bar = 50 m. C ) Compa rison of CEM cell capture efficiency in PBS between AuNP aptamer and aptamer alone when they were coated in a flat channel device, at flow rate s from 0.4 L/s to 2.4 L/s D ) Comparison of the capture purity of target CEM cells between AuNP aptamer and apt amer alone at the same flow rate ; no statistical difference was observed. Error bars represent standard deviations (n=3).

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92 Figure 3 8 Capture of CEM cells from blood using DNA nanospheres in the flat channel device. A B ) Spatial distribution of surfac e captured CEM cells along the 50 mm long microchannel in the flat channel device at different flow rate s of ( A ) 1.2 L/s and ( B ) 2.4 L/s C ) Capture efficiency for 100,000, 10,000, 1000 and 100 CEM cells spiked in 1 mL of lysed blood, with flow rate of 1 .2 L/s D ) CEM cell capture efficiency from lysed blood or whole blood at the same flow rate (1.2 L/s); 1000 CEM cells were spiked in 1 mL lysed blood or whole blood. Error bars represent the standard deviations of triplicate experiments

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93 Figure 3 9 I solation of cancer cells from whole blood using DNA nanospheres in micromixer device. A ) Device layout and dimensions of a microfluidic device containing herringbone mixer s B ) Representative image of captured CEM cells (DiI+, DAPI+) from whole blood ; the DAPI+ cells (blue only) are nonspecifically captured white blood cells ; scale bar = 50 m C ) CEM cell capture efficiency in whole blood at various flow rates using AuNP aptamer, aptamer alone and anti PTK7 antibody, respectively. D ) C alibration plot of ca ncer cell capture from whole blood and lysed blood with different cell concentrations at 1 .5 L/s, solid lines represent linear fitting. Error bars represent standard deviations (n=3).

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94 Table 3 1. Detailed aptamer sequence information. Underscore ind icates the full sequence of sgc8 aptamer or TD05 aptamer; for flow cytometric test, fluorescein isothiocyanate (FITC) is used instead of biotin linker. Name Sequence sgc8 ATC TA A CTG CTG CGC CGC CGG GAA AAT ACT GTA CGG T TA GA T TTT TTT TTT biotin Thiol sgc8 thiol (PEG) 24 ATC TA A CTG CTG CGC CGC CGG GAA AAT ACT GTA CGG T TA GA biotin TD05 AAC ACC GTG GAG GAT AGT TCG GTG GCT GTT CAG GGT CTC CTC CCG GTG TTT TTT TTT T biotin 3 Thiol TD05 thiol (PEG) 24 AAC ACC GTG GAG GAT AGT TCG GTG GCT GTT CAG GGT CTC CTC CCG GTG biotin 3

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95 CHAPTER 4 A MICROFLUIDIC GEOMETRICALLY ENHANCED MIXING CHIP FOR CAPTURE, RELEASE AND CULTURE OF CIRCULATING TUMOR CELLS FROM PANCREATIC CANCER PATIENTS 4.1 Background Pancreatic cancer is the f ourth leading cause of cancer deaths in the United States, with the poorest 5 year survival rate (6%) for all cancer stages. Over 90% of pancreatic cancers progress to become metastatic. 2 141 The poor prognosis of pancreatic cancer patients is related to the early dissemination of the disease and the lack of early detection. 142 Circulating tumor cells (CTCs) are tumor cells disseminated from primary tumors which subsequently travel through the blood circulation to distant organs. CTCs are thus responsible for the initiation of metastasis and the in tr ansit spread of cancer to distant sites. 67 143 Therefore, CTCs hold the key to track metastasis, and they can be used for cancer diagnosis and monitoring of cancer status. Clinical studies have demonstrated that CTCs are correlated with disease progression for a wide range of cancers, such as breast, colorectal and prostate cancer. 12 144 While biopsy is the current gold standard of cancer diagnosis, it involves removal of tissues or cells from the body and examination by experienced surgeons and pathologists. 145 The invasive nature of biopsy prevents patients from being tested in an ongoing or repetitive basis. CTC examination, on the other hand, is m uch less invasive, with only 5 10 mL of patient blood needed; it is like a blood test for cancer. CTC monitoring is regarded as 146 which enables noninvasive cancer diagnosis and real time monitoring of therapeutic response. Capture, Release a nd Culture o f Circulating Tumor Cells from Pancreatic Cancer Patients Using an Enhanced Mixing Chip Lab on a Chip, 201 4 14, 89 98

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96 However, CTCs are extraordinarily rare, with only a few CTCs circulating amidst billions of blood cells, making their isolation and characterization a tremendous technical challenge. 13 Thus, high efficiency a nd high purity isolation of CTCs from patient blood is urgently needed to obtain accurate information of CTCs. Currently, the only FDA approved technology is the CellSearch system (Janssen Diagnostics, LLC, Raritan, NJ). Unfortunately, this system is limit ed by low efficiency, low purity and high cost, and does not fully address the issue of isolating the extremely low abundance of CTCs. 37 147 Recently, microfluidic devices with high affinity ligands, including antibodies 17 47 49 72 and aptamers, 75 78 127 have provided distinctive opportunities for efficient and specific isolation of CTCs from patient blood. Microfluidic devices, due to their large surface area t o volume ratio and short diffusion distance, can substantially increase the interaction between cells and the ligand coated surface. 14 103 148 Staggered herringbone micromixers have been developed for fluid mixing in microchannel s 133 and have been explo ited for enhancing the cell capture. 51 72 Yet, limited research has been reported on the optimization of herri ngbone mixers for high performance cell capture. Different from mixing solutions through transverse flow, inducing cell surface interactions requires cells with nearly zero diffusivity for advection to microchannel surface. 149 Herein, we have developed a geometrically enhanced mixing (GEM) chip for high performance CTC capture ( high efficiency purity throughput and cell viability). 150 With experimental optimization of the herringbone micr omixers, we achieved capture of spiked tumor cells with >90% capture efficiency and >84% purity In addition, the time required to process 1 mL blood sample is <17 min, much faster than those reported in literature. 47 51 72 Since very limited work has been

