Single-Cell Assays: Microfluidics, Genomics, and Drug Discovery (Methods in Molecular Biology, 2689) 1071633228, 9781071633229

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Methods in Molecular Biology 2689

Paul C.H. Li Angela Ruohao Wu  Editors

Single-Cell Assays Microfluidics, Genomics, and Drug Discovery

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

For further volumes: http://www.springer.com/series/7651

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-by step fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

Single-Cell Assays Microfluidics, Genomics, and Drug Discovery

Edited by

Paul C.H. Li Department of Chemistry, Simon Fraser University, Burnaby, BC, Canada

Angela Ruohao Wu Division of Life Science, Hong Kong University of Science and Technology, Kowloon, Hong Kong, China

Editors Paul C.H. Li Department of Chemistry Simon Fraser University Burnaby, BC, Canada

Angela Ruohao Wu Division of Life Science Hong Kong University of Science and Technology Kowloon, Hong Kong, China

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3322-9 ISBN 978-1-0716-3323-6 (eBook) https://doi.org/10.1007/978-1-0716-3323-6 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Cover Illustration Caption: Cover image courtesy of Elaine Lai-Han Leung. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Dedication To Debby and others who have found a new way to live their lives.

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Preface Drug researchers have discovered the opportunities offered by single-cell assays enabled by microtechnology, while people from the microfluidics area now realize the application potential of these technologies in drug discovery. On the one hand, the microfluidic community has prepared single cells and extracellular vesicles in microdevices for drug research. On the other hand, the pharmaceutical community has demonstrated that drug discovery can be realized by micro/nanofluidic chips using cell-based assays. As for microtechnology, while droplet microfluidics have been utilized by many groups (Ko, Han, DiCarlo, Hu, Nagl) for single-cell assays, the capture of single cells has also been achieved by hydrodynamic flow (Leung, Li), physical traps (Ding, Shintaku), region patterning (Chen), electrophoresis (Shintaku), and dielectrophoresis (Qin). Other than singlecell assays (Chaps. 1–15), there are also studies of single extracellular vesicles (Chap. 16). In terms of drug discovery, multidrug resistance has been studied by Leung’s group in lung cancer cells and Wang’s team in ovarian cancer cells. Cancer research is also carried out on pancreatic cancer cells by Hu’s group. Drug candidates, such as small molecules like paclitaxel (Chap. 1), curcumin (Chap. 2), resveratrol (Chap. 2), daunorubicin (Chap. 8), have been investigated, or biologics, like circRNA (Chap. 13), have been studied. Regarding cell types, the Li group studied single glioma cell; Chen’s group examined single osteoblast and adipocyte; Hu’s group investigated single pancreatic cancer cells; and the DiCarlo’s group studied single yeast cells. Single cells obtained from patients with diseases such as non-small cell lung cancer, sarcoma, and pancreatic cancer are studied in Chap. 1, 7, and 13, respectively. Single CTCs have been identified based on the immunofluorescence assay in lung cancer cells (Chap. 1) and sarcoma cells (Chap. 7). Changes in various measurable quantities, such as cell calcium, pH, and cell differentiation, have been measured on single cells in Chaps. 2 and 4, respectively. Nucleic acid molecules were detected by LAMP (BCR-ABL gene) in K562 cells, digital RT-PCR for RNA in EV (Chap. 16), and in pancreatic cancer cells (Chap. 13). Single-cell transfection is also achieved on MCF-7 cells by Ding’s group. PD-1-EGFP plasmid was introduced to the cell for co-expression of genes; the protein expression of EGFP and PD-1 was detected fluorescently. Intracellular coupling is also studied on single fibroblasts by Huang’s group. Furthermore, single cells have been investigated for studies of the genome and transciptome, either after separation of nuclei and cytoplasm (Chap. 14) or without separation in one tube (Chap. 15). Single-cell assays have been achieved in high throughput in a valvebased system by Tamiya’s group, a chip with microtraps by Ding’s group, in single-emulsion droplets by Nagl’s and Hu’s groups, and double-emulsion droplets by Han’s group. This book serves to enable researchers from both microfluidic and pharmaceutical communities to obtain a rapid overview in the state-of-the-art microfluidic single-cell assays, and to get an impression of what possibilities these assays offer to drug discovery. Burnaby, BC, Canada Kowloon, Hong Kong, China

Paul C.H. Li Angela Ruohao Wu

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Contents Dedication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . About the Editors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1 Detection of Single Non-small Cell Lung Cancer Cell Multidrug Resistance with Single-Cell Bioanalyzer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jun Cai, Ya-Bing Cao, and Elaine Lai-Han Leung 2 Cytosolic Calcium Measurement Utilizing a Single-Cell Biochip to Study the Effect of Curcumin and Resveratrol on a Single Glioma Cell . . . . . . . . . . . . . . Abolfazl Rahimi, Hamide Sharifi, and Paul C.H. Li 3 Dielectrophoresis-Assisted Self-Digitization Chip for High-Efficiency Single-Cell Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yuling Qin, Li Wu, and Daniel T. Chiu 4 Optical pH Monitoring in Microdroplet Platforms for Live Cell Experiments Using Colloidal Surfactants. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xuyan Lin, Wenting Qiu, Steevanson Bayer, and Stefan Nagl 5 PicoShells: Hollow Hydrogel Microparticles for High-Throughput Screening of Clonal Libraries . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cayden Williamson, Mark van Zee, and Dino Di Carlo 6 Single-Cell Micropatterning by Non-fouling Hydrogels . . . . . . . . . . . . . . . . . . . . . Wengang Liu, Ting-Hsuan Chen, and Jiandong Ding 7 Microfluidics-Enabled Isolation and Single-Cell Analysis of Circulating Tumor Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Minh-Chau N. Le, Kierstin A. Smith, Morteza Alipanah, Kangfu Chen, Joanne P. Lagmay, and Z. Hugh Fan 8 Discrimination of Multidrug Resistance in Cancer Cells Achieved Using Single-Cell Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Haiyan Wang, Runxuan Zhang, Di Yang, and Xin Wang 9 Microfluidic Approach for Modeling Coupled Circadian Clock . . . . . . . . . . . . . . . Kui Han and Yanyi Huang 10 A High-Throughput Single-Cell Assay on a Valve-Based Microfluidic Platform Applied to Protein Quantification, Immune Response Monitoring, and Drug Discovery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jonathan C. Briones, Wilfred V. Espulgar, Shohei Koyama, Hyota Takamatsu, Masato Saito, and Eiichi Tamiya 11 Single-Cell Exogenous Gene Transfection Analysis Chip. . . . . . . . . . . . . . . . . . . . . Haiyang Xie and Xianting Ding 12 Double Emulsion Flow Cytometry for Rapid Single Genome Detection . . . . . . . Thomas Cowell and Hee-Sun Han

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Single-Cell Analysis of circRNA Using ddPCR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jiayi Peng, Feng Li, Xiangdong Xu, and Shen Hu 14 A SINC-Seq Protocol for the Analysis of Subcellular Gene Expression in Single Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mahmoud N. Abdelmoez and Hirofumi Shintaku 15 Profiling Single-Cell Genome and Transcriptome by scONE-Seq . . . . . . . . . . . . . Lei Yu and Angela Ruohao Wu 16 Single Extracellular Vesicle Analysis Using Droplet Microfluidics . . . . . . . . . . . . . David Eun Reynolds, George Galanis, Yongcheng Wang, and Jina Ko

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Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors MAHMOUD N. ABDELMOEZ • Cluster for Pioneering Research, RIKEN, Wako, Saitama, Japan; Department of Mechanical Power Engineering, Faculty of Engineering, Assiut University, Assiut, Egypt MORTEZA ALIPANAH • Interdisciplinary Microsystems Group, Department of Mechanical and Aerospace Engineering, University of Florida, Gainesville, FL, USA STEEVANSON BAYER • Department of Chemistry, Hong Kong University of Science and Technology, Kowloon, Hong Kong, China JONATHAN C. BRIONES • Life and Medical Photonics Division, Institute for Open and Transdisciplinary Research Initiatives, Osaka University, Osaka, Japan JUN CAI • Faculty of Pharmacy and Food Science, Zhuhai College of Science and Technology, Zhuhai, China; Dr. Neher’s Biophysics Laboratory for Innovative Drug Discovery, State Key Laboratory of Quality Research in Chinese Medicine, Macau Institute for Applied Research in Medicine and Health, Macau University of Science and Technology, Macau, SAR, China YA-BING CAO • Department of Oncology, Kiang Wu Hospital, Macau, SAR, China KANGFU CHEN • Interdisciplinary Microsystems Group, Department of Mechanical and Aerospace Engineering, University of Florida, Gainesville, FL, USA TING-HSUAN CHEN • Department of Biomedical Engineering, City University of Hong Kong, Hong Kong, China DANIEL T. CHIU • Department of Chemistry and Bioengineering, University of Washington, Seattle, WA, USA THOMAS COWELL • Department of Chemistry, University of Illinois at Urbana-Champaign, Urbana, IL, USA DINO DI CARLO • Department of Bioengineering, University of California, Los Angeles, CA, USA; Department of Mechanical and Aerospace Engineering, University of California, Los Angeles, CA, USA; California NanoSystems Institute, University of California, Los Angeles, CA, USA; Jonsson Comprehensive Cancer Center, University of California, Los Angeles, CA, USA JIANDONG DING • State Key Laboratory of Molecular Engineering of Polymers, Department of Macromolecular Science, Fudan University, Shanghai, China XIANTING DING • Institute for Personalized Medicine, School of Biomedical Engineering, Shanghai Jiao Tong University, Shanghai, China WILFRED V. ESPULGAR • Department of Physics, College of Science, De La Salle University, Manila, Philippines Z. HUGH FAN • Interdisciplinary Microsystems Group, Department of Mechanical and Aerospace Engineering, University of Florida, Gainesville, FL, USA; J. Crayton Pruitt Family Department of Biomedical Engineering, Gainesville, FL, USA GEORGE GALANIS • Department of Bioengineering, University of Pennsylvania, Philadelphia, PA, USA HEE-SUN HAN • Department of Chemistry, University of Illinois at Urbana-Champaign, Urbana, IL, USA; Carl R. Woese Institute for Genomic Biology, University of Illinois at Urbana-Champaign, Urbana, IL, USA KUI HAN • Changping Laboratory, Beijing, China

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SHEN HU • Division of Oral and Systemic Health Sciences, School of Dentistry, University of California, Los Angeles, CA, USA; Jonsson Comprehensive Cancer Center, University of California, Los Angeles, CA, USA; California NanoSystems Institute, University of California, Los Angeles, CA, USA YANYI HUANG • Biomedical Pioneering Innovation Center (BIOPIC), College of Chemistry, and Peking-Tsinghua Center for Life Sciences, Peking University, Beijing, China JINA KO • Department of Bioengineering, University of Pennsylvania, Philadelphia, PA, USA; Department of Pathology and Laboratory Medicine, University of Pennsylvania, Philadelphia, PA, USA SHOHEI KOYAMA • Department of Respiratory Medicine and Clinical Immunology, Graduate School of Medicine, Osaka University, Osaka, Japan JOANNE P. LAGMAY • Department of Pediatrics, Gainesville, FL, USA MINH-CHAU N. LE • Interdisciplinary Microsystems Group, Department of Mechanical and Aerospace Engineering, University of Florida, Gainesville, FL, USA ELAINE LAI-HAN LEUNG • Cancer Center, Faculty of Health Science, University of Macau, Macau, SAR, China; MOE Frontiers Science Center for Precision Oncology, University of Macau, Macau, SAR, China FENG LI • Division of Oral and Systemic Health Sciences, School of Dentistry, University of California, Los Angeles, CA, USA PAUL C.H. LI • Department of Chemistry, Simon Fraser University, Burnaby, BC, Canada; Department of Molecular Biology & Biochemistry, Simon Fraser University, Burnaby, BC, Canada XUYAN LIN • Department of Chemistry, Hong Kong University of Science and Technology, Kowloon, Hong Kong, China; Division of Life Sciences, Hong Kong University of Science and Technology, Kowloon, Hong Kong, China WENGANG LIU • Department of Biomedical Engineering, City University of Hong Kong, Hong Kong, China STEFAN NAGL • Department of Chemistry, Hong Kong University of Science and Technology, Kowloon, Hong Kong, China JIAYI PENG • Division of Oral and Systemic Health Sciences, School of Dentistry, University of California, Los Angeles, CA, USA YULING QIN • School of Public Health, Nantong University, Nantong, Jiangsu, P. R. China WENTING QIU • Department of Chemistry, Hong Kong University of Science and Technology, Kowloon, Hong Kong, China ABOLFAZL RAHIMI • Department of Chemistry, Simon Fraser University, Burnaby, BC, Canada DAVID EUN REYNOLDS • Department of Bioengineering, University of Pennsylvania, Philadelphia, PA, USA MASATO SAITO • Life and Medical Photonics Division, Institute for Open and Transdisciplinary Research Initiatives, Osaka University, Osaka, Japan; National Institute of Advanced Industrial Science and Technology, PhotoBIO Open Innovation Laboratory, Osaka, Japan HAMIDE SHARIFI • Department of Chemistry, Simon Fraser University, Burnaby, BC, Canada HIROFUMI SHINTAKU • Cluster for Pioneering Research, RIKEN, Wako, Saitama, Japan KIERSTIN A. SMITH • Interdisciplinary Microsystems Group, Department of Mechanical and Aerospace Engineering, University of Florida, Gainesville, FL, USA