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97 done on cellular studies after CTC capture, 46 we have investigated the release, the viability and the culture of the captured cells. Captured cells can be efficiently released with the combined methods of trypsinization and high flow rate washing. Experiments also sho wed that released cells grew as well as intact cells that had not been subjected to the capture and release process. Further, we applied the device for isolation of CTCs from pancreatic cancer patients, with CTCs observed in 17 of 18 patient samples. We al so demonstrated the potential of using CTC enumeration as a surrogate for radiographic monitoring of chemotherapy response in pancreatic cancer patients. Our device sensitivity enables isolation and enumeration of CTCs from pancreatic cancer patients, a di sease where invasive biopsies are difficult and the commercial CellSearch system has proven to be inefficient. 151 Compared with reported efforts, 47 49 51 72 103 this work demonstrated a systematic study of the following aspects: geometric optimization of micromixer for enhanced target CTC capture, release and re culture of captured tumor cells, c ell viability before and after release, cell binding behaviors after release and re culture, isolation and counting of understudied pancreatic CTCs, comparison of CTC enumeration with CT scans for monitoring chemotherapy response in pancreatic cancer patie nts. A comprehensive study of these aspects would further improve CTC isolation performance help understand post capture processing of CTCs and push forward CTC isolation for cancer diagnosis. In this study we first developed a geometrically enhanced mi xing chip (GEM chip) based on patterned herringbone or chevron structures. The mixer design was inspired by several groups, 51 133 149 and the dimensions were optimized for high efficiency and high purity cell capture. As shown in Figure 4 1, the GEM chip is the same size as a

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98 microscope slide (3 in. 1 i n.), having 8 parallel channels with uniform flow to form a high throughput device. Each channel is 2.1 mm wide, 50 m deep, with 50 m deep herringbone grooves repeating over a total length of 50 mm. The staggered herringbone grooves disrupt streamlines a nd induce chaotic mixing, which maximize collisions and interactions between target cells and device surface, leading to increased cell capture efficiency. The groove width and the groove pitch were carefully selected for high performance cell capture, as discussed in detail in Results and Discussion. 4.2 Experimental Section 4.2.1 Microfluidic Device Fabrication The microfluidic GEM chip consists of a polydimethylsiloxane (PDMS) structure fabricated using two layer soft lithography, according to literature. 30 134 The two layer SU 8 structure (a main channel layer and a herringbone mixer layer) was fabricated via two spin coating and exposure ste ps and a single developing step. The device layout was designed in AutoCAD and then sent to CAD/Art Services, Inc. (Brandon, OR) to produce a high resolution transparency photomask (Figure 4 2) As shown in Figure 4 3, s ilicon wafers were first spin coated with 50 m thick SU 8 2035 photoresist (MicroChem, Newton, MA) as the main channel layer. After soft baking, UV light exposure, and post exposure baking, another layer of SU 8 was added to form the herringbone mixer layer. With precise alignment between t he main channel and the mixer, a second exposure was performed to create the herringbone mixer pattern (Figure 4 4) After development, a silicon master patterned with the complementary structures was obtained. PDMS structures were fabricated by casting a liquid PDMS precursor against the master using Sylgard 184 silicone elastomer kit (Dow Corning, Midland, MI), according to the

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99 (10:1 ratio of base to curing agent) Inlet and outlet wells were created at the channel ends by punc hing holes in the PDMS sheet. The channel depth, which was controlled by the spin speed of the SU 8, was measured using a Dektak 150 profilometer. A high aspect ratio micropillar based PDMS device was also fabricated for cell capture, as shown in Figure 4 5. Further experiments show that mixing device is better than the micropillar device in terms of purity when processing blood samples, thus mixing device was selected for future experiments. Study of f low and shear stress in microchannel is shown in Figur e 4 6, as detailed in Appendix A. 4.2.2 Cell C ulture L3.6pl cells 152 lab (Department of Surge ry, University of Florida). BxPC 3 cells (CRL 1687, human pancreatic adenocarcinoma) and MIAPaCa 2 cells (CRL 1420, human pancreatic carcinoma) were purchased from American Type Culture Collection (ATCC). Cells were cultured in DMEM medium (ATCC) supplemen ted with 10% fetal bovine serum (FBS; heat inactivated; GIBCO) and 100 units/mL penicillin streptomycin (PS, Cellgro, Manassas, VA) and incubated at 37C under 5% CO 2 atmosphere. Cells were grown as adherent monolayers in 60 mm 15 mm culture dishes to 90 % confluence, subsequently detached with 0.05% Trypsin 0.53 mM EDTA (0.05%, Cellgro) and re seeded at a lower concentration. 4.2.3 Reagents and B uffers Biotinylated anti EpCAM (Anti Human CD326, eBioscience, San Diego, CA) immobilized on device surface wa s used as the CTC capture agent. Anti cytokeratin FITC (CAM 5.2, conjugated with fluorescein isothiocyanate, BD Biosciences, San Jose,

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100 CA) and anti CD45 PE (conjugated with phycoerythrin, BD Biosciences) were used to label CTCs and white blood cells, respe diamidino 2 phenylindole, Invitrogen, Carlsbad, CA), which stains DNA in cell nuclei, was used to label all phosphate buffered saline with calcium and m agnesium (PBS, Fisher Scientific, Hampton, NH) was used to wash cells. A buffer containing 10 mg/mL (1%) bovine serum albumin (BSA, Fisher Scientific) and 0.05% Tween 20 (Fisher Scientific) in PBS was used for rinsing the unbound molecules from the channel surface, and resuspending cells for cell capture. BSA and Tween 20 in PBS was used to fully passivate the surfaces to reduce nonspecific adsorption of cells in the channels. Flow cytometry analysis was used to test the binding capabilities of anti EpCAM t o pancreatic cance r cell lines. Fluorescence measurements were performed with a FACScan cytometer (BD Immunocytometry Systems, San Jose, CA). Briefly, 200,000 cells were incubated with 10 g/mL biotinylated anti (containing 0.1% BSA) for 20 min on ice After incubation, the cells were washed three times with PBS. Then streptavidin phycoerythrin (SA PE) Cy5 (Invitrogen) was added and incubated for another 20 min. After washing, 10,000 counts were measured in the flow cytometer to determine the fluoresce nce. The cells incubated with SA PE Cy5 alone were used as a negative control to determine nonspecific binding. Figure 4 7A & B show the strong binding of the anti EpCAM antibody with L3.6pl cells and BxPC 3 cells, respectively. Figure 4 7C which shows no binding between anti EpCAM and MIAPaCa 2 cells, indicates that MIAPaCa 2 cells can be used as a negative control.

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101 4.2.4 Capture o f Spiked Tumor Cells i n Microfluidic Device Immediately before experiments, cells were detached from the culture dish and the n rinsed with PBS and resuspended at 10 6 instructions, the target cells and control cells were stained with Vybrant DiI (red) and Vybrant DiD (blue) cell labeling solutions (Invitrogen), then rinsed with PBS, and r esuspended at 10 6 cells/mL in the PBS containing BSA and Tween 20. Labeled cells were stored on ice and further diluted or spiked into blood to the desired concentrations before experiments. Anti coagulant containing human whole blood from healthy partici pants was experiments. For some experiments, CTC capture from whole blood samples was preceded by red blood cell lysis performed as previously described. 153 Briefly, lysed blood was obtained by treating whole blood with red blood cell (RBC) lysing buffer, prepare d by adding 155 mM (8.3 g/L) ammonium chloride in 0.01 M Tris HCl buffer, with pH=7.5. Different concentrations of cancer cell lines were then spiked in whole blood or lysed blood. The detailed RBC lysis procedure is shown in Appendix B. To initiate cell c apture experiments, one channel volume (~100 L) of 1 mg/mL avidin (Invitrogen) in PBS was first introduced into the device, followed by incubation for 15 min and then three rinses with PBS. Then, one channel volume of biotinylated anti EpCAM (20 g/mL) wa s introduced into the device and incubated for 15 min, followed by three rinses with the PBS containing BSA and Tween 20. Finally, 1 mL of cell mixture or blood sample was pumped into the device at a flow rate of 1 L/s (or other flow rates specified in th e text). At the end of the experiment, the microchannel was

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102 washed three times with PBS, followed by acquiring fluorescent images for the determination of the number of cells captured. 4.2.5 Instrument S etup The cell suspension or blood sample was intro duced into the device by pumping using a syringe pump (KD Legato 111, KD Scientific, Holliston, MA) with a BD syringe connected to the inlet of the device via polymer tubing and a female luer to barb adapter (IDEX Health & Science, Oak Harbor, WA). To avoi d cell settling, a tiny magnetic stirring bar was placed inside the BD syringe, with a stir plate beneath the syringe. The magnetic stirring bar kept cells in suspension while the cell mixture or blood was being pumped through the device. An Olympus IX71 f luorescence microscope (Olympus America, Melville, NY) with an automated ProScan stage (Prior Scientific, Rockland, MA) was used to image and count the captured cells on the device. 4.2.6 Cell R elease and R e culture Cell release was achieved by trypsin an d high flow rate washing. After cells captured inside the channel, proteolytic enzyme trypsin (0.25%) was introduced into the device and incubated for 5 min at 37 C. Then, cell culture medium was pumped into the device at a flow rate of 5 L/s to dislodge the bound cells. The release flow rate was much higher than the cell capturing flow rate of 1 L/s. Released cells were collected in a new cell culture dish (60 mm 15 mm size), with a total volume of 4 mL culture medium. Then the cells were put into the incubator for propagation in culture. To test the viability of cells captured by the device, propidium iodide (PI) and acridine orange (AO) staining (Invitrogen) assays were performed. PI is a membrane impermeant stain that labels only dead cells with re d fluorescence. AO is a membrane permeable dye that binds to nucleic acids of all cells and induces green fluorescence.

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103 contain 2 M PI and 2 M AO in PBS. After incubatin g the working solution with cells for 10 min, fluorescent images were taken to evaluate the viability of the captured cells (Figure 4 12C ). 4.2.7 Patient Blood Specimen Collection a nd Processing Blood samples of patients with metastatic pancreatic cancer were obtained from the University of Florida Health Cancer Center after informed consent through a University of Florida Institutional Review Board (IRB) approved protocol. Blood samples from normal healthy participants were obtained through the Gainesvill e LifeSouth Community Blood Center following a University of Florida IRB approved protocol. Specimens were collected into BD Vacutainer tubes containing anti coagulant sodium heparin and were processed within 6 hours after being drawn. The blood processing safety protocol is shown in Appendix C. CTC capture was performed by the same protocols as described above. Unlike the pre Three color immunocytochemistry (DAPI, FITC anti cy tokeratin, PE anti CD45) was conducted to identify CTCs from nonspecifically captured blood cells. Cell staining began with cell fixation and permeabilization by incubation for 20 min with 4% paraformaldehyde and 0.2% Triton X 100, respectively. Then, a mi xture of 10 g/mL PE anti CD45, 10 g/mL FITC anti cytokeratin and 500 nM DAPI were introduced into the device and incubated for 20 min. After washing, the microfluidic device was examined under the fluoresce microscope. Only cells that were DAPI positive, CD45 negative, cytokeratin positive, with the appropriate size and morphology were counted as CTCs (DAPI+, CD45 cytokeratin+). Cell debris, red blood cells (DAPI ) white blood cells