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HYOTA TAKAMATSU • Department of Respiratory Medicine and Clinical Immunology, Graduate School of Medicine, Osaka University, Osaka, Japan EIICHI TAMIYA • National Institute of Advanced Industrial Science and Technology, PhotoBIO Open Innovation Laboratory, Osaka, Japan; The Institute of Scientific and Industrial Research, Osaka University, Osaka, Japan MARK VAN ZEE • Department of Bioengineering, University of California, Los Angeles, CA, USA HAIYAN WANG • Department of Chemistry and Chemical Engineering, Shanxi Datong University, Datong, Shanxi, China XIN WANG • Department of Chemistry and Chemical Engineering, Shanxi Datong University, Datong, Shanxi, China YONGCHENG WANG • Liangzhu Laboratory, Zhejiang University Medical Center, Hangzhou, China; Department of Laboratory Medicine, the First Affiliated Hospital, Zhejiang University School of Medicine, Hangzhou, China CAYDEN WILLIAMSON • Department of Bioengineering, University of California, Los Angeles, CA, USA ANGELA RUOHAO WU • Division of Life Science, The Hong Kong University of Science and Technology, Kowloon, Hong Kong, China; Department of Chemical and Biological Engineering, The Hong Kong University of Science and Technology, Hong Kong, China LI WU • School of Public Health, Nantong University, Nantong, Jiangsu, P. R. China HAIYANG XIE • Institute for Personalized Medicine, School of Biomedical Engineering, Shanghai Jiao Tong University, Shanghai, China XIANGDONG XU • Division of Oral and Systemic Health Sciences, School of Dentistry, University of California, Los Angeles, CA, USA DI YANG • Department of Chemistry and Chemical Engineering, Shanxi Datong University, Datong, Shanxi, China LEI YU • Division of Life Science, The Hong Kong University of Science and Technology, Kowloon, Hong Kong, China RUNXUAN ZHANG • Department of Chemistry and Chemical Engineering, Shanxi Datong University, Datong, Shanxi, China

About the Editors PAUL C.H. L I obtained his PhD in Analytical Chemistry at the University of Toronto in 1995. Then he developed the microfluidic lab-on-a-chip at the University of Alberta during his postdoctoral work. Dr. Li joined Simon Fraser University in 1999, and he was promoted to full professor in 2010. Dr. Li is interested in integrating microfluidics for single-cell analysis in order to study the transport kinetics of chemical compounds (from herbs) on cancer single cells. This study is particularly useful for characterizing multidrug resistant cancer cells, leading to potential improvement in chemotherapy. He is also interested to combine microfluidics with the nucleic acid bioarray for detection of low volumes and low concentrations of fungal pathogenic DNA, KRAS cancer gene DNA, and influenza viral RNA. Dr. Li has written Microfluidic Lab-on-a-chip for Chemical and Biological Analysis and Discovery (2006) and Fundamentals of Lab on a Chip for Biological Analysis and Discovery (2010); he also co-edited Microarray Technology: Methods and Applications (2016) and Multidisciplinary Microfluidic and Nanofluidic Lab-on-a-Chip: Principles and Applications (2021). Dr. Li is invited speaker of major national and international conferences. Dr. Li is the inventor of 4 granted patents and 5 pending patents, and he is the founder and vice president of ZellChip Technologies Inc. specializing in microfluidic-based instruments for life science applications. ANGELA RUOHAO WU completed her PhD and post-doctoral training in Bioengineering at Stanford University. She joined the Hong Kong University of Science and Technology in 2015, jointly appointed in the Division of Life Science and in the Department of Chemical and Biological Engineering, and was promoted to tenured associate professor in 2022. Dr. Wu is one of the earliest in the world to work in single cell sequencing and pioneered the field of microfluidic chromatin immunoprecipitation (ChIP), inventing the first microfluidic chip for automated ChIP. Dr. Wu’s research interests lie at the nexus of bioengineering and molecular biology, using interdisciplinary approaches to build technologies that enable quantitative interrogation of biological systems. Examples include scONE-seq, a single-cell multi-omics sequencing technology to study tumor heterogeneity; MINERVA for viral genome and metatranscriptome sequencing, which was extensively used in COVID-19 clinical research; and algorithms for accurate and rapid integration of single-cell transcriptomic

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datasets. Dr. Wu has been invited to speak at many major international conferences, including a Keynote speech at microTAS, one of the largest microfluidics conferences in the world. Dr. Wu is a recipient of the MIT Technology Review Innovators under 35 Asia award and is also a World Economic Forum Young Scientist. She was a co-founder of Agenovir Corporation, an anti-viral genome editing therapeutics company that was acquired in 2018.

Chapter 1 Detection of Single Non-small Cell Lung Cancer Cell Multidrug Resistance with Single-Cell Bioanalyzer Jun Cai, Ya-Bing Cao, and Elaine Lai-Han Leung Abstract Non-small cell lung cancer (NSCLC) is one of the leading causes of cancer death in the world. Despite the development of various lung cancer treatment methods, including surgery, radiation therapy, endocrine therapy, immunotherapy, and gene therapy, chemotherapy remains the most common approach for treating cancer. The risk of tumors acquiring resistance to chemotherapy remains a significant hurdle to the use of this approach for the successful treatment of various types of cancer. The majority of cancer-related deaths are related to metastasis. Circulating tumor cells (CTCs) are cells that have been detached from the primary tumor or have metastasized and entered the circulation. CTCs can cause metastases in various organs by reaching them through the bloodstream. The CTCs exist in peripheral blood as single cells or as oligoclonal clusters of tumor cells along with platelets and lymphocytes. The detection of CTCs is an important component of liquid biopsy which aids in the diagnosis, treatment, and prognosis of cancer. Here, we describe a method for extracting CTCs from the tumor of patients and using the microfluidic single-cell technique to study the inhibition of multidrug resistance due to drug efflux on a single cancer cell, to propose novel methods that can provide clinicians with more appropriate choices in their diagnostic and treatment approaches. Key words Non-small cell lung cancer, Circulating tumor cells, Chemoresistance, Clinical utility, Liquid biopsy, Single-cell microfluidic chip, Single-cell bioanalyzer

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Introduction Lung cancer is one of the most commonly diagnosed cancers and the leading cause of cancer-related death worldwide, with an estimated 2 million new cases and 1.76 million deaths each year [1, 2]. Lung cancer is classified into the categories of NSCLC and small cell lung cancer (SCLC). NSCLC is less aggressive, but it is generally identified only at the advanced stages. Around 80% of lung cancer are NSCLC, and they can be further subdivided into adenocarcinoma, squamous cell carcinoma, bronchoalveolar carcinoma, and large cell carcinoma [3–5]. Current effective therapies for NSCLC include chemotherapy, surgical resection,

Paul C.H. Li and Angela Ruohao Wu (eds.), Single-Cell Assays: Microfluidics, Genomics, and Drug Discovery, Methods in Molecular Biology, vol. 2689, https://doi.org/10.1007/978-1-0716-3323-6_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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immunotherapies, and targeted therapy [6]. Chemotherapy has been established in the treatment of patients with lung cancer for many years. Nevertheless, the therapeutic effectiveness of chemotherapeutic agents is limited due to the development of multidrug resistance (MDR) in cancer cells [7]. The general mechanism of cancer MDR is the expression of an energy-dependent efflux pump called adenosine triphosphate (ATP) binding cassette (ABC) transporter. In humans, there are 48 members of this transporter protein family, which are involved in a variety of physiological functions, such as transporting lipids, sterols, peptides, and small ions. Among these members, there are at least three transporters, including one that is called P-gp, also known as MDR1 or ABCB1. These transporters have broad drug specificities and can transport a range of structurally diverse compounds, thereby reducing the accumulation of drugs in the cells and diminishing the efficacy of drugs [8, 9]. Therefore, early prediction of MDR and the search for compounds that can inhibit and kill multidrug-resistant tumor cells in patients in the clinic are important. CTCs are cells that have been detached from the primary tumor or have metastasized and entered circulation [10]. CTCs can cause metastases in various organs after reaching them by passing through the bloodstream [11]. The enrichment and identification of CTCs in patient blood samples play an important role in the diagnostic evaluation and personalized treatment of cancer. The single-cell bioanalyzer (SCB) has been recently developed, functioning as an in vitro diagnostic instrument in CTC analysis. This instrument, which uses a microfluidic chip, has been used to study MDR by monitoring the drug efflux of cancer cells in real time. This new microfluidic method has been used to investigate the effect of MDR on drug modulation by studying the accumulation of daunorubicin in MDR leukemia cells [12, 13]. This novel technology contains three key advantages. First, SCB is simple because it does not require multiple cycles of drug absorption and drug efflux. Second, it is faster than the traditional liquid biopsy method. Third, due to its ability to compare the time points before the MDR modulation test, SCB provides a more reliable control. Therefore, here we describe a method for capturing CTCs using the microfluidic chip and using SCB to identify NSCLC patients who are resistant to specific types of chemotherapy; such knowledge can provide clinicians with more appropriate choices in their diagnostic approaches. In our study, CTCs have been isolated from the peripheral blood of NSCLC patients. The CTCs are tested with two reagents. Reagent A is fluorescently labeled paclitaxel, which is a well-known chemotherapy drug used to treat lung cancer patients. Reagent B is a P-glycoprotein (P-gp) inhibitor. At the beginning of the experiment, a single CTC is captured, and its background fluorescence value is measured first. After the signal becomes stable, Reagent A is added, the absorption of Reagent A by the captured single cell is

Measuring Multi-Drug Resistance of Tumor Cells

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measured, and the fluorescence value is recorded. After the fluorescence value is stable, Reagent B is added, and the fluorescence value is observed again. If the uptake of Reagent A by the CTC increases after the addition of Reagent B, it means that the cell is resistant to drug absorption and the cancer patent is resistant to chemotherapy.

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Materials

2.1 Microfluidic Chip and Single-Cell Bioanalyzer (SCB)

1. Obtain the single-cell bioanalyzer (SCB-402) and microfluidic chip (SCC-001) from ZellChip Technologies Inc. 2. The microfluidic chip (2.5 cm × 7.5 cm) is shown in Fig. 1a. It is composed of four channels, four reservoirs, and a chamber (S) containing the cell-retaining structure (Fig. 1b–c). In Fig. 1b, the lower left and right reservoirs (1,2) serve as the sample inlet and waste reservoirs, respectively. The role of reservoirs 3 and 4 is to allow anticancer drugs or P-gp inhibitors to be delivered. The channel depth is 40 μm, while the reservoir is 1 mm deep and 1.5 mm in diameter. The microchip is made of methyl methacrylate. In Fig. 1c, d, the retention structure of the cells within the chamber is used to select and retain individual cancer cells (Fig. 1d) [14]. Based on this active cell trapping strategy, a well-conditioned cell will be established that ensures the successful removal of other unwanted cells [15]. 3. SCB is composed of an imaging system, optical measurement system, fluorescence detection, and analysis system (Fig. 2).

2.2 Reagents and Apparatus

1. 10% FBS. 2. Phosphate buffered saline (PBS). 3. RPMI 1640 medium. 4. Hanks’ Balanced Salt Solution (HBSS). 5. Dimethyl sulfoxide (DMSO). 6. Deionized water. 7. Lymphocyte separation Ficoll solution. 8. EDTA-K2 anticoagulant. 9. Vacuum the blood collection tube. 10. Plastic sample tubes: 50 mL and 1.5 mL. 11. Red blood cell lysis buffer. 12. Circulating Tumor Cell Enumeration Kit (IsoFlux)IsoFlux magnetic beads pre-conjugated with anti-EpCAM and 50 μL anti-EGFR beads. 13. Binding buffer. 14. 1X Fixative solution. 15. Normal donkey serum 10%.

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Fig. 1 The microfluidic chip. It allows the isolation and drug treatment of single cells for measurements on the single-cell bioanalyzer (SCB). (a) Image of the microfluidic device or microchip. (b) The layout of the microchip indicates that reservoirs 3 and 4 are used for drug delivery, whereas reservoirs 1 and 2 serve as the cell inlet and waste, respectively. The letter “S” corresponds to the chamber that consists of the cell retention structure. (c) Schematic diagrams displaying the sorting of a single circulating tumor cell (CTC) and its retention near the cell retention structure. (d) The cancer cell retained within the cell retention structure is shown

Fig. 2 Image of the SCB-402 showing the LED light source, microscope, power supply, and photomultiplier tube (PMT) detector, with a laptop personal computer (PC)

Measuring Multi-Drug Resistance of Tumor Cells

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16. Anti-CD45 Rabbit Primary Antibody. Dilute 1 μL CD45 Primary Antibody with 99 μL binding buffer. 17. Anti-Rabbit Cy™3-conjugated Secondary Antibody. Dilute 1 μL Anti-Rabbit Cy™3-conjugated Secondary Antibody with 199 μL binding buffer. 18. Anti-Cytokeratin FITC-conjugated Antibody. Dilute 1 μL Anti-Cytokeratin FITC-conjugated Antibody with μL binding buffer. 19. Hoechst 33342. 20. Solution C 1X: Add 9 mL of deionized water to 10X buffer C to make 1X buffer C. 21. Reagent A (fluorescently labeled paclitaxel). Add 500 μL of DMSO to Reagent A to make a stock of 2 mg/mL. Take 25 μL of the stock (2 mg/mL) and add 975 μL 1X buffer C to make a solution of 50 μg/mL. This solution will be further diluted 25 times to 2 μg/mL by 1X buffer C. 22. Reagent B (P-gp inhibitor): Add 500 μL of DMSO to Reagent B to make a stock of 20 mg/mL. When it is used, add 475 μL 1X buffer C into the stock to make a solution of 1 mg/mL. This solution will be further diluted five times by 1X buffer C to give 200 μg/mL.