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104 (DAPI+, CD45+) double positive cells ( both CD45+ and cytokeratin + with DAPI+) were excluded from counting. CTC capture purity was defined as the ratio of the number of CTCs captured to the total number of nucleated cells (DAPI+) bound to the device. For another set s of experiments, we released the specifically capture d CTCs along with nonspecifically captured leukocytes into culture dish (instead of staining and counting). And fresh medium was added once a week (with leukocytes washed away). We observe d a few cells (probably CTCs) adhered to the culture dish after 1 we ek of culture. However, these adhered cells did not proliferate even after 4 months of culturing (unlike the spiked tumor cells which grew into clusters within 2 weeks). 4.3 Results and Discussion 4.3.1 Target Cell Capture f rom a Homogenous Cell Mixture T he performance of the device was first evaluated by sorting a mixture of pancreatic cancer cell lines: target L3.6pl cells (EpCAM+) and control MIAPaCa 2 cells (EpCAM ). Flow cytometry results show that L3.6pl cells bind strongly with anti EpCAM, while MIA Paca 2 cells do not bind with anti EpCAM (Figure 4 7 ). This means that L3.6pl cells express a significant number of EpCAM receptors, while MIAPaCa 2 cells express negligible surface EpCAM, which is consistent with data already reported in literature. 154 To start the cell capture, biotinylated anti EpCAM was first immobilized on the surface of microchannel. Then a cell mixture containing 10 6 L3.6pl cells (stained with Vybra nt DiI, red) and 10 6 MIAPaCa 2 cells (stained with Vybrant DiD, blue) per mL sample were introduced into the microchannel. Figure 4 8A shows a representative image of the cell mixture prior to sorting, with same number of target cells and control cells. Fi gure 4 8B shows a typical image after the cell mixture was processed through the device, with L3.6pl cells in the majority, while most control MIAPaCa 2 cells were removed by

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105 washing. Figure 4 8 A & B clearly indicate that significant enrichment of target c ells can be achieved using the antibody coated microfluidic device. A fter the initial experiments different flow rates were used to study the effects of flow rate on cell capture efficiency defined as the ratio of the number of target cells captured to the number of target cells initially introduced As shown in Figure 4 9A the capture efficiency of L3.6pl cells was >90% at low flow rate s but decreased dramatically at flow rates above 1 L/s, primarily due to the reduced interaction time between the ce lls and antibody coated surfaces as well as the increased shear stress at higher flow rates. To obtain both efficient capture and sufficient throughput, a n optimal flow rate of 1 L/s was chosen, with a flow velocity of 0.75 mm/s and maximum shear stress o f 0.38 dyn/cm 2 at the wall. As shown in Figure 4 9B t he capture efficiency was ( 90 2) % for L3.6pl cells and ( 9 2 4) % for BxPC 3 cells at 1 L/s 4.3.2 Micromixer Device Optimization f or High Performance Cell Capture When we used the traditional microm ixer design dimensions (HB chip, Figure 4 1C ) 51 for pancreatic tumor cell capture, we found that non target cells were easily trapped in the device (causing low C TC capture purity) and cells were not captured on the same focus plane (making imaging and counting difficult). We suspected that cell trapping took place in narrow grooves (with high aspect ratio) as illustrated in Figure 4 1C and hypothesized that an in creased groove width would give better purity. Thus we made two new designs by increasing the groove width from 50 m (narrow groove, Figure 4 1 C ) to 80 m and 120 m (wide groove, Figure 4 1 D ). Experimental results proved that a wider groove with increase d groove pitch achieved high purity cell capture, while maintaining cell capture efficiency. As shown in Figure 4 10 with a groove width of 120 m, we obtained a capture purity of 84%, while the traditional 50 m groove

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106 width yield only 61% purity. In add ition, the capture efficiency for the wide groove design was not reduced even slightly higher, which agrees with simulation study by Forbes et al. 149 4.3.3 T umor Cell Capture f rom Lysed Blood a nd Whole Blood T o test cell capture under more physiological conditions and to mimic CTC capture from patient blood we conducted a series of experiments in which labeled L3.6pl cells were spiked in lysed or whole blood. S ample s were prepared by spiking 50 50,000 L 3.6pl cells in 1 mL lysed blood or whole blood. A fter being pumped through the micromixer device, as many as ~92% of L3.6pl cells were captured from lysed blood (Figure 4 11A ), and ~89% of L3.6pl cells were captured from whole blood (Figure 4 11 B ), proving that the device and the conditions are suitable for capturing CTCs from patient blood specimens with or without prior red blood cell lysis 4.3.4 Cell Release a nd Cell Viability The detachment and release of captured cells in antibody coated microchannels was achieved by using a combination of trypsinization (enzymatic release) 4 8 and high flow rate washing (high shear stress). 71 Detached cells were collected in a cell culture dish with fresh medium for propagation in cell culture. As s hown in Figure 4 12A the release efficiency of L3.6pl cells increased to >60% by using the combined releasing method, while high flow washing alone gave only ~30% release. The release efficiency is defined as the ratio of the number of cells released to t he number of cells captured. The trypsin release and shear stress based release procedures cause minimum cell damage as proved by cell viability assay and flow cytometry. PI/AO assay was used to test the viability of released cells, with >85% cells remaini ng viable after the capturing and release process (Figure 4 12B & C ), making the isolated tumor cells suitable for

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107 subsequent cellular analysis. Flow cytometry tests also showed that released L3.6pl cells retain their binding with anti EpCAM, as shown in F igure 4 12D 4.3.5 Re c ulture of C aptured Tumor C ells To determine whether isolated tumor cells can be re cultured, 5,000 L3.6pl cells were spiked into whole blood and subjected to the capture and release process as discussed above. The released cells were then seeded into cell culture dishes for propagation in culture A s comparison, 5,000 intact L3.6pl cells (not subjected to the culture and release process) were directly seeded for culture with the same conditions. Results showed that both adhered well a nd proliferated on the culture dishes, forming large clusters and colonies by day 9 (shown in Figure 4 13 A & B ) and grow ing to confluence with longer time (14 days), although the captured cells took a little longer to reach confluence than intact cells. T hen we were able to trypsinize these cells and seed them to other culture dishes, where they grew as adherent monolayers. The isolated cells have successfully undergone multiple (>8) passages without loss of viability or detectable changes in behavior. F lo w cytometry tests indicated that the isolated cells maintain binding behavior with anti EpCAM, as shown in Figure 4 13C T hese results clearly demonstrate that tumor cell lines isolated from whole blood retain both their viability and their proliferation a bility, which are crucial for CTC cellular analysis. 4.3.6 Isolation o f C TCs f rom Patients w ith Pancreatic Cancer Using t he GEM Chip Blood samples from patients with metastatic pancreatic cancer (stage I V) were analyzed for CTC enumeration using the above optimized device and conditions. Since EpCAM has been known to be overexpressed in pancreatic adenocarcinoma, anti EpCAM was used as the capture agent. 155 Milliliters of pati ent blood were pumped through the antibody coated device. After fixation and permeabilization, three color

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108 immunocytochemistry was utilized to identify and count CTCs from nonspecifically captured white blood cells, using FITC labeled anti Cytokeratin (CK, green), PE labeled anti CD45 (red) and DAPI (blue) for staining. As shown in Figure 4 14 CTCs are DAPI+/CK+/CD45 cells, while WBCs are DAPI+/CK /CD45+ cells. More images of CTCs were shown in Figure 4 16 and WBCs were shown in Figure 4 17. A significant population of positive cells with both hematopoietic and epithelial markers (CK+/CD45+) were found in quite a few patient samples (average ~2 double cells in 1 mL patient blood). Since the origin and significance of these cells are und er debate, we temporarily excluded them from CTC counting. 51 156 For the 18 pancreatic cancer patient sa mples processed, CTCs were found in 17 cases (>94%), with an average number of 3 CTCs per mL of blood, as shown in Table 4 1. To examine the possibility of false positives, we investigated capturing CTCs from whole blood of normal healthy individuals. Simi lar volumes of blood were run through our device using the same protocol. Table 4 2 shows the results from blood samples of nine healthy donors. Zero CTCs were detected from blood samples of all normal healthy individuals studied, thus showing a false posi tive rate of zero. Additionally, we found much fewer double positive cells in healthy donor blood than in patient blood, indicating that most of the double positive cells could be the heterogeneous CTCs or the nonspecific binding of anti CD45 to CTCs Further studies with additional markers are required to understand and explain these double cells. For capturing CTCs from patient blood, much more nonspecific capture of white blood cells was observed than the spiking experimen ts using health which could due to complexity of patient blood conditions. The high purity of the GEM

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109 chip shows more advantage over the traditional mixing chip when enumerating patient CTCs. The GEM chip would have been able to detect an average of ~23 C TCs from 7.5 mL blood, much higher than the cut off number of CellSearch system. Considering that CellSearch is inefficient for pancreatic cancer, the GEM chip reported here could become a powerful tool for CTC enumeration in pancreatic cancer. I n addition with a flow rate of 1 L/s (3.6 mL/h), 1 mL blood sample can be processed within 17 min, which gives sufficient throughput for clinical applications. We also released and attempted to culture the captured patient CTCs in vitro, using the above mentioned protocol for re culturing spiked tumor cells. Similar release efficiency was obtained for patient CTCs. However, CTCs were not able to proliferate or propagate in the culture dish, although they were found adhered to the culture dish. We processed 12 pancr eatic patient blood samples (with 5 10 mL volume for each sample), but CTCs from them were not proliferat ing till now. We suspect that during the progression of metastasis, CTCs shed from a primary tumor and entered into blood stream might lose their abil ity to proliferate 4.3.7 Monitoring A nti cancer T reatment R esponse U sing CTCs To demonstrate the unique clinical potential of our device and system, we evaluated the relation between the CTC number and tumor size in patients with pancreatic cancer underg oing chemotherapy. Three patients with stage IV metastatic pancreatic cancer (deemed unresectable) were included in the analysis. Each patient received identical standard treatments with the identical palliative chemotherapy and with X ray computed tomogra phy (CT) scans done at the same intervals. B lood samples were collected at baseline and at the first day of each subsequent treatment cycle. CTCs were captured and counted using the device and methods discussed above.

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110 Investigators were blinded to the demo graphic and clinicopathological characteristics of the patients. The number of CTCs captured at different treatment cycles is plotted in Figure 4 15 A C In general the CTC number decreased with continuation of treatment and modeled the CT scan results (w hich represent standard clinical response measurements). The CTC number correlated proportionally with CT scan measured tumor size in each of the three patients Figure 4 15D & E show that tumor size decreased as treatment progressed for patient #3 which was reflected by the trend of CTC number in Figure 4 15C CT scan data from patient #1 and patient #2 also indicated either reduced primary tumor size or reduced metastatic tumor burden (data not shown). Together, t hese results indicate that CTC quanti fica tion using our device correlates with clinical response and findings from CT imaging, but causes significantly less harms to patients than standard clinical radiographic measurements. W ith the noninvasive nature of our approach, it could provide a powerful tool for monitoring early response or failure to cancer treatment and potentially early cancer diagnosis and relapse predict ion 4.4 C onclusion In this chapter we demonstrated an efficient CTC capture platform based on a geometrically enhanced mi xing (GE M) chip The device achieved >90% capture efficiency, >8 4% purity with a throughput of processing 3.6 mL blood in 1 hour T he system was then utilized to isolate CTCs from pancreatic cancer patient blood sample s with CTCs detected in 1 7 of 18 samples We also successfully demonstrated positive correlation in monitoring anti cancer treatment response using the CTC numbers obtained from our device. In addition the captured cells were released from device with > 6 1% release efficiency, and with > 86% viability Furthermore, we demonstrated

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111 the ability to culture the captured cells, a critical requirement for post isolation cellular analysis. Alt hough it is extremely challenging to culture the isolated CTCs from patient blood and to develop a new cell line, our system shows the possibility to culture spiked tumor cells after the sophisticated capture and release process while maintain ing their viability and proliferation capability. Therefore, o ur CTC capture system shows great potential for efficient CTC enric hment, isolation, and cellular / genetic analysis leading to Our future efforts include further improving CTC capture purity, culturing the captured CTCs from patient, cellular and genetic study of isolated CTCs.