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Methods

3.1 Extraction of CTCs from Blood

1. Use EDTA-K2 anticoagulant vacuum blood collection tube to obtain about 7 mL of a patient peripheral blood sample, and turn the tube upside down five times to adequately mix the anticoagulant and blood. Store the sample at 4 °C. 2. Add lymphocyte separation solution Ficoll to a 50 mL centrifuge tube. 3. Take the anticoagulated peripheral blood and mix it well with sterile PBS at a ratio of 1:1, and use a pipette to slowly superimpose the mixture on the Ficoll surface along the tube wall. Ensure the liquid flow movements are gentle and pay attention to maintaining a clear solution interface (see Note 1). The final volume ratio of peripheral blood, PBS, and lymphocyte separation medium is 1:1:1. 4. Centrifuge the tube at 400 g for 30 min (see Note 2). 5. After centrifugation, the liquids in the tube should be seen to be divided into three layers: the upper layer is plasma and PBS, the lower layer is mainly red blood cells and granulocytes (or polymorphonuclear leukocytes), and the middle layer is lymphocyte separation liquid Ficoll. At the interface between the upper and middle layers, there is a narrow zone of white

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Fig. 3 Extraction of CTCs by Ficoll density gradient centrifugation. CTCs are to be found in the middle layer of mononuclear cells, with the plasma layer on top, and red blood cells, and polymorphonuclear leukocytes settled to the tube bottom

cloudy layer dominated by mononuclear cells, including lymphocytes and monocytes. In addition, the layer contains platelets (Fig. 3). 6. Remove part of the supernatant liquid, leaving about 1 mL of supernatant liquid, insert the pipette into the cloudy layer, and aspirate the mononuclear cells. Put them into another centrifuge tube, add PBS at more than five times the volume of the mononuclear layer, centrifuge at 300 g for 10 min, and wash the cells twice with PBS. 7. After the final centrifugation, discard the supernatant, add red blood cell lysis buffer, and incubate the tube at room temperature for 5 min to lyse the erythrocytes (red blood cells). The lysis time can be appropriately increased or decreased according to the situation. 8. Add 10 mL of PBS to the tube, centrifuge it at 300 g for 10 min, and wash the cells twice with PBS. 9. After the final centrifugation, discard the supernatant, add 50 μL RPMI 1640 containing 10% FBS to resuspend the cells, and then count the cells and calculate the cell viability.

Measuring Multi-Drug Resistance of Tumor Cells

3.2 Purification and Identification of CTCs

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Use the Circulating Tumor Cell Enumeration Kit for the detection of CTCs from biological samples (whole blood, fractionated blood, ascites, etc.). The kit contains antibodies and reagents for purification and immunofluorescence staining of cells to define CTCs as Cytokeratin+, EGFR+, CD45-, nucleated, and intact cells: 1. Put 50 μL magnetic beads pre-conjugated with anti-EpCAM and 50 μL EGFR beads in a sample tube. Use a magnet over the tube to attract the beads, remove the preservative solution, wash once with 100 μL binding buffer, remove the magnet, and add 50 μL binding buffer to resuspend the beads. 2. Add 30 μL EpCAM beads and 30 μL EGFR beads to the cell suspension obtained above (see Subheading 3.1, step 9), mix the beads gently with a pipette tip, and incubate at room temperature for 1.5 h. 3. Use a magnet over the tube to attract the beads, wash twice with 100 μL binding buffer, remove the magnet, and resuspend with 20 μL binding buffer. 4. Place the cell samples in two 1.5 mL sample tubes with 10 μL each, one for single-cell retention and detection (see Subheading 3.3) and the other tube for identification of CTC as follows. 5. Take one tube of the above samples, add 40 μL of the 1X Fixative solution, incubate at room temperature for 20 min, remove the Fixative solution, wash once with 100 μL binding buffer, add 40 μL of 10% Normal donkey serum, and incubate at room temperature for 5 min. 6. Remove 10% Normal donkey serum, add 40 μL diluted AntiCD45 Primary Antibody (see Subheading 3.1, step 16), and incubate at room temperature for 20 min. 7. Remove Anti-CD45 Primary Antibody, wash once with 100 μL binding buffer, and then remove binding buffer; add 40 μL diluted Anti-Rabbit Cy™3-conjugated Secondary Antibody (see Subheading 3.1, step 17) at room temperature for 20 min in the dark. 8. Remove Cy™3-conjugated Secondary Antibody and wash once with 100 μL binding buffer. 9. Remove the binding buffer, add 40 μL diluted AntiCytokeratin FITC-conjugated antibody (see Subheading 3.1, step 18), and store at room temperature for 40 min in the dark. 10. Remove Anti-Cytokeratin FITC-conjugated Antibody, wash once with 100 μL binding buffer, add 40 μL Hoechst 33342, and protect the cells from light at room temperature for 20 min. Place the cell sample on a glass slide, put a coverslip on top of it, and observe and image the cells under a fluorescence microscope (Fig. 4).

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Fig. 4 Purification of CTCs from peripheral blood of patients with non-small cell lung cancer. (a) Images of cancer cells with PBMCs. (b) CTCs are identified by staining with fluorescence-labeled antibodies (EpCAM, Hoechst, CD45) and imaged using a fluorescent microscope. The top four panels indicate the presence of a CTC (stained by EpCAM and Hoechst) with a lymphocyte (stained by CD45); the bottom four panels depict four CTCs 3.3 CTC Retention and Detection

For single-cell analysis, a single CTC must be selected from 10 μl of purified CTC suspension. The same CTC must be able to be retained in a convenient location in the microfluidic chip. This cell retention will help to measure changes in the fluorescence of paclitaxel within the CTC during its delivery. Since NSCLC cells are adherent, they will stick to the appropriate location in the microfluidic chip. The procedure for single-cell selection and drug treatment is as follows: 1. Take a microfluidic single-cell chip out of the package. 2. Put the microfluidic chip on the bench so that the chip label faces up and is on the left side. 3. Place the microfluidic chip on the microscopic stage of the SCB, and adjust the focus of the microscopic objective until the internal structure of the microfluidic chip is clearly visible. 4. Slowly add 5 μL of RPMI 1640 medium to Reservoir 1 (Fig. 1b) with a 10 μL pipette, allowing the medium to fill the entire channel of the microfluidic chip, making sure that all channels and chambers are filled without air bubbles (see Note 3). 5. Use a pipette to fully resuspend the cells in the above cell suspension containing CTCs (see Note 4). Pipette 2 μL of the suspension into reservoir 1. 6. Observe the flow of cells into the cell retention structure S toward reservoir 2.

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7. Adjust the amount of liquid put in reservoir 1 to increase, or in reservoir 2 to decrease, the liquid flow. When the selected CTC reaches the entrance of the retention structure S, wait for 15 min at room temperature to allow cells to attach to the channel surface of the microfluidic chip. 8. After single CTC cells are selected and captured in the chip, start measuring (λex = 480 nm; λem = 527 nm) to monitor the increase in fluorescence intensity due to the accumulation of Reagent A (fluorescently labeled paclitaxel) in the cells. To measure cellular signal and background, two different detection locations, intracellular and extracellular, should be monitored. To accomplish this, the position of the chip is precisely controlled by an automated translation stage, which allows the chip to be moved back and forth between the two detection locations. The translation of the chip is performed in such a way that the cell fluorescence is measured for 10 s and background fluorescence for 60 s. After giving minutes later until the background fluorescence value stabilized, treat the cells with drugs. 9. Add a solution of Reagent A (2 μL) to reservoir 4 (see Note 5). Note that 2 μL reagent A (2 μg/mL) in reservoir 4 and 2 μL buffer C 1X in reservoir 3 will result in 1 μg/mL reagent A interacting with the cell. 10. Note the fluorescent intensity of the cell, and take an image after 5 min. 11. Replace reservoir 3 with 2 μL of reagent B solution (see Note 5). 12. Observe the changes in the fluorescence intensity of the cells on the computer, image them after 5 min, and analyze the data. 3.4 Analysis of Microfluidic Assay Results

Treat the captured CTCs with fluorescently labeled paclitaxel (PTX) for 300 s after acquiring the background fluorescence values. Observe the change in fluorescence intensity; the fluorescence signal should become stable after about 500 s. Then add the P-gp inhibitor (Reagent B), until the fluorescence signal does not increase further, as shown in Fig. 5a. This suggests that no ABC transporters in the captured CTCs are involved in drug efflux. As shown in Fig. 5b, the uptake of PTX by CTCs increases after the addition of P-gp inhibitor. This indicates that the ABC transporters in the captured CTCs, which have caused a certain efflux effect on the drug, are inhibited or blocked, leading to enhanced signals. This enhancement of accumulated PTX in the cell is indicated by the fold increase, which is defined as the ratio of the fluorescence signal of the inhibitor-blocked cell to that of the unblocked cell, as described [16, 17] as follows:

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Fig. 5 Graphs of fluorescence signal value versus time after drug treatment of single CTCs by the drug (fluorescently labeled PTX, 2.0 μg/mL) and the P-gp inhibitor. (a) One CTC does not show signal enhancement after being treated by the P-gp inhibitor; (b) another CTC shows an increase in fluorescence signal after treatment by the P-gp inhibitor

½Fluorescence IntensityPTX = ½Fluorescence IntensityPTXþBackground - ½Fluorescence IntensityBackground Fold Increase of PTX =

4

Fluorescence Signal of the Inhibitor Blocked Cell Fluorescence Signal of the Unblocked Cell

Notes 1. In extracting CTCs from peripheral blood, the whole blood sample can be added to the upper or lower layer of Ficoll, but the two solutions must be clearly separated in the end. 2. The centrifuge speed can be adjusted according to the different Ficoll of the lymphocyte separation medium. In the first step of centrifugation to isolate CTCs, do not set braking or set it at a low level of braking. Otherwise, the separation will result in a layered mess. 3. During centrifugation of CTCs extracted from peripheral blood, if air bubbles appear during the process of adding medium to fill in the channel, use vacuum suction to remove air bubbles and liquid, and then repeat this filling step. 4. The cell suspension must be fully resuspended before being added to the microfluidic chip channel. 5. In the experiment of measuring the drug absorption of a single CTC, choose the appropriate chemotherapeutic drugs (Reagent A) and P-gp inhibitors (Reagent B) according to the needs.

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Acknowledgments This work has been supported by the 2020 Young Qi-Huang Scholar, funded by the National Administration of Traditional Chinese Medicine, and the project is also financially supported by the Start-up Research Grant of University of Macau (SRG202200020-FHS) and the Faculty of Health Sciences, University of Macau. References 1. Thai AA, Solomon BJ, Sequist LV, Gainor JF, Heist RS (2021) Lung cancer. Lancet 398: 535–554 2. Ferlay J, Colombet M, Soerjomataram I, Par˜ eros M, Znaor A, Bray F (2021) kin DM, Pin Cancer statistics for the year 2020: an overview. Int J Cancer 149:778 3. Rami-Porta R, Bolejack V, Giroux DJ, Chansky K, Crowley J, Asamura H, Goldstraw P (2014) The IASLC lung cancer staging project: the new database to inform the eighth edition of the TNM classification of lung cancer. J Thorac Oncol 9:1618–1624 4. Chansky K, Detterbeck FC, Nicholson AG, Rusch VW, Vallie`res E, Groome P, Kennedy C, Krasnik M, Peake M, Shemanski L, Bolejack V, Crowley JJ, Asamura H, Rami-Porta R (2017) The IASLC lung cancer staging project: external validation of the revision of the TNM stage groupings in the eighth edition of the TNM classification of lung cancer. J Thorac Oncol 12:1109–1121 5. Wahbah M, Boroumand N, Castro C, El-Zeky F, Eltorky M (2007) Changing trends in the distribution of the histologic types of lung cancer: a review of 4,439 cases. Ann Diagn Pathol 11:89–96 6. Collins LG, Haines C, Perkel R, Enck RE (2007) Lung cancer: diagnosis and management. Am Fam Physician 75:56–63 7. Jabir NR, Tabrez S, Ashraf GM, Shakil S, Damanhouri GA, Kamal MA (2012) Nanotechnology-based approaches in anticancer research. Int J Nanomedicine 7:4391–4408 8. Dallavalle S, Dobricˇic´ V, Lazzarato L, Gazzano E, Machuqueiro M, Pajeva I, Tsakovska I, Zidar N, Fruttero R (2020) Improvement of conventional anti-cancer drugs as new tools against multidrug resistant tumors. Drug Resist Updat 50:100682 9. Gottesman MM, Ling V (2006) The molecular basis of multidrug resistance in cancer: the early

years of P-glycoprotein research. FEBS Lett 580:998–1009 10. Ashworth TJMJ (1869) A case of cancer in which cells similar to those in the tumours were seen in the blood after death. Aust Med J 14:146 11. Lin D, Shen L, Luo M, Zhang K, Li J, Yang Q, Zhu F, Zhou D, Zheng S, Chen Y, Zhou J (2021) Circulating tumor cells: biology and clinical significance. Signal Transduct Target Ther 6:404 12. Khamenehfar A, Gandhi MK, Chen Y, Hogge DE, Li PC (2016) Dielectrophoretic microfluidic Chip enables single-cell measurements for multidrug resistance in heterogeneous acute myeloid leukemia patient samples. Anal Chem 88:5680–5688 13. Li X, Chen Y, Li PC (2011) A simple and fast microfluidic approach of same-single-cell analysis (SASCA) for the study of multidrug resistance modulation in cancer cells. Lab Chip 11: 1378–1384 14. Li X, Huang J, Tibbits GF, Li PC (2007) Realtime monitoring of intracellular calcium dynamic mobilization of a single cardiomyocyte in a microfluidic chip pertaining to drug discovery. Electrophoresis 28:4723–4733 15. Shen F, Li X, Li PC (2014) Study of flow behaviors on single-cell manipulation and shear stress reduction in microfluidic chips using computational fluid dynamics simulations. Biomicrofluidics 8:014109 16. Khamenehfar A, Wan CP, Li PC, Letchford K, Burt HM (2014) Same-single-cell analysis using the microfluidic biochip to reveal drug accumulation enhancement by an amphiphilic diblock copolymer drug formulation. Anal Bioanal Chem 406:7071–7083 17. Parekh K, Sharifi H, Khamenehfar A, Beischlag TV, Payer RTM, Li PCH (2019) Chapter six – the microfluidic capture of single breast cancer cells for multi-drug resistance assays. In: Allbritton NL, Kovarik ML (eds) Methods in enzymology. Academic

Chapter 2 Cytosolic Calcium Measurement Utilizing a Single-Cell Biochip to Study the Effect of Curcumin and Resveratrol on a Single Glioma Cell Abolfazl Rahimi, Hamide Sharifi, and Paul C.H. Li Abstract A microfluidic method has been developed for real-time measurement of the effects of curcumin on the intracellular calcium concentration in a single glioma cell (U87-MG). This method is based on quantitative fluorescence measurement of intracellular calcium in a cell selected in a single-cell biochip. This biochip consists of three reservoirs, three channels, and a V-shaped cell retention structure. Because of the adherent nature of glioma cells, a single cell can adhere within the aforementioned V-shaped structure. The singlecell calcium measurement will minimize cell damage caused by conventional cell calcium assay methods. Previous studies have shown that curcumin increased cytosolic calcium in glioma cells using the fluorescent dye: Fluo-4. So in this study, the effects of 5 μM and 10 μM solutions of curcumin on the increases of cytosolic calcium in a single glioma cell have been measured. Moreover, the effects of 100 μM and 200 μM of resveratrol are measured. At the final stage of the experiments, ionomycin was used to increase the intracellular calcium to the highest possible level due to dye saturation. It has been demonstrated that microfluidic cell calcium measurement is a real-time cytosolic assay that requires small quantities of reagent, which will have potential uses for drug discovery. Key words Curcumin, Resveratrol, Fluorescence measurements, Intracellular calcium, Biochip, Microfluidic chip, Single glioma cell