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112 Figure 4 1. Picture and d esign of the microfluidic geometrically enhanced mixing chip (GEM chip) A ) Picture of the 3 in. 1 in. microfluidic GEM chip, consisting of eight parallelized channels with single inlet and outlet. B ) Micrograph (4 bright field) of the staggered herringbone grooves inside a channel, showing their asymmetry and periodicity, scale bar = 200 m. C ) A narrow groove design based on reported herringbone (HB) chip, 51 with 50 m groove width, purple dots show cells captured inside channel. D ) Cross sectional view of the wide groove GEM chip, with channel depth of 50 m and groove depth of 50 m; the groove pitch is set to be 200 m, the groove width is chosen to be 120 m.

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113 Figure 4 2 Picture of the photomask used for fabrication of the two layer microfluidic mixing device. A) The mask for the main channel layer; B) the mask for the herringbone mixer layer. Alignment makers were designed on the both sides of the mask; Blank region was designed surround ing the markers, which make the alignment easier.

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114 Figure 4 3 Schem e of SU 8 mold fabrication process for the two layer mixing device. A) With t wo developme nt process, the coating of the second layer is uneven ; B) Single developing process with multiple coating and exposure, with even coating of the second layer

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115 Figure 4 4. 3 D view and SEM image of the herringbone micromixer structure inside the microfluidic channel. A) The overall layout of the device. B) Scanning electron microscopy image (SEM) of the herringbone groove structure in PDMS. The groove width here is 80 m with a groove pitch of 160 m. C D) The patterned SU8 structure on silicon m old, with one layer of large channel, and another layer of herringbone ridges.

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116 Figure 4 5. A high aspect ratio micropillar device tested for cell capture. A) Layout of the device, with overall channel design the same as the micromixer device. The inset is the scanning electron microscopy ( SEM ) image of the array of high aspect ratio PDMS micropillar s B) Microscopy image shows the specific capture of target cells inside the micropillar based device (red), with few control cells (green). The ima ge is an overlay of fluorescence image with bright field image (s cale bar = 50 m ) This micropillar device have more nonspecific cell capture than micromixing device, especially for blood samples, thus mixing device is used for all the subsequent experime nts.

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117 Figure 4 6 Sch em atic illustration of flow and shear stress on cells inside channel. A) The flow inside channel is pressure driven with parabolic velocity profile; while shear stress varies from channel bottom to top ; B) Diagram of the motion of target cells under hydrodynamic flow ; cells experienced shear force from flow and binding force from ligands (antibodies or aptamers).

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118 Figure 4 7 Flow cytometry test of anti EpCAM binding with different types of pancreatic cancer cells. A) L3.6pl cell s; B) BxPC 3 cells ; C) MIAPaCa 2 cells. Data shows that anti EpCAM binds well L3.6pl cells or BxPC 3 cells, while does not bind with MIAPaCa 2 cells, indicating that L3.6pl and BxPC 3 cells express EpCAM, while MIAPaCa 2 cells do not express EpCAM. Strepta vidin phycoerythrin Cy5 (SA PE Cy5 ) was used to label the biotinylated anti EpCAM.

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119 Figure 4 8 Representative image s of cells before and after capture. A ) 1:1 mixture of target L3.6pl cells (red) and control MIAPaCa 2 cells (blue) before sorting; B ) L 3.6pl cells (red) among MIAPaCa 2 cells (blue) after sorting. Scale bar = 50 m. Target cells were efficiently captured while most control cells were removed by washing

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120 Figure 4 9 L3.6pl cell capture efficiency as a function of flow ra te A ) R educed capture occurred at a high flow rate because of a larger shear force and the reduced interaction time between cells and antibody coated surfaces. B ) Capture efficiency of L3.6pl cells and BxPC 3 cells at the optimal flow rate of 1 L/s, with >90% capture efficiency for both types of cells. Error bars represent standard deviations (n=3). Figure 4 10 Comparisons of capture efficiency and purity of L3.6pl cells with different groove width: 50 m (conventional narrow groove HB chip), 80 m, and 120 m (wide groove GEM chip) Capture purity is defined as the ratio of the number of target cells captured to the number of total cells captured. Error bars represent standard deviations (n=3).

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121 Figure 4 11 Regression analysis of the number of the L3.6pl cells captured by the microfluidic device versus the number of the cells spiked in 1 mL of lysed or whole blood. A ) lysed blood ; B ) whole blood. The x axis indicates the number of spiked cells, y is the number of captured cells. Error bars represen t standard deviations (n=3).

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122 Figure 4 12 Cell release and cell viability testing. A ) By high flow rate washing alone, a release efficiency of 34% was obtained; with a combination of trypsinization and high flow rate washing, the release efficienc y reached 62% for L3.6pl cells. B ) Cell viability before cell capture process (extracted directly from culture) is ~99%. Cell viability immediately after cell capture in device is ~89%; after release the viability is ~86%, without significant difference T he high viability indicates that released cells are suitable for subsequent cell culture. Error bars represent standard deviations (n=3). C ) Fluorescence image of the L3.6pl cells after capture and release with PI/AO staining. T he orange ( red merged with g reen ) color indicates nonviable cells (PI and AO staining) while the green color alone indicates viable cells (AO staining alone) Scale bar = 50 m. D ) Flow cytometry test shows that the captured and then released L3.6pl cells maintain their binding capa bility with anti EpCAM, without any differences compared to normal L3.6pl cells

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123 Figure 4 13 Phase contrast micrograph (10 ) of re cultured cells : A ) r e cultured BxPC 3 cells ; B ) r e cultured L3.6pl cells after 9 days of growth. Scale bar = 100 m. C ) Flow cytometry test showing that the captured and then recultured cells maintained their binding capability with anti EpCAM, without any differences compared to intact cells. Figure 4 14 Fluorescence microscope images ( 40 ) of CTCs captured from patie nt blood A ) A representative image of CTCs, with DAPI+, Cytokeratin+ and CD45 ; B ) typical image of white blood cells (WBCs), with DAPI+ CK and CD45+. Scale bar = 10 m.

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124 Figure 4 1 5 CTC number under different treatment cycle correlates with tumor si ze by CT scans. A C ) The number of CTCs per mL of blood from pancreatic cancer patients at different treatment cycles for three patients: A ) patient # 1; B ) patient # 2; C ) patient # 3. D E ) CT scan image of patient #3 at D ) the beginning of the treatment (c ycle 1); E ) the latter stage of treatment (cycle 11); the red arrows indicate regression of the primary pancreatic cancer. E ach treatment cycle is 14 days.

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125 Figure 4 1 6 Fluorescence image s of white blood cells, with DAPI positive, cytokeratin negat ive and CD45 positive (scale bar = 10 m)

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126 Figure 4 1 7 Fluorescence image s of circulating tumor cells, with DAPI positive, cytokera tin positive and CD45 negative (scale bar = 10 m).

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127 Table 4 1. Quantification of CTCs per mL of blood among 18 samples from patients with metastatic pancreatic cancer. Sample No. Cancer type Volume processed (mL) Raw number of CTCs CTCs/mL 1 Pancreas 2 4 2 2 Pancreas 4 14 4 3 Pancreas 2 9 5 4 Pancreas 1 2 2 5 Pancreas 2 2 1 6 Pancreas 2 0 0 7 Pancreas 2 5 3 8 Pancreas 1 2 2 9 Pancreas 2 4 2 10 Pancreas 4 19 5 11 Pancreas 2 5 3 12 Pancreas 2 4 2 13 Pancreas 4 5 1 14 Pancreas 4 15 4 15 Pancreas 4 16 4 16 Pancreas 4 29 7 17 Pancreas 2 6 3 18 Pancreas 2 2 1 Table 4 2. Quantification of CTCs in healthy donor blood. Healthy Sample 1 2 3 4 5 6 7 8 9 Number of CTCs 0 0 0 0 0 0 0 0 0

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128 CHAPTER 5 SUMMARY AND FUTURE WORK 5.1 Summary CTC isolation and enumeration provides an alternative to invasive biopsy for cancer diagnosis and monitoring of c ancer progression. The isolation of CTCs involves two challenging steps: 1) maximizing the capture of target cells; 2) minimizing the capture of non target cells. These two steps correspond to the CTC capture efficiency and purity, or the sensitivity and s pecificity of CTC detection This dissertation presents three novel methods for high performance CTC analysis, including capture, enrichment release, re culture as well as purity and viability testing Devices, techniques and experimental conditions were studied and optimized for achieving these specific goals. First, we developed a microfluidic device with unique micropillar geometry, together with high affinity nucleic acid based ligands, for high efficiency high purity and high throughput cancer cell capture The channel geometry and flow rate was systematically studied to find the optimal geometry and flow rate for the best efficiency and purity of cancer cell capture. With the optimized condition, we achieved efficient capture of as few as 10 colore ctal tumor cells from 1 mL whole blood. The viability of captured cells was proved to be high, making captured cells suitable for subsequent cellular and genetic study. Efficient release of captured cells was achieved using toehold mediated DNA hybridizati on. We also demonstrated cell capture in thermoplastic (COC) device with one step s urface functionalization of COC. Secondly, gold nanoparticle incorporated microfluidic device s w ere developed for high efficiency and high throughput cancer cell capture. I nstead of engineering complicated microstructures inside channel s we used nanoparticle s conjugated with

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129 DNA aptamers for efficient cell capture in a flat channel device The DNA aptamer conjugated nanoparticles allowed multivalent binding, offered increas ed surface area, and packed ligands with high density thus enhanc ing the cancer cell capture. With a passive mixing microdevice, the DNA nanospheres enabled rapid cancer cell capture from whole blood with high efficiency and purity. At last, a geometrical ly enhanced mixing (GEM) microfluidic chip was de velope d for capturing CTCs from patient blood. The antibody coated GEM chip achieved efficient capture of spiked pancreatic tumor cells. We also investigated the release and the re culture of the captured tu mor cells; and captured cells were successfully released and re cultured. Then the device was applied to the capture of CTCs from patients with metastatic pancreatic cancer CTCs w ere precisely verified and enumerated and the CTC number was compared with t he therapeutic treatment response The CTC counts from the GEM chip correspond ed well with clinical CT scans of tumor size Thus, the GEM chip has a great potential for cancer diagnosis treatment response monitoring and cancer prognosis 5.2 F uture Work 5.2.1 Aptamer E nabled Cancer Cell Isolation We have successfully demonstrated isolation of cancer cells from whole blood using aptamer and aptamer conjugated nanoparticle s in microfluidic devices. In the future, we can use EpCAM aptamer s or gold nanoparti cle s conjugated with EpCAM aptamer s for captur ing CTCs from patients with non hematological cancers Using aptamers targeting cancer stem cells (CSCs) we can identify CSCs from patient blood. A ptamer based cell capture will lead to many facile method s for viable release of capture d cells such as synthesis of a disulfide bond on aptamer and synthesis of a

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130 photocleavable linker or biotin on aptamer Aptamers can also be incorporated into hydrogels or branched DNA polymer s coated on device surface s for enha nced cell capture. Besides gold nanoparticles, other nanomaterials such as graphe n e oxides, 157 DNA micelles, 158 and quantum dots can be used for enhanced CTC capture or detection inside microfluidic device 5.2.2 CTC Isolation from Patient Blood We have demonstrated the efficient isolation of CTCs from patients with metas tatic cancer using a geometrically enhanced mixing chip. In the future, we need to further improve the purity of cell capture and reduce the nonspecific capture of non CTCs, making CTC enumeration more accurate. We can also use negative capture methods to capture blood cells and collect CTCs. 159 We can use different antibodies other than anti EpCAM for CTC capture such as anti HER2, anti EGFR and anti PSA We can also incorporate mesenchymal markers to capture t hose CTCs that have already undergone EMT. 160 In addition to the traditional CTC definition by CellSearch system (DAPI+, CD45 and cytokeratin+) new CTC def inition and standards should be set with more statistic data. We need to successfully culture CTCs with cell stimulation and develop new cancer cell line s On chip cell culture techniques can be developed. The biology study of CTCs will lead to the findin g of the mechanism of metastasis. Certain genes in CTCs can be studied to discover tumor micrometastasis and can potentially be used for gene therapy. CTCs can be collected and lysed and tumor genes can be studied with p olymerase chain reaction ( PCR ) 161 and gene sequencing 162 Circulating tumor DNA (CTDNA) 163 can be detected and studied with droplet microfluidics based digital PCR. The combination of CTC and CTDNA analysis 164 will provide a new platform for cancer