1

Introduction The application of a single-cell biochip for drug accumulation was previously published by Noghabi et al. [1]. The study was done by fluorescence analysis of a single cell. A similar biochip platform was previously used to measure intracellular calcium induced by isoliquiritigenin [2] and for monitoring intracellular drug accumulation [3]. In recent years, huge efforts were made to study Chinese herbs such as Curcuma longa [4]. For instance, the polyphenol in Curcuma longa, such as curcumin or (E,E)-1,7-bis(4-hydroxy-3-methoxyphenyl)-1,6-heptadiene-3,5-dione, has shown anticancer and

Paul C.H. Li and Angela Ruohao Wu (eds.), Single-Cell Assays: Microfluidics, Genomics, and Drug Discovery, Methods in Molecular Biology, vol. 2689, https://doi.org/10.1007/978-1-0716-3323-6_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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analgesic effects. Curcumin can affect the transient receptor potential vanilloid 1 which is an important mediator of sensory signals that are related to analgesic effects [5]. On the other hand, resveratrol is a polyphenol compound that acts like an antioxidant and anti-inflammatory which exists in plants such as blueberry, grapes, etc. Calcium measurement induced by curcumin has been performed on a single U87-MG cell. This is an adherent human glioblastoma cell line derived from glioma [6]. Since calcium is a second messenger that controls a diverse range of cellular functions, calcium measurement is one important assay for identifying beneficial drugs that bind to membrane receptors [7]. Flow cytometry is another cytosolic calcium measurement technique [8], but its application is limited since this method requires a large number of cells and it has been designed to perform a bulk analysis but not a single-cell assay. Furthermore, there is time lag between the introduction of a new reagent and cell measurement in the flow cytometry technique which makes it incapable of performing real-time measurements [9]. Single-cell microfluidic analysis can overcome the aforementioned obstacles and will provide a real-time alternative technique to resolve the time lag issue inherited in the flow cytometry technique. Here, a microfluidic single-cell assay has been reported for fast and real-time measurements of cytosolic calcium in a single U87-MG cell after the addition of curcumin and resveratrol [10]. As previously reported, intracellular calcium is elevated due to different cellular mechanisms. The reason for this is the increase in intracellular calcium ion concentration in glioma cell activation of transient receptor potential vanilloid 1 [11]. The single-cell assay that includes different steps such as single-cell selection and retention, cell attachment, addition of reagent, and quantitative measurement of cytosolic calcium will be described in this chapter. Compared to conventional methods, the microfluidic single-cell method obtains intracellular calcium changes in a real-time manner. Furthermore, microfluidic single-cell calcium measurement, which has potential uses for drug discovery, requires low number of cells (i.e. 50–500), uses only 5 μL of test reagent solution, and reduces cell damage with the on-chip dye loading and reagent introduction techniques.

2

Materials

2.1 Solvents and Growth Medium

1. DMSO (99.9%). 2. DMEM/high glucose medium. Supplement this growth medium with sodium pyruvate, 10% fetal bovine serum (FBS), and 1% penicillin.

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2.2

Cells

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1. Obtain U87-MG cells from cryopreserved storage. 2. The cells are cultured in the DMEM growth medium in a 5% CO2 atmosphere at 37 °C. The doubling time for the cell line is ~39 h; hence, change the growth medium twice a week and passage the cells once a week.

2.3 Buffers and Enzymes

1. Hanks’ Balanced Salt Solution (HBSS). 2. Phosphate-buffered saline (PBS). 3. 0.05% trypsin-EDTA.

2.4

Dyes

1. Fluo-4 AM ester (50 μg, special packaging). Dissolve it in 50 mL DMSO to prepare a 1 μg/μL stock solution of the fluorescent calcium probe. Since Fluo-4 AM is a light-sensitive dye, it must be stored in darkness at -20 °C. Freshly dilute the stock before use in HBSS to make a 5.0 μM solution. 2. Trypan blue solution (0.4%). Use this solution to test the viability of the cell under the microscope. Dead cells will become stained after the addition of trypan blue.

2.5

Test Reagents

1. Curcumin. Dissolve curcumin in DMSO to make a stock solution of 100 μM. Dilute it in HBSS to produce 5 μM and 10 μM working solutions. 2. Resveratrol. Dissolve resveratrol in DMSO to make a stock solution of 1000 μM. Dilute it in HBSS to produce 100 μM and 200 μM working solutions. 3. Ionomycin salt (calcium salt). Use this to saturate Fluo-4 in the cell by Ca2+ for calibration purposes. Dissolve ionomycin in DMSO to make the stock solution. Dilute it in HBSS to prepare a working solution containing 10 μM ionomycin and 50 μM CaCl2.

3 3.1

Methods Chip Fabrication

1. Design a glass biochip containing three reservoirs, three channels, and one chamber, see Figs. 1 and 2a. 2. Fabricate the biochip by standard chip cleaning, thin film deposition, photolithography, photoresist development, hydrofluoric acid wet etching, reservoir forming, and chip bonding. Similar to a previously reported paper [12], this chip comprises a chamber that contains a V-shaped cell retention structure for single-cell isolation; see Figs. 1a and 2a, c.

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15 mm

a

1

2

3

30 mm

b c 1

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Fig. 1 Single-cell biochip. (a) Schematic diagram of the biochip with three reservoirs. Reservoir 1 was used for cell suspension introduction and washing and Reservoir 2 for introducing the reagent, and Reservoir 3 is a waste well. The depth of each channel was 40 μm and the depth and diameter and reservoirs was 600 μm and 2.5 mm, respectively. (b) Image of the biochip with channels filled with trypan blue solution beside a Canadian 25-cent coin for size comparison. (c) Detection window locations, sites 1 and 2 (blue boxes) are marked to show the locations to measure the background and cellular signal, respectively, for background correction [18] 3.2 Microscopic System

1. Use an optical system that comprises an inverted microscope (TE300, Nikon) and a CCD camera for bright-field imaging; see Fig. 2b. 2. For easy and clear microscopic observation, use a TV monitor. For image capture, use a video capture card installed in a computer.

Calcium Measurement of a Single Glioma Cell on a Biochip

(a)

Cell inlet reservoir (1)

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Reagent reservoir (2)

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Single Cell Biochip

Excitation source Chip holder CCD Camera

Photomultiplier Tube (PMT)

Inverted microscope

Fig. 2 The microfluidic chip and the imaging/measurement system. (a) The schematic diagram of the microfluidic chip consisting of three solution reservoirs and a cell retention structure. (b) An image of the instrument setup that consists of an inverted microscope, chip holder, excitation source, CCD camera, and PMT. (c) A single U87-MG cell is retained in the cell retention structure that is opposite to the reagent channel leading to the chamber 3.3 Optical Imaging and Fluorescence Measurement

1. Use a Xenon arc lamp coupled with a monochromator for dye excitation; see Fig. 2b. 2. Use a dichroic filter (620 nm) to let only the red light enter the video camera for cell imaging, but allow for the green fluorescent emission to reach the microphotometer system consisting of the photomultiplier tube (PMT) through a detection aperture. 3. Acquire the data by the Felix software. Download the data as an ASCII file for analysis.

3.4 Cell Thawing and Culturing 3.4.1 Thawing Cells

1. Prepare the wash medium (no serum) and DMEM culture medium (with FBS). Make sure the CO2 cell incubator is operating properly. 2. Wear the cryo-glove and remove a cell tower from the liquid nitrogen (LN) tank (Fig. 3a); place the tower on the bench (Fig. 3b); carefully remove the long safety pin from the front side of the tower; make sure the front side always faces upward when handling the tower; remove a box from it (Fig. 3c); and from the box, take out a cryovial and read the label, and place the selected vial in ice in a Styrofoam box on the bench. 3. Carefully replace the box to the tower, put back the safety pin, and put the tower back to LN tank.

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Fig. 3 Cryovials stored in a LN2 tank. (a) A tower being taken out of the tank. (b) A tower, with a handle on top and with a long safety pin securing the boxes, is placed on a bench. (c) A box opened to show labeled cryovials

4. Take out the cryovial, which consists of 1 mL of cryopreserved cells, from the Styrofoam box, and put the vial into a water bath at room temperature (see Note 1). 5. Dry off the outside of the cryovial and spray with a 75% ethanol solution before placing the vial in the biosafety cabinet. 6. Empty the cells from the vial into a 15 mL centrifuge tube; add 9 mL of wash medium (no serum) to mix with the cells; centrifuge the cell suspension down to produce a cell pellet; and remove the supernatant. 7. Add 9 mL of culture medium (with FBS) to the cell pellet; pipette up and down to resuspend the cells; and put the cell suspension in a culture plate or flask, and keep it in the cell incubator. 8. Check the cells for adhesion inside the flask after 24 h (Fig. 4a). If the cells do not attach, add 4–5 mL fresh culture medium. 3.4.2 Subculturing (Passaging) Cells

1. Remove the cell culture plate from the incubator. Check for adherent cells under the microscope (see Note 2). The cells’ confluency should be more than 80%; see Fig. 4b. 2. Remove the culture medium from cell culture plate. 3. Briefly rinse the cell layer with 1 mL PBS solution to remove all traces of serum that contains trypsin inhibitor (see Note 3). 4. Add suitable amount of trypsin-EDTA (usually 2–3 mL) solution to the flask to detach the cells. Wait for 5–15 min and observe cells to disperse under the microscope (see Note 4). 5. Transfer the cell suspension into a centrifuge tube. 6. Rinse the plate with 2 mL of complete growth medium (with FBS), or pure FBS, and transfer the liquid into the centrifuge tube.

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Fig. 4 Culture of adherent cells. (a) Adherent cells (star-shaped) and detached cells (round). (b) Estimated guide of cell confluency (10%, 30%, 50% and 90%)

7. Do step 6 twice. Centrifuge the tube at 1000 rpm for 5 min. Check ATCC website for correct centrifugation force and time suitable for the cell line. 8. After centrifugation, ensure to see a cell pellet at the bottom of the centrifuge tube. Remove the supernatant carefully without disturbing the pellet or removing the cells. Afterward, add 1 mL of complete growth medium into the centrifuge tube and pipette up and down to resuspend the cells. 9. Immediately withdraw 0.5 mL of the cell suspension, and put it in a 1.5 mL vial for the microfluidic experiment. 10. Transfer the rest of the cell suspension into a new plate, and add 9 mL of growth medium to the plate for subculturing (see Note 5). 11. Write the name of cell line, passage date, passage number, and name of the lab on cell plate (see Note 6). Incubate the cell plate at 37 °C. 3.5 Single Glioma Cell Selection and Retention in a Microfluidic Biochip

The first step of single-cell analysis is to select a single live and intact cell from a cancer cell suspension. Then the single cell must be retained in a specific location inside the chip within the observation region defined by the detection aperture during the experiment. Because of the adherent nature of glioma cells, the cells will attach within the V-shaped cell retention structure after incubation for 10 min. This retention structure will provide the opportunity for the measurement of the changes in intracellular curcumin concentrations during the delivery of reagents to the cell without it being flushed away.

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3.5.1 Washing and Priming the Biochip

1. Place the microfluidic biochip onto the microscope stage; adjust the focus knob until the cell retention structure is clearly visible. Ensure the microphotometer is set to “View” (see Note 7). 2. Add an aliquot (5 μL) of 70% ethanol to Reservoir 1, and allow the liquid to fill and prime the channels. Ensure that there are no air bubbles in all channels and the chamber; remove ethanol from Reservoir 1 by gentle suction (see Note 8). 3. Add an aliquot (5 μL) of 70% ethanol to Reservoirs 1 and 2. Examine all channels and the chamber to be free of air bubbles, and pipette to remove ethanol via Reservoir 2, and then Reservoir 1 (see Note 9). 4. Add an aliquot (5 μL) of cell medium solution to Reservoir 2 and then Reservoir 1; examine all channels and the chamber to be free of any air bubbles. Pipette to remove the medium solution via Reservoir 2 and then Reservoir 1. 5. Repeat step 4 two times. 6. Add 5 μL of cell medium solution to Reservoirs 2 and 1, and then Reservoir 3. Pipette to remove the medium solution via Reservoir 2, then Reservoir 1, and then Reservoir 3. 7. Repeat step 6 once. 8. Add an aliquot (5 μL) of cell medium solution to Reservoir 2, then Reservoir 1, and then Reservoir 3, and now, the microfluidic chip is ready for experiment.

3.5.2 Single-Cell Selection and Drug Delivery

1. After priming the chip, pipette to remove as much of the medium solution as possible from only Reservoir 1. Then gently pipette up and down the cell suspension in the vial (see Note 10). Inject a small cell aliquot of 5 μL into Reservoir 1. 2. After the addition of the cell suspension to Reservoir 1, remove a small amount of medium solution to ensure the free movement of cells in channels. 3. Select a single cell by placing a small aliquot (1 μL) of medium to push the cell into the V-shaped cell retention structure. Let the cell be incubated for 5–10 min for adhesion (see Note 11). 4. Load the selected glioma cell with the dye by adding 5 μM of Fluo-4 AM ester. Adjust the aperture window of the microphotometer system to enclose the single cell (see Note 12). Start the fluorescence measurement. After Fluo-4 AM introduction, wait around 600 s for the cell to fully become loaded with the fluorescence calcium probe; see Fig. 5. 5. Treat the cell with 5 μM and 10 μM of curcumin by applying the test reagent to Reservoir 2 (see Note 13). The increase in fluorescence intensity indicates the elevations in the cellular calcium ion concentration; see Fig. 5.