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131 diagnosis, monitoring treatment response, caner prognosis, biology and genetic study of cancer progression and metastasis. 5.2.3 Other Potential Methods for CT C Detection One method includes the use of microfluidic droplets for partition ing blood into nanoliter aliquots and detecting CTCs from blood aliquots which will substantially increase the throughput Activatable aptamer probes 165 can be added simultaneously with cells into droplets, so that enhanced staining of cancer cells can be achieved within droplets. Signal amplification method can be applied in droplets for sensitive CTC detection. 166 167 Another method includes incorporating magnetic beads inside microfluid ic device for enhanced CTC isolation with controllable CTC release Other methods include combining size based, di electrophoresis based and magnetic beads based methods together inside a micro chip for efficient CTC isolation. Ideally, a high performance mi crochip should be designed so that blood samples flow into chip and come out with two streams: one is CTCs and the other is blood cells. The microchip should show its simplicity with complicated parts embedded inside chip, and should be easily operated by clinicians and untrained personnel

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132 APPENDIX A VELOCITY AND SHEAR STRESS IN MICROCHANNEL Given the Navier Stoke s equation for incompressible flow driven by a pressure gradient between two parallel stationary plates (2 D channel flow, fully developed, Newtonian fluids): (1) where is the dynamic viscosity of the fluid, is the hydrodynamic pressure, and is flow velocity in x direction, as shown in Fig ure 4 6 S ince it is 2 D, fully developed flow then : (2) Upon integration: (3) w here c 1 and c 2 are constant. W ith boundary conditions at the wall and at the center of channel: (4) where is the channel hei ght T hus: (5) The velocity profile is parabolic.

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133 Volumetric flow rate per unit width (the width is defined to be the length in the z direction) is : (6) where is the channel width. Then, shear stress: (7) The shear stress is linear across the channel, with zer o shear at the channel center (y=0) due to symmetry. S hear stress at the wall: (8)

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134 APPENDIX B PROTOCOL OF RBC LYSIS FOR WHOLE BLOOD SAMPLES 1) C entrifuge blood sample at 2 500 rpm for 8 min 2) C arefully obtain the buffy coat using a disposable, plastic transfer pipet 3) A dd red blood cell ( RBC ) lysis buffer (buffer to blood ratio = 10:1) RBC lysis buffer: 155 mM ammonium chloride 10 mM Tris pH 7.5 4) Incubate in RBC lysis buffer for 15 min 5) Centrifuge at 1200 rpm for 5 min 6) Discard supernatant 7) Repeat step 3 6 till there is minimal presence of RBCs 8) Wash off remaining RBCs with PBS: Add 5 m L of PBS, centrifuge at 1200rpm for 5min, discard supernatant, re suspend pellet in 1 m L PBS 9) P erfor m CTC capture with microfluidic device using the lysed blood

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135 APPENDIX C BLOOD PROCESSING SAFETY PROTOCOL 1) Safety Training: P ersonnel who handle the human blood samples must go through the University of Florida Bloodborne Pathogen (BBP) Training a nd Biomedical Waste (BMW) Training. An initial training and subsequent annual trainings are required. 2) BBP: BBPs are pathogenic microorganisms present in blood and other potentially infectious material (OPIM) that can cause disease in humans. The most comm on BBPs are Hepatitis B virus, Hepatitis C virus and Human immunodeficiency virus (HIV). T he primary route of occupational exposure to BBPs is percutaneous exposure (e.g., cut or puncture with contaminated sharp objects, such as needles, scalpels, glass) Pe rsonnel should have Hepatitis B v accine before handling human blood samples; the vaccine is safe and effective for preventing infection. 3) Exposure Prevention: Always take universal precautions against BBPs. Controls need to be done to protect against BBP exposures, including engineering controls (sharp containers, biosafety cabinet sharps with safety features ) ; safe work practices (e.g., do not recap needles, hand washing) ; administrative controls (decontamination and disinfection ) and personal protective equipment (gloves, goggles lab coats ). 4) Exposure handling : Part of this section is adapted from the University of Florida Bloodborne Pathogen (BBP) Training and Biomedical Waste (BMW) Training handout material.

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136 If one gets an exposure, he or she should wash wound with soap and water for 5 min ute s and flush mucous membranes for 15 min ute s and seek immediate medical attention. Afterwards, he or she should notify supervisor, contact compensation office and allow medical to follow up with appropriate testing and required written opinion. 5) Biomedical waste processing: Biomedical waste must be inactivated by bleach or autoclaving and segregated properly (sharp containers, autoclave bags, biological waste box) then transported by a registered BMW transporter (Stericycle, Inc. Eaton Park, FL). Bio spill kit (bleach, absorbent material, autoclave bags, gloves, safety glasses dust pan and scoop or tongs for broke n glass ) must be stored in lab.

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151 BIOGRAPHICAL SKETCH Weian Sheng was born in Anhui, China in 1988 He received his bachelor degree in mechanical engineering from Huazhong University of Science and Technology, Wuhan, China, in June 200 9 optoe lectronics With strong interest s in mechanical and electronic engineering h e came to the United States for graduate study in January 2010 He enrolled in the PhD program of mechanical engineering at the University of Florida and worked as a graduate rese arch assistant at the Interdisciplinary Microsystems Group He got a Master of Science in mechanical engineering in May 2012. He received his Doctor of Philosophy in mechanical engineering from the University of Florida in December 2013.