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F max

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F lu o re s c e n c e I n te n s ity

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10 µM curcumin

2000

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5 µM curcumin

1000

Fluo -4 dye

500

0

0

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1000

1500

2000

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Time (second)

Fig. 5 Intracellular calcium measurement in a glioma cell retained in a microfluidic biochip. During loading of the dye (Fluo-4), data is acquired from 0–500 s for Fmin (see Eq. 1). Increases are observed after addition of 5 and 10 μM of curcumin. Finally, after ionomycin is added, data for Fmax (see Eq. 1) is obtained from 2000 to 2500 s

6. Introduce 10 μM ionomycin (containing 50 μM calcium chloride solution) to the cell at 2000 s to acquire the highest fluorescence intensity (Fmax), as shown in Fig. 5; this is to saturate Fluo-4 inside the cell. 7. Convert the fluorescence intensity to intracellular calcium concentration using Eq. 1: Ca2þ i = K d

F - F min F max - F

ð1Þ

where F shows measured fluorescence, Kd represents dissociation constant of Fluo-4 that has the value of 0.35 μM, Fmin is the background fluorescence intensity (in the calcium-free surrounding solution), and Fmax indicates maximum fluorescence induced by ionomycin that facilitates calcium ion entry through the cell membrane to saturate Fluo-4 [12]. 8. Before curcumin addition or in the resting condition, obtain Fmin from 0 to 500 s where dye loading has begun. Obtain Fmax at the end of the experiment from 2000 to 2500 s. The resting

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Fig. 6 Resveratrol-stimulated intracellular calcium measurement in a glioma cell. During loading of the dye (Fluo-4), data is acquired from 0 to 1000 s. Increases are observed after addition of 100 and 200 μM of resveratrol (RES). Finally, after ionomycin is added, data for Fmax (see Eq. 1) is obtained from 5500 to 6500 s

cytosolic calcium concentration in the cell is determined to be 9.1 ± 1.7 nM. After adding 5 μM curcumin, the concentration increases to 38.7 ± 1.8 nM. In the next step, a second significant increase to 420 ± 11 nM occurs after the treatment with 10 μM curcumin. The results are consistent with some literature reports (see Note 14). 9. In a second study, measure the emitted fluorescent intensity from a single glioma cell after adding different concentrations of resveratrol. Select a single glioma cell and load it with Fluo4 AM. Then measure the intracellular calcium levels of the cell as stimulated by 100 μM and 200 μM of resveratrol; see Fig. 6. 3.6 Maintenance of the Glass Microfluidic Chip

During the microfluidic experiment, small particles or cellular debris may cause a channel or reservoir to be clogged or result in fragment residues that may impact future experiments. Therefore, the following protocol outlines the appropriate steps to efficiently and effectively clean a glass microfluidic chip in order to maintain the integrity of future experimental results: 1. Apply a constant suction to Reservoir 3 of the microfluidic chip.

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2. Add 5 μL ddH2O through Reservoirs 1 and 2 to generate a steady liquid flow in the microchannel. 3. Repeat step 2 twice. 4. Add an aliquot (5 μL) of soap solution to Reservoirs 1 and 2, while continuing to apply a constant suction to Reservoir 3 to flush the cleaning solution through the microfluidic chip (see Note 15). 5. Repeat step 4 four times. 6. Remove excess soap solution by adding 5 μL ddH2O through Reservoirs 1 and 2 to generate a steady liquid flow. 7. Repeat step 6 twice. 8. Add 5 μL of 70% ethanol to Reservoirs 1 and 2, while applying a constant suction to Reservoir 3 to remove the liquid through the microfluidic chip. 9. Repeat step 8 twice. Then use suction to each reservoir to allow the removal of excess ethanol from the channels and/or reservoirs. 10. Wait for 30 min until the microfluidic chip is air-dried. Check under the microscope to ensure that no channels are clogged and that the chamber is clean and free of particles or debris. The chip is stored for future experiments.

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Notes 1. Do not leave the cryovial unattended during the cell thawing process. (It is important for cell viability that the cells are thawed and processed quickly to remove DMSO – thawing only takes a few seconds.) When 80% of the frozen volume has thawed, it is ready for the next step. 2. You can confirm that the cells are completely detached from the plate by observing the movement of the cells under the microscope while swirling the plate. 3. FBS contains protease inhibitors, such as α1-antitrypsin, that can deactivate trypsin. So the inhibitor must be removed by washing the cells with PBS before trypsinization [13]. 4. To avoid cell clumping, do not agitate the cells by hitting or shaking the flask while waiting for the cells to detach. Cells that are difficult to detach may be placed at 37 °C to facilitate dispersal. 5. Cells should be subcultured every 3–4 days when cell confluence is more than 80% (see Fig. 4b).

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6. A growing body of literature demonstrates that the passage number affects a cell line’s characteristics over time. Cell lines at high passage numbers experience alterations in morphology, response to stimuli, growth rate, protein expression, and transfection efficiency, as compared to lower-passage cells. 7. To prevent any damages to the PMT, ensure microphotometer dial is switched to “View” while the room light is on. When a red lamp is used, and during fluorescence measurement, the dial is switched to “Measure.” 8. If an air bubble is observed during the loading of 70% ethanol, use suction to remove the air bubble. Then allow the remaining ethanol to evaporate before repeating the step. 9. After successful completion of the priming step by 70% ethanol, all subsequent steps should be swiftly performed to prevent the evaporation of residual ethanol that is left in channels, which would finally defeat the purpose of ethanol priming. 10. While a cell pellet is resuspended in the medium solution by pipetting it up and down, a P1000 micropipette must be used. The use of lower-volume micropipette tips, such as those used by P200 or P10, can damage the cells and reduce their viability. 11. To test if the desired single cell appears to have adhered to the chamber of the microchip during the 10-min incubation period, an aliquot of medium solution can be removed or added to Reservoir 3 to induce liquid flow in order to confirm no change of cell position. 12. During the adjustment of the measurement aperture window, ensure there is some space around the cell. It is because the cell may increase in size during the experiment. 13. Before adding test reagents to Reservoir 2, ensure that it is completely vacant by pipetting out excess solution. This is to prevent dilution of the reagent concentration. 14. There is no literature value regarding intracellular calcium measurements on the U87-MG cell induced by curcumin. But there was reported increase of calcium concentration of 200 nM as induced by 25 μM curcumin in a human parasite [14]. In Jurkat T cells, the effects of curcumin on the intracellular calcium level were measured using fluorimeter cuvettes; but no value of calcium change was reported because there were problems of inconsistent treatment times on test and control cells [15]. Moreover, the influence of 25 μM curcumin on mouse cells loaded with Fluo-4 was monitored using cuvettes, but the cell calcium levels were not reported because curcumin and ionomycin were applied in two separate batches of cells (2 × 106 cells/mL) [16]. Flow cytometry was also used to study the intracellular calcium homeostasis in thyroid cancer cells treated with curcumin for 24 h [17].

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15. Don’t use strong acids (e.g., HCl) or strong bases (e.g., NaOH) to clean the reservoirs or chambers of the microfluidic chip. The soap used in this experiment was made by adding 5 mL of Sparkleen 1 (Fisherbrand) to 1 L of warm water and then stored in a disposable syringe for easy injection into the microfluidic reservoirs and channels.

Acknowledgments Financial support from the Natural Sciences and Engineering Research Council of Canada is sincerely acknowledged. References 1. Noghabi HS, Soo M, Khamenehfar A et al (2019) Dielectrophoretic trapping of single leukemic cells using the conventional and compact optical measurement systems. Electrophoresis 40(10):1478–1485 2. Li X, Xue X, Li PCH (2009) Real-time detection of the early event of cytotoxicity of herbal ingredients on single leukemia cells studied in a microfluidic biochip. Integr Biol 1(1):90–98 3. Khamenehfar A, Wan CP, Li PCH et al (2014) Same-single-cell analysis using the microfluidic biochip to reveal drug accumulation enhancement by an amphiphilic diblock copolymer drug formulation. Anal Bioanal Chem 406: 7071–7083 4. Xu X, Chen D, Ye B et al (2015) Curcumin induces the apoptosis of nonsmall cell lung cancer cells through a calcium signaling pathway. J Mol Med 35(6):1610–1616 5. Kaneko Y, Szallasi A (2014) Transient receptor potential (TRP) channels: a clinical perspective. Br J Pharmacol 171(10):2474–2507 6. Jiang Y, Marinescu VD, Xie Y et al (2017) Glioblastoma cell malignancy and drug sensitivity are affected by the cell of origin. Cell Rep 18(4):977–990 7. Ebashi S (1961) Calcium binding activity of vesicular relaxing factor. J Biochem 50(3): 236–244 8. Adan A, Alizada G, Kiraz Y et al (2017) Flow cytometry: basic principles and applications. Crit Rev Biotechnol 37(2):163–176 9. Zhang X, Yin H, Cooper JM et al (2006) A microfluidic-based system for analysis of single cells based on Ca2+ flux. Electrophoresis 27(24):5093–5100 10. Noghabi HS, Ahmed AQ, Li PCH (2021) Intracellular calcium increases due to curcumin measured using a single-cell biochip. Anal Lett 54:2769–2776

11. Lu J, Ju YT, Li C et al (2016) Effect of TRPV1 combined with lidocaine on cell state and apoptosis of U87-MG glioma cell lines. J Trop Med 9(3):288–292 12. Khamenehfar A, Gandhi MK, Chen Y et al (2016) Dielectrophoretic microfluidic chip enables single-cell measurements for multidrug resistance in heterogeneous acute myeloid leukemia patient samples. Anal Chem 88(11): 5680–5688 13. Chen L, Mao SJ, Larsen WJ (1992) Identification of a factor in fetal bovine serum that stabilizes the cumulus extracellular matrix. A role for a member of the inter-alpha-trypsin inhibitor family. J Biol Chem 267(17):12380–12386 14. Das R, Roy A, Dutta N et al (2008) Reactive oxygen species and imbalance of calcium homeostasis contributes to curcumin induced programmed cell death in Leishmania donovani. Apoptosis 13(7):867–882 15. Shin DH, Seo EY, Pang B et al (2011) Inhibition of Ca2+-release-activated Ca2+ channel (CRAC) and Kchannels by curcumin in Jurkat-T cells. J Pharmacol Sci 115(2): 144–154 16. Takikawa M, Kurimoto Y, Tsuda T (2013) Curcumin stimulates glucagon-like peptide-1 secretion in GLUTag cells via Ca2+/calmodulin-dependent kinase II activation. Biochem Biophys Res Commun 435(2):165–170 17. Zhang L, Cheng X, Xu S et al (2018) Curcumin induces endoplasmic reticulum stress associated apoptosis in human papillary thyroid carcinoma BCPAP cells via disruption of intracellular calcium homeostasis. Medicine 97(24): e11095 18. Peng XY, Li PCH (2005) Extraction of pure cellular fluorescence by cell scanning in a single-cell microchip. Lab Chip 5(11): 1298–1302

Chapter 3 Dielectrophoresis-Assisted Self-Digitization Chip for High-Efficiency Single-Cell Analysis Yuling Qin, Li Wu, and Daniel T. Chiu Abstract Single-cell analysis of cell phenotypic information such as surface protein expression and nucleic acid content is essential for understanding heterogeneity within cell populations. Here the design and use of a dielectrophoresis-assisted self-digitization (SD) microfluidics chip is described; it captures single cells in isolated microchambers with high efficiency for single-cell analysis. The self-digitization chip spontaneously partitions aqueous solution into microchambers through a combination of fluidic forces, interfacial tension, and channel geometry. Single cells are guided to and trapped at the entrances of microchambers by dielectrophoresis (DEP) due to local electric field maxima created by an externally applied AC voltage. Excess cells are flushed away, and trapped cells are released into the chambers and prepared for in situ analysis by turning off the external voltage, by running reaction buffer through the chip, and by sealing the chambers with a flow of an immiscible oil phase through the surrounding channels. The use of this device in single-cell analysis is demonstrated by performing single-cell nucleic acid quantitation based on loopmediated isothermal amplification (LAMP). This platform provides a powerful new tool for single-cell research pertaining to drug discovery. For example, the single-cell genotyping of cancer-related mutant gene observed from the digital chip could be useful biomarker for targeted therapy. Key words Single-cell analysis, Cell heterogeneity, Dielectrophoresis, Self-digitization chip, Loopmediated isothermal amplification (LAMP)

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Introduction Heterogeneity within cell populations poses challenges for bulk analyses, in which information from important but rare cell types is masked by averaging nature of bulk measurements [1–5]. Molecular analysis of single cells is required to understand the function and development of cell subpopulations in disease research and therapeutic drug discovery. An ongoing challenge in single-cell analysis is manipulation of cells of interest to be isolated into low-volume (100 mm) (see Note 15). 5. Desiccator connected to a vacuum line. 6. Oven (capable of at least 60 °C). 7. Razor blade or scalpel (see Note 16). 8. A cyclic olefin copolymer (COC) sheet or a clean plastic sheet. 9. Biopsy punch (1.5 mm) (see Note 17). 10. Microscope glass slides (3 in. × 1 in.). 11. Detergent. 12. Ethanol (100%). Store in a wash bottle. 13. Pressurized nitrogen. 14. UV-ozone cleaner (see Note 18). 15. Parafilm. 2.3 Device Functionalization

1. Programmable syringe pumps. 2. Kimwipes. 3. Syringe (5 mL) with Luer Lock tip. 4. Female Luer-to-barb adapters (1/8 in. inner diameter). 5. Chromatography tubing (1/16 in. outer diameter × 0.010 in. inner diameter). 6. Phosphate-buffered saline (PBS): PBS without calcium and magnesium. 7. Blocking solution: 2% bovine serum albumin (BSA) in PBS (see Note 19). 8. Avidin solution: Add 2 μL of 100 mg/mL avidin to 98 μL of PBS. 9. Biotinylated antibodies: Add 4 μL of 0.5 mg/mL biotin antihuman GD2 monoclonal antibodies and 4 μL of 0.5 mg/mL biotin antihuman CSV monoclonal antibodies to 92 μL of PBS.

2.4 Blood Sample Processing

1. Programmable syringe pumps. 2. Magnetic stirrer. 3. Magnetic stir bar (2 mm × 5 mm). 4. Centrifuge tubes (15 mL and 50 mL). 5. Biosafety cabinet. 6. Serological pipette (10 mL). 7. Blunt-end fill needle (18 G, 1 ½ in. long). 8. Blocking solution: 2% BSA in PBS. 9. Paraformaldehyde (PFA): 4% PFA in PBS. The fixative reagent is used to fix the cells before staining and imaging. 10. Triton X-100: 0.02% Triton X-100 in PBS.

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11. Fluorescence cocktail: Add 0.4 μL of 0.5 mg/mL anti-panCKAlexa Fluor 488, 0.4 μL of 0.5 mg/mL anti-CSV-FITC, and 20 μL of anti-CD45-PE (a commercial solution, see Note 20) to 79.2 μL of blocking solution. Store in a microcentrifuge tube wrapped in aluminum foil (see Note 21). 12. DAPI (4′,6-diamidino-2-phenylindole): 300 μM DAPI in DI water. Store in microcentrifuge tube wrapped in aluminum foil. 13. Petri dish (100 mm) wrapped in aluminum foil. 2.5

CTC Detection

1. Fluorescence microscopy setup: Agilent BioTek Lionheart LX automated microscope, objectives (4×, 10×, 20×), imaging LED (light emitting diode) and filter cubes to match the selected fluorophores in the fluorescence cocktail (see Note 22), computer, and Gen5 Microplate Reader and Imager Software. 2. Transparent adhesive tape.

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Methods The lab personnel performing these procedures should adhere to all regulations regarding the safe use and waste disposal of chemicals, biohazardous materials and reagents, and sharps. Appropriate personal protective equipment (PPE) should always be worn. For the procedures in Subheadings 3.2, 3.3, 3.4 and 3.5, it is recommended that powder-free gloves be used to minimize contaminations.

3.1 Device Design and Silicon Master Mold Fabrication

1. Draw the microfluidic device design in a CAD software. Export the design file to a commercial photomask vendor (see Note 1). Obtain the photomask from the vendor and inspect the quality of the photomask received. 2. Prepare three Pyrex dishes filled with the following reagents: one dish filled with acetone, one dish filled with IPA, and one dish filled with DI water. The level of the reagent inside the dish should be about 2 cm deep. 3. Place a new silicon wafer in the acetone dish. Place the Pyrex dish containing the wafer in an ultrasonic bath and sonicate for 5 min. 4. Transfer the wafer to the IPA dish using a wafer tweezer. Sonicate the wafer in IPA for 5 min. 5. Transfer the wafer to the DI water dish using a wafer tweezer. Sonicate the wafer in the DI water for 5 min.

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6. After sonication, use a wafer tweezer to take the wafer out of the DI water. Immediately place the wafer under a running stream of DI water (either from a wash bottle or a DI faucet). Rinse the wafer with DI water for 10 s. 7. Dry the wafer using pressurized N2. If the tweezer does not provide a strong hold during the drying process, hold the wafer by a gloved hand at the edges to avoid touching the polished side of the wafer. 8. Bake the wafer in an oven for 10 min at 125 °C to further remove moisture on the wafer. 9. Transfer the wafer to an HMDS vapor prime station equipped with a hot plate (see Note 5). Leave the wafer on the hot plate for 3 min to drive off any remaining moisture. Deposit HMDS vapor to the wafer for 30 s. Incubate the wafer on the hot plate for another 1 min. Transfer the wafer from the hot plate to the cold plate and leave the wafer on the cold plate for at least 20 s to cool down (see Note 6). 10. Place the wafer on a spin coater (see Note 8). Run the recipe to clean the nozzle for dispensing AZ1512 photoresist. 11. Once nozzle cleaning is finished, run the recipe to automatically dispense the photoresist and uniformly coat the wafer at a 2 μm resist thickness. 12. Remove the wafer and run the recipe to clean the spin coater. 13. Immediately place the wafer on a leveled hot plate and bake at 112 °C for 2 min. 14. Load the wafer onto the aligner. Take the photomask out of its case and use pressurized N2 to remove dust particles from the feature side of the photomask. Load the photomask and the wafer onto the aligner. Expose the wafer using a UV light source of power output 6.0 mW/cm2 at 365 nm, a desired dosage of 220 mJ/cm2 recommended for 2 μm AZ1512 photoresist thickness, hard contact mode, and an exposure time of 37 s (see Note 9). 15. Immerse the exposed wafer in a Pyrex dish filled with the developer solution (see Note 10). Lightly rock the dish in a circular motion to agitate the developer to speed up the development process. Develop the wafer for 75 s, the duration recommended for 2 μm of the photoresist. 16. After 75 s of development, retrieve the wafer from the developer using a wafer tweezer. Immediately place the developed wafer under a running stream of DI water (either from a wash bottle or a faucet). Rinse the wafer with DI water for 10 s. Rinse the wafer with fresh developer for 5 s to remove any photoresist residues on the wafer. Place the wafer under running DI water again for 10 s. Dry the wafer surface using pressurized N2.

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17. Perform etching on the developed wafer using a DRIE system to achieve an etched depth of about 45 μm (see Note 11). Use a profilometer to check the final etched depth (see Note 12). Make sure that the wafer is completely dry before moving on to step 18. 18. After the etching process, remove the remaining photoresist on the wafer by placing the wafer in a heated photoresist strip bath. It is recommended that the wafer be anchored on a wafer holder for this step and step 19 to avoid wafer breakage during handling. Leave the wafer in the strip bath for 10 min at 70 °C. 19. Immediately place the wafer holder with the wafer in a quick dump rinser. Start a program of three dump-and-fill cycles. 20. After rinsing is completed, remove the wafer from the holder and dry the wafer completely using pressurized N2. 21. Check the final features of the silicon master under a light microscope to make sure that there is no photoresist left. It is optional to check the final depth of the features after removing the photoresist using a profilometer (see Note 12). 22. Perform this step inside a fume hood. Place the silicon master in a desiccator. Using a 100 μL pipette and a clean pipette tip, deliver 30 μL of PFOCTS into a small weigh boat. Place this weigh boat into the desiccator next to the master. Close the desiccator and turn on the vacuum line. Check to make sure that the desiccator is well sealed; this can be verified by lifting up the entire desiccator by grabbing from the lid. Leave the desiccator under vacuum for 30 min to let PFOCTS evaporate into the entire desiccator space to treat the surface of the silicon master. The silicon master is ready for use in device fabrication. 3.2 Device Fabrication with Soft Lithography

1. Weigh out the PDMS prepolymer mixture which contains 10 parts of Sylgard 184 elastomer base and one part of the curing agent on a weigh boat (see Note 23). It is recommended to weigh out 10 parts of the base in the weigh boat first, followed by zeroing the scale and finally adding one part of the curing agent to the same weigh boat. 2. Mix the base and curing agent in the weigh boat thoroughly for 3 min using a metal or plastic spatula. Air bubbles will form as the stirring and mixing continue. Use the number of bubbles observed to gauge the extent of mixing (i.e., the more thorough the mixing, the more bubbles will be seen in the mixture). 3. Put the weigh boat containing the thoroughly mixed PDMS prepolymer mixture in a desiccator and close the lid. Turn on the vacuum to remove the bubbles from the mixture. Check to make sure that the desiccator is well sealed by lifting up the

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Fig. 1 Top view showing the device mold containing the silicon master and the casted PDMS after curing. The PDMS piece is cut around the device features but inside the border of the master. This cutting allows the silicon master to remain adhered to the petri dish after the PDMS piece is peeled off from the mold

entire desiccator from the lid. Be careful to not let the PDMS overflow as the vacuum line is operating. If the mixture appears to overflow from the weigh boat, “purge” the desiccator (see Note 24). Degassing the mixture may take about 15–30 min, depending on the number of bubbles generated during mixing. 4. Once all bubbles are removed from the mixture, turn off the vacuum. “Purge” the desiccator by opening its valve and letting atmospheric air enter the chamber and stabilize the inside pressure. Once the “hissing” sound of air entering the desiccator has stopped, open the lid to retrieve the PDMS prepolymer mixture. 5. Place the silicon master in a petri dish (Fig. 1) (see Note 25). Carefully and slowly pour the degassed mixture directly onto the center of the silicon master mold; carefully and slowly pour the mixture to avoid generating new bubbles. If additional bubbles form during this pouring process, use a clean needle to carefully pop the bubbles without touching the device features on the master. 6. Place the mold containing the PDMS prepolymer mixture into an oven. Bake the mixture at 60 °C for 3 h to make sure it is completely cured (i.e., polymerized). Alternatively, to speed up this curing process, bake at 80 °C for 2 h. 7. Take the cured PDMS out of the oven and let it cool down to room temperature. Using a razor blade or a scalpel, cut out the desired PDMS device out from the mold. Be careful to cut around the device features while staying inside the border of the master mold (Fig. 1). To avoid possibly breaking the master, do not apply excessive pressure on it during cutting.

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8. Gently peel the cut PDMS piece from the master. Place the PDMS piece on a clean COC film or plastic sheet with the device features facing down. Use a biopsy punch to drill holes in the PDMS to create the inlets and outlets of the microfluidic device (see Note 17). 9. Using a razor blade or scalpel, resize the PDMS piece to match the size of the microscope slide (3 in. × 1 in.). 10. Apply 1 mL of detergent (or pump once on a detergent dispenser) onto the feature side of the PDMS piece. Use gloved fingers to gently scrub the PDMS surfaces. Place the PDMS piece under running DI water to rinse off the detergent. Use an ethanol wash bottle to rinse the feature side of the PDMS piece. Repeat the scrubbing of the surface using fingers while wearing gloves. Rinse the feature side of the PDMS piece with the ethanol wash bottle for 5 s. Immediately dry all surfaces of the PDMS piece using pressurized N2. Place the PDMS piece inside the UV-ozone machine with the feature side facing upward. 11. Clean a microscope glass slide using an acetone wash bottle for 5 s. Then, place the microscope slide under running DI water to rinse off acetone. Repeat the acetone washing and DI water rinsing. Using a wash bottle, apply ethanol to the glass slide for 5 s and use gloved fingers to gently scrub the surfaces. Rinse the glass slide under running DI water. Apply ethanol to the glass slide for 5 s again. Immediately dry all surfaces of the microscope glass slide using pressurized N2. Place the glass slide inside the UV-ozone machine. 12. Turn on the UV-ozone machine to treat the exposed surfaces of the PDMS piece and the glass slide for 5 min. After the UV-ozone treatment, wear a pair of new powder-free gloves, and assemble the two device components together by pressing the treated feature side of the PDMS piece against the treated side of the microscope glass slide. Figure 2 shows the completed microfluidic device, depicting main channels and lateral filters. 13. Place the completed microfluidic device inside an oven at 60 °C for at least 20 min to dehydrate the device and strengthen the permanent PDMS-glass bond. 14. The microfluidic device is now ready for surface modification as discussed in Subheading 3.3. If the microfluidic device does not need to be used right away, it can be stored at room temperature in a petri dish, with the device’s inlets and outlets wrapped with Parafilm to prevent dust particles from entering.

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Fig. 2 The microfluidic chip. (a) The chip with four parallel serpentine main channels, one inlet and one outlet. (b) Lateral filters shown in each main channel. (c) A cell captured at the filter based on both immunoaffinity and cell size. Reproduced in part from [11] with permission from the Chemical and Biological Microsystems Society (CBMS). (Copyright 2020 CBMS)

3.3 Device Functionalization

1. Attach 8 in. of tubing to a Luer-to-barb adapter by pushing the one end of the tubing securely into the barbed end of the adapter. Assemble three of these adapter-tubing attachments. To aid the insertion of the tubing into the barbed end of the tubing, cut the end of the tubing at a 45° angle. 2. Fill three sterile 5 mL syringes with Luer-Lock tips with the following reagents. Fill the first syringe with 100% ethanol. Fill the second syringe with PBS. Fill the third syringe with the blocking solution (2% BSA). 3. Connect the three adapter-tubing attachments onto the three 5 mL reagent syringes. Expel air bubbles from the syringes by holding the syringes so that the tips are pointing upward, by tapping the body of the syringe with your fingers to allow the bubbles to rise toward the tip of the syringe, and by pushing the plunger slowly to force the bubbles out of the syringe and tubing (see Note 26). 4. Load the syringe pump with the ethanol syringe. Dispense a few drops of ethanol onto the inlet of the device to start wetting the interior of the device. The ethanol can easily fill the microchannels of the device via capillary action due to the low surface tension of ethanol. Once it has partially filled the device, dispense a few more drops of ethanol onto the inlet to completely cover the opening before inserting the tubing into the inlet of the device. Program the syringe pump to infuse 250 μL at 2 μL/s and start the infusion of ethanol. Place a folded Kimwipe at the outlet of the device to absorb the ethanol waste (see Note 27).

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5. During the ethanol infusion, make sure that there are no bubbles trapped inside the microchannels of the device. If any bubbles are seen, use a needle cap (or an object of similar size) to tap lightly on the PDMS piece during ethanol infusion to drive the bubbles out of the device. Once the infusion is completed, pipette a 100 μL droplet of PBS around the tubing at the inlet and on top of the outlet of the device, before removing the tubing from the device inlet (see Note 28). 6. Load the PBS syringe onto the syringe pump. Connect the PBS syringe to the device and wash the ethanol out of the device (450 μL at 2 μL/s). Again, place a folded Kimwipe at the outlet of the device to absorb the waste. Replace the Kimwipe with a new one before the current wipe is saturated with the reagent waste. 7. Using a 100 μL pipette and new pipette tip, load the tip with 100 μL of avidin. Insert the pipette tip into the device inlet until the tip touches the glass slide. At this point, the pipette and the device form a 90° angle (see Note 29). Slowly push down on the pipette to the second stop to deliver avidin in the pipette tip into the device. Excess reagent will flow out of the outlet (see Note 30). Once the level of the content inside the pipette tip has dropped down to the top of the PDMS piece, carefully depress the pipette to the first stop before removing the tip from the inlet. Dispense a droplet of PBS at the inlet of the device (see Note 31). Incubate the device at room temperature for 15 min. 8. Following avidin incubation, wash the device with PBS (250 μL at 1 μL/s; see step 6). 9. Deliver 100 μL of biotinylated antibodies into the device (see step 7). Allow the antibodies to incubate in the device at room temperature for 15 min (see Note 32). 10. Following the antibodies’ incubation, load the blocking solution syringe onto the syringe pump. Rinse the device with the blocking solution (250 μL at 1 μL/s; see step 6). Incubate the blocking solution in the device at room temperature for 20 min. The device is now ready to be used for blood sample processing. If the blood sample is not ready to be processed, the device can be stored at 4 °C to curtail antibody degradation. 3.4 Blood Sample Processing

1. Prepare an aliquot of PBS of a volume equal to at least that of the blood sample. 2. Prepare new PBS and blocking solution syringes or reuse the ones from those in Subheading 3.3 if they are not contaminated or degraded.

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3. Prepare two new syringe-tubing units: one filled with paraformaldehyde (PFA) and the other with Triton X-100 (see Subheading 3.3, steps 1–3). 4. Assemble an adapter-tubing attachment. Place a magnetic stir bar inside a 5 mL syringe. Fill the syringe with the blocking solution (see Note 19). Connect the adapter-tubing attachment to the syringe. Expel air bubbles from the syringe to make sure that the entire syringe-tubing unit is filled with the blocking solution (see Subheading 3.3, steps 1–3). Let the blocking solution coat the syringe-tubing unit for 20 min at room temperature. 5. Inside a biosafety cabinet, fill a 50 mL centrifuge tube with 40 mL of the blocking solution (2% BSA) to coat the inside of the tube. Place one 10 mL serological pipette in the tube to allow the blocking solution to fill the inside of the pipette and coat it (see Note 33). Let the blocking solution coat a partial length of the pipette and the centrifuge tube at room temperature, and incubate for at least 20 min. 6. After the incubation, transfer the BSA-coated 10 mL serological pipette to a second 50 mL centrifuge tube. Transfer the blocking solution from the first centrifuge tube into the second centrifuge tube (which now contains the serological pipette). The first centrifuge tube is now BSA-treated and ready to contain the blood sample. The second centrifuge tube now functions to hold the pipette. 7. Take the vacutainer tube containing the blood sample out of 4 °C storage and put it into the biosafety cabinet (see Note 34). Invert the vacutainer tube several times to ensure that the blood sample is homogeneous. If the entire volume in the vacutainer tube is subjected to processing by the microfluidic device, uncap the vacutainer tube and empty the entire content into the BSA-treated centrifuge tube. Estimate the volume of the blood sample by reading the volume markings on the centrifuge tube. If only an aliquot of the blood sample is subjected to microfluidic device processing, use the BSA-coated serological pipette to transfer the aliquot to the BSA-treated centrifuge tube (see Note 35). 8. If the entire content in the vacutainer is transferred, using the BSA-coated pipette, draw up a volume of PBS equal to that of the blood sample, and deliver the PBS to the vacutainer for rinsing. Set the serological pipette controller at the lowest operating speed for liquid transfer to minimize damage to the blood sample during handling. Rinse the inside of the vacutainer by pipetting PBS up and down several times. Be very careful not to exceed the BSA-coated region inside the serological pipette every time a volume is drawn up inside it. This is to

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ensure that all operations pertaining to the handling of the blood sample stay within the regions treated with the blocking solution. 9. Once the inside of the vacutainer is rinsed, deliver the content to the centrifuge tube containing the blood sample. Again, be cautious not to exceed the BSA-coated region inside the serological pipette during transferring, even if this means multiple transfers are to be done. At this point, the blood sample has been diluted 1:1 with PBS to reduce the viscosity of blood. 10. Thoroughly mix the diluted blood sample by pipetting up and down the blood-PBS mixture while staying inside the BSA-coated region of the serological pipette. Perform mixing until the diluted blood sample becomes homogenous. 11. Retrieve the BSA-coated syringe-tubing unit. Detach the adapter-tubing attachment from the syringe. Attach a bluntend fill needle to the syringe (see Note 36). Dispose of the blocking solution content inside the syringe through the blunt fill needle. 12. Draw up the diluted blood sample via the blunt fill needle. Once the syringe is filled (or when all the contents have been withdrawn), carefully detach the blunt fill needle, and discard it according to applicable biohazardous waste disposal guidelines. Carefully reconnect the adapter-tubing attachment to the syringe to prevent blood from spilling from the syringe. Expel air bubbles from the syringe now filled with the blood sample (see Subheading 3.3, step 3). 13. Load the syringe containing the blood sample onto the syringe pump. Place an object underneath one end of the syringe pump to tilt it so that tip of the sample syringe is angled down by ~30° (Fig. 3). Place a magnetic stirrer centered below the tip of the

Fig. 3 The setup for the infusion of the blood sample into the microfluidic device. An object is used to tilt the syringe pump down with an angle of 30°. The magnetic stirrer is placed to activate the stir bar placed inside the syringe barrel. A 15 mL tube is placed at the chip outlet to collect the infusion waste

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syringe so that the magnetic stir bar inside the syringe is stirring its content continuously (Fig. 3). This stirring is done to keep the blood cells from settling down while the sample infusion is in progress. 14. Program the syringe pump to dispense the first 200 μL of the blood sample at 2 μL/s to ensure that there are no more air bubbles present in the syringe-tubing unit. While the blood is being dispensed, collect the waste at the end of the tubing with a 15 mL centrifuge tube (Fig. 3). From this step onward, all waste collected in the 15 mL tube and at the device outlet are considered biohazardous waste. The biohazardous waste should be handled according to applicable safety and disposal guidelines. 15. Connect the syringe tubing to the inlet of the microfluidic device. Connect another small piece of the chromatography tubing to the outlet of the device. Place the other end of this small tubing in the 15 mL centrifuge tube previously used as a waste collector in step 14. Program the syringe pump to introduce the rest of the blood sample to the microfluidic device at 1 μL/s (see Note 37). The setup for the sample infusion step is illustrated in Fig. 3. 16. At the end of the blood sample infusion, disconnect the sample syringe from the device inlet, and connect the PBS syringe to the device inlet to wash out nonspecifically adhered cells from the device by pumping 450 μL PBS at 2 μL/s. At the end of the PBS washing step, disconnect the tubing at the device outlet. 17. Introduce 150 μL of the PFA solution into the device at 1 μL/ s (see Note 38). Incubate at room temperature for 10 min. After the incubation, wash the device with PBS (250 μL at 2 μL/s; see Note 27). 18. Introduce the Triton X-100 solution into the device (150 μL at 1 μL/s). Incubate at room temperature for 10 min. After the incubation, wash the device with PBS (250 μL at 2 μL/s; see Note 27). 19. Introduce the blocking solution into the device (150 μL at 1 μL/s; see Note 27). Incubate at room temperature for 30 min. 20. Using a 100 μL pipette and new pipette tip, load the tip with 100 μL of the prepared fluorescence cocktail. Deliver the fluorescence cocktail to the device (see Subheading 3.3, step 7). Place the device inside a petri dish wrapped in aluminum foil. Incubate the mixture in the device at room temperature for 1 h. After incubation, wash the channels with PBS (450 μL at 2 μL/s; see Note 27).

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21. Deliver 100 μL of the prepared DAPI solution into the device (see Subheading 3.3, step 7). Place the device inside a petri dish wrapped in aluminum foil. Incubate it at room temperature for 10 min. After incubation, wash the device with PBS (450 μL at 2 μL/s; see Note 27). Store the device at 4 °C in the foilwrapped petri dish until ready for CTC detection. 3.5

CTC Detection

1. If the microfluidic device has been stored at 4 °C prior to detection, take the device out and wait for 10–15 min while shielded from light. This waiting time will allow the sample to return to room temperature before imaging. 2. Turn on the computer, the fluorescence microscope, and the image capture and analysis software associated with the microscope. 3. Once the device has returned to room temperature, wipe the outside of the device and dry with Kimwipes. Cover the inlet and outlet of the device with a small piece of transparent adhesive tape. This is done to prevent the evaporation of the fluid inside the device during imaging. Place the device in the microscope slide slot of the fluorescence microscope. 4. Use the bright-field mode and the 4× objective to briefly inspect all microchannels of the device. Then, specify the boundaries of the four microchannels of the device. The boundaries will guide the automated imaging of the device. 5. Switch to the RFP channel (see Note 22) while staying on the 4× objective. This fluorescence imaging channel will show the signals coming from the blood cells expressing CD45 conjugated with the PE fluorophore. Adjust the exposure settings in the RFP channel by referencing the exposure histogram. Finetune the focus in all four microchannels, and take the average focal plane value as the focus used for imaging in the RFP channel. 6. Switch to the GFP channel (see Note 22) while staying on the 4× objective. The GFP imaging channel recognizes the signals coming from Alexa Fluor 488 and FITC fluorophores. Use the average focal plane value previously found in step 5, because both steps are detecting signals coming from either the cell membrane or the cytoplasm. Adjust the exposure settings in the GFP channel by referencing the exposure histogram. 7. Repeat step 5 for the DAPI channel (see Note 22) to find the appropriate exposure and focus settings for this channel. The focal plane value for DAPI should be slightly different from that used for RFP and GFP, because DAPI detects the signal coming from the cell nuclei. 8. Program for the microscope to image the device microchannels in all four imaging channels (bright-field, DAPI, GFP, and RFP) at the specified exposure settings and focal plane values.

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Fig. 4 Representative images of a sarcoma CTC detected during image analysis. The signal intensity in each fluorescence channel is considered as positive based on the signal thresholds previously established using controls. Potential CTCs are identified (circled in red) if they satisfy the criteria of DAPI+(panCK+CSV)+CD45-, as shown in the merged image

9. Following the imaging of the microchannels, perform the merging of the three fluorescence imaging channels (DAPI, GFP, and RFP) using the imaging and analysis software (Gen5), while keeping the copies of the four separate imaging channels for future use. The images are now ready for analysis. 10. Image analysis can be done by specifying the signal detection threshold for each of the three fluorescence channels (DAPI, GFP, and RFP). Potential CTCs are defined as cells being positive for DAPI in the DAPI channel, positive for panCKAlexa Fluor 488 and/or CSV-FITC in the GFP channel, and negative for CD45-PE in the RFP channel. Signals meeting such criteria are identified as potential CTCs by the analysis software, as shown in Fig. 4. It is strongly recommended that a trained lab personnel manually confirms the potential CTCs identified by the software as true CTCs. Morphological features that can help this confirmation process include cell size, cell shape, cytoplasmic characteristic of the GFP signal, and the position of the cell nucleus in relation to the cytoplasm.

4 Notes 1. Check with the photomask fabrication vendor for the acceptable design file format (e.g., DXF, GDSII, and OASIS). The file format will help determine the best CAD software to use for chip feature designing. 2. Use a wafer tweezer purchased from the Electron Microscopy Sciences. 3. DI water can be stored in a wash bottle or be used directly from a DI water lab faucet. The DI water must meet the quality standard required for cleanroom use. 4. Pyrex glass dishes may be reused for another reagent after three rinses with DI water.

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5. HMDS is used to promote photoresist adhesion. It is important that the silicon wafer be heated to 90–130 °C to drive off moisture prior to applying HMDS. Here, use an HMDS vapor prime station equipped with a hot plate. Different cleanrooms may have different types of HMDS prime stations. Consult the standard operating procedure (SOP) for the HMDS equipment provided by the cleanroom staff. 6. If a cold-temperature device or plate is not available, a stack of a few cleanroom wipes placed underneath the silicon wafer can also be used to facilitate the cooling of a hot wafer. 7. Positive photoresist AZ1512 is recommended. The material recommended in Subheading 2.1 is based on the choice of AZ1512 as a photoresist. However, other positive photoresists capable of a resist film thickness of Laminar Flow > Single-Phase Flow > Laminar Flow (spf). Click “Add.” Add Multiphysics and Fluid-Structure Interaction (fsi). Click Study. 4. In the “Select Study” menu, choose “General Studies” and click “Stationary.” Click Done. 5. Using the CAD design, convert the 2D drawing of the microchamber unit to a 3D model. This can be imported to the COMSOL Multiphysics software by clicking the “Geometry” tab and choosing “Import.” The geometry of the microchamber can also be built in the user interface by choosing the shapes in the “Primitive” tab. Input the radius, height, and position (x,y,z) of each object. The axis type can be set to z-axis. Choose the appropriate “Booleans and Partitions” action such as “intersection, difference, and union” to create the target geometry. Click “build or build all” to build the design. In this setup, the cylindrical-ring design is compared to a step-like design to investigate the geometry’s effect on the sealing and rupture pressures (Fig. 1). 6. To set the material of the object comprising the microchamber unit, click “materials” in the menu and select “browse materials” in the built-in materials library. For PDMS, select MEMS, polymers, then PDMS. Click done. In the material setting for PDMS, select the domain by clicking on the geometry, the region for PDMS. The material properties can be changed from their default values. For a linear

Fig. 1 COMSOL Multiphysics simulation was used to evaluate the characteristics of varying designs and specifications of a microvalve-chamber unit: (a) mesh structures of valve designs indicating the outer radius R, inner radius r, and height h and (b) modeling of the valve actuation and chamber sealing

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elastic material, the density can be set to 950[kg/m3], Young’s modulus to 2.63 MPa, and Poisson’s ratio to 0.49. Select the air material for the valve channel and glass for the substrate. 7. In the “Solid Mechanics (solid),” select the domains in the geometry displaced in the “Graphics” window (see Note 3). 8. Under the “Solid Mechanics (solid),” select Linear Elastic Material. Select user defined in “Absolute pressure” changing it to 1[atm]. In the same menu, select “Hyper Elastic Material.” In its setting window, select “Neo-Hookean” in material model and “Nearly incompressible material” in compressibility (see Note 4). In the “Physics” tab, select “Boundary” then “fixed constraint” in the Solid Mechanics section. Select in the boundary selection the glass substrate surface, the surfaces of pillars and traps facing the substrate, and the top surface of the control layer as a fixed constraint (see Note 5). Add “free” boundary and set boundary selection to all boundaries. Boundaries identified as fixed constraints are automatically set as not applicable. 9. Under the “Laminar Flow (spf),” select the domains of the valve/valve channel in the domain selection. Set the compressibility to “Weakly compressible flow” (see Note 6). Under the “Physics” tab, select “Boundaries” and then Wall in the Laminar flow section. Select all the boundaries making up the valve/valve channel in the Boundary Selection. In the menu setting, select “No slip” in the Wall condition. Under the “Physics” tab, select “Boundaries” and then Inlet in the Laminar flow section. Select one end of the valve channel in the Boundary Selection. Select “Pressure” in the Boundary Condition. Enter “para*1[MPa]” as P0 in the Pressure Condition. 10. Select Mesh in the Model Builder menu. Select “Physics-controlled mesh” in the Sequence type and “Finer” in the Element size. Click Build All. 11. In the “Definitions” tab, click “Pairs” and then select “Contact Pair.” Select the top surface of the glass substrate in the “Source Boundaries” and the pillar surfaces connecting to the control layer in the “Destination Boundaries.” Add another “Contact Pair” in the “Definitions.” Select the top surface of the glass substrate in the “Source Boundaries” and the bottom surface of the control layer (on top of the pillars) in the “Destination Boundaries.” 12. In the “Definitions” tab, click “Moving Mesh” and select “Deformation Domain.” Select all the domains comprising the valve/valve channel. In the settings window, choose “Hyperelastic” in the Mesh smoothing type.

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13. In the Model Builder tree, click “Parameters” under “Global Definitions.” In the settings window, enter the following in the Parameters: “para” under Name, “0” under Expressions, 0 under Value, and “pressure value” under Description column. 14. Set up an auxiliary continuation sweep for the parameter “para.” Click “Study” and then “Stationary” under the Model Builder tree. In the settings window for Stationary, click to expand the Study extensions section. Select the “Auxiliary sweep” check box. Click add (+) and in the table select “para (pressure value)” under Parameter name and input “range(0,0.01,0.3)” under Parameter value list. The range is arbitrary (see Note 7). 15. In the “Home” tab, click “Compute” to see the results of the simulation (see Note 8). 3.2 Microfluidic Device Fabrication 3.2.1 Design Considerations of the Microvalve Chamber

3.2.2 SU8 Mold Fabrication

To create the multilayer microfluidic device, an epoxy-based mold is fabricated according to the design of the flow and control layers (Fig. 2). AutoCAD is a common choice for making the designs for these layers. In the flow layer, a major consideration is the size of the main channel, separation distance of the microstructure array, and the trap design. For the control layer, a ring-shaped structure is preferred over a rectangular shape for the reason and purpose stated in the previous section. The separation distance between each structure must also be considered to determine the density of the microchambers. 1. Prepare a 4 inch (or any size depending on design) silicon wafer and heat it on a hot plate at 120 °C for 15 min to remove any moisture on its surface. Mount it on a spin coater. SU8-negative photoresist is preferred in this mold fabrication via photolithography (see Note 9). 2. Determine the desired thickness and set the spin coater program to the desired RPM at different spin steps. For a 25 μm thickness and using SU8 3025, the spin-coating program steps can be as follows:

Fig. 2 Image of the fabricated molds for flow layer (left) and control layer (right)

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Table 1 Negative photoresist formulation and spin coating speed used to fabricate the mold layers on silicon wafer. Different combinations of other SU8 formulations and spin speed (in RPM) can also be used to achieve desired height Pre-exposure bake time (min)

Post-exposure curing time (min)

Desired height SU8 photoresist RPM 65 °C

95 °C

65 °C

95 °C

15

3010

1600 1

10

1

3

25

3025

3000 1

10

1

3

50

3025

1600 1

15

1

5

50

3035

2000 1

15

1

5

Step 0. Set slope at 5 second (0 rpm to 500 rpm ramp). Step 1. Set RPM to 500 and time to 10 s (spin at 500 rpm for 10 seconds). This will allow the spread of SU8 on the substrate. Step 2. Compute ramp rate from 500 to 3000 rpm. 3000–500 rpm/300 rpm s-1 = 8.3 s. Set slope to 8.3 s. Step 3. Set RPM to 3000 and time to 30 s (maximum rotation speed for 30 s). This controls the layer thickness. Step 4. Compute the negative slope rate from 3000 to 0. 3000 rpm/ 100 rpm s-1 = 30 s. Step 5. End. The photoresist mold for the designs in this device needs a height of 15 and 25 μm for the flow control layer and 25 and 50 μm for the flow layer. Different spin rates (in RPM) and SU8 formulations are used for each target height as shown in Table 1 (see Note 10). 3. Subject the coated wafer to pre-exposure soft-bake by placing it on a hot plate at 65 °C for 1 minand then 95 °C for 10 mins (for a layer height of 15 and 25 μm) or 15 mins (for 50 μm height) (see Note 11). 4. Expose the coated wafers to UV light (20,000 mJ cm-2 at 405 nm). In this work, use a maskless exposure apparatus. This exposure follows the design pattern data with a 1 μm resolution in the design file (see Note 12). 5. After the multilayer exposure, subject the coated wafer to postexposure curing at a temperature of 65 °C (1 min) and 95 °C (3 mins for 15 and 25 μm and 5 mins for 50 μm height).

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6. Submerge the cured photoresists in SU8 developer, the time of which depends on the thickness of the resist on the wafer (20–50 μm for 5–8 min). Rinse with ethanol and dry using N2 air gun. 7. Glue metal posts (optional) to the inlet or outlet mold structure to serve as a scaffold during soft lithography. 8. Subject the wafer with the mold to silanization using 1H,1H,2H,2H-perfluorodecyltrichlorosilane to enable an easy release of PDMS during pattern transfer. The resulting molds before silanization are shown in Fig. 2. 3.2.3 PDMS Channel Pattern Transfer and Device Assembly

For a valve-based microfluidic device assembly, prepare different off-ratios of the mixture of PDMS oligomer and curing agent for each layer (see Note 13). In this work, the mixtures with the ratios of 20:1 and 5:1 are preferred for the flow and control layer, respectively. 1. Spin-coat the prepared mixtures of PDMS oligomer and curing agent with certain off-ratios on the SU8 molds. In this work, about 50 μm thickness is preferred for the flow layer mold and about 2 mm thickness for the control layer mold. The 20:1 mixture is coated at 2000 rpm for the flow layer while the 5:1 mixture is coated at 100 rpm for the control layer. 2. Place the PDMS coated wafers on an 80 °C hot plate and soft cure for about 8–10 mins for the flow layer and 20 mins for the control layer. 3. After curing, cut out the control layer PDMS to a shape enclosing the mold design. Remove from the mold, degrease by soaking in IPA for 3–5 mins, and air-dry using N2 gun. For IPA rinsing explanation, see Note 14. 4. Align the cutout control layer on top of the flow layer. This can be done manually under a compound microscope, ensuring that the hydrodynamic trap at the bottom aligns with the ring-shaped valve on top. 5. Pour the remaining 5:1 PDMS mixture around the aligned layers and cure for at least 2 h at 80 °C in an oven. Overnight curing is preferred for PDMS bonding (see Note 15). 6. Cut the bonded PDMS layers off the mold and rinse with IPA for 3–5 mins. Dry with an N2 gas gun afterward. 7. Subject the PDMS layers and clean micro cover glass to oxygen plasma exposure using a Plasma Dry Cleaner for 50 s at the maximum power of 75 W (see Note 16). 8. Bind the PDMS block and glass substrate through conformal contact using an N2 gas gun. An illustration of the layer composition and image of the microfluidic device is shown in Fig. 3.

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Fig. 3 The microfluidic chip is a multilayer device that is made up of a control layer, a flow layer, and a glass substrate: (a) illustration of the layer composition of the device and (b) image of the chip with the channels filled with colored liquid, red for the control channels and blue for the flow channels

9. Place the assembled microfluidic device on an 80 °C hot plate for 10 mins. Fill the channels with liquid (i.e., PBS, ultrapure water, or deionized water). This is to prevent hydrophobic recovery (hydrophilic loss) by keeping the flow channels wet during storage [11]. The plasma treatment done to the PDMS increased its surface energy, making it hydrophilic [12]. 10. Store the microfluidic device at 4 °C (see Note 17). 11. In preparing the chip for use in experiments, apply the BSA coating to the PDMS surface to reduce adhesion of cells on it. Fill the flow channels with 1% BSA in PBS solution and incubate at 4 °C for 1 h or at room temperature overnight. 3.3 Cell Preparation, Culture, and Sorting

The model cells used to test the capability of the device in detecting and quantifying GzmB in the single-cell assay system include granzyme B (GmzB)-transduced Jurkat T-lymphocyte, NK92 natural killer, and THP1 monocyte cells. Human peripheral blood mononuclear cells (PBMCs) derived from healthy individuals and lung cancer patients were also used in the platform. For information about the cell sample source, see Note 18.

3.3.1 Culture Media and Cell Culture

1. To create the culture media for the Jurkat, THP1, and PBMC cells, put RPMI 1640 medium, fetal bovine serum (FBS), and penicillin-streptomycin in a water bath at 37 °C. 2. Mix them in the composition of 10% FBS and 1% penicillinstreptomycin using a vortex mixer or manually by flipping the sealed container up and down, repeatedly. 3. To create the media for NK92 cells, mix alpha medium with 2 mM L-glutamine, 0.2 mM i-inositol, 20 mM folic acid, 0.1 mM 2-mercaptoethanol, 12.5% FBS, and 12.5% horse serum in the presence of 500 IU/ml human interleukin2 (IL-2).

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4. The media can be stored in a 4 °C refrigerator. 5. Fast thaw a vial of cryopreserved cells by quickly dipping it in and out of a 37 °C water bath. 6. Once the cell suspension is thawed, transfer them dropwise to a 10 mL centrifuge tube containing 4 mL RPMI medium at 37 °C. 7. Resuspend the cells by gently pipetting up and down (see Note 19). 8. Put the tube with the cell suspension in a centrifuge and spin at 900 rpm for 5 mins at 4 °C. Afterward, aspirate the supernatant and dead cells. 9. Add 4 mL RPMI at 37 °C dropwise to the cell pellet; in order to avoid thermal shock on cells, gently pipette up and down to resuspend the cells again. 10. Determine the cell concentration using a disposable hemocytometer. Afterward, transfer the volume of the cell suspension according to the desired number of cells to a culture flask containing the prepared media. 11. Stored the cell culture in an incubator at 37 °C and 5% CO2. 3.4 Sample Preparation Prior to Single-Cell Assays 3.4.1

Cell Harvesting

In preparation for the single-cell assay experiment, the model cells are stimulated through incubation in a media containing IL-2 (500 IU/ml) at 37 °C and 5% CO2 for 12 h. 1. From the prepared culture, harvest and rinse the cells. 2. Adjust the number of cells in the suspension to a concentration of 1–5 × 106 cells/ml in an ice-cold running buffer (see Subheading 2.4, step 1) (see Note 20).

3.4.2

For PBMCs

For human PBMCs, conduct DNase and/or collagenase treatment to reduce the clumping and sticking of cells to the PDMS channel: 1. Centrifuge the cells at 300 g for 5 mins. Aspirate and discard the supernatant and resuspend the cells with 1 mL of DNase/ collagenase solution (see Subheading 2.2.4). Treat the cell samples for 30 min at 37 °C. 2. Centrifuge the cells again at 300 g for 5 mins. Discard the supernatant and resuspend the cells in 3 mL running buffer (see Subheading 2.4, step 1). 3. Make the cells pass through a 4 μm cell strainer that is placed on a 5 mL round bottom tube. Then, wash the strainer with 2 mL running buffer. 4. Centrifuge the cells at 300 g for 5 mins. Discard the supernatant and resuspend the cells in the prepared RPMI cell medium.

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3.5 GzmB Enzymatic Assays 3.5.1 Single-Cell GzmB Assay

Use the high-throughput valve-based device here to conduct the GzmB enzymatic activity assay, to quantify the protease expression of each compartmentalized cell, and to conduct surface marker staining for cell classification. Under the same concept, other extracellular protein secretions can be quantified using the same platform with corresponding peptide substrates, antibodies, and probes. An image of the microfluidic platform used for single-cell assay is shown in Fig. 4. The following are the steps in conducting the single-cell GzmB enzymatic activity assay: 1. Prewarm the assay buffer (see Subheading 2.3) in a water bath at 37 °C. Thaw the peptide substrate (Ac-IEPD-AFC) to room temperature. Connect external equipment like syringe pump and air pump to the microfluidic device’s inlets/outlet (Fig. 4). Turn on the stage heater and set at 37 °C. 2. Wash the microfluidic channel (with BSA-treated surface for protein blocking) by rinsing it with PBS or assay buffer through the action of a syringe pump. 3. Pipette 50 μL of the prepared cell suspension into the device inlet. Flush the cells into the flow channel at a flow rate of 30 μL/min with the aid of the syringe pump. Cells that flow in the channel pass through the hydrodynamic traps, allowing the cell capture (see Note 21).

Fig. 4 Microfluidic platform with external equipments attached to the system

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4. Flush out excess or untrapped cells by rinsing the flow channel with PBS buffer medium at 50 μL/min. 5. Actuate the ring-shaped pneumatic microvalves by applying air pressure (0.04–0.05 MPa) to deflect the thin PDMS membrane around each trap. This membrane deflection creates sealed microchambers that compartmentalize the cells. Both the 1000- and 5000-chamber chip can be sealed using the abovementioned pressure. 6. Scan and capture images of the microchamber field (preferably at 10x magnification) using a fluorescence microscope both in bright-field and in fluorescence modes. The fluorescence intensity under the 360 nm/460 nm filter cube, which is also called DAPI, serves as background measurement. 7. While the microchambers are sealed, flush in the flow channel the antibody/antibodies (see Note 23) for staining desired cell subset (i.e., anti-CD14, anti-CD56, anti-TCRαβ), see Table 2, and for dead cell staining (using PI). Briefly, open and close the microchambers by switching the valve actuation off and on, respectively. Incubate for 30 min. Open the valves and rinse the microfluidic channels with the assay buffer. For the model cells, the antibody stains are individually perfused in the microchambers according to the cell being evaluated. 8. Flush in the peptide substrate (Ac-IEPD-AFC) (see Note 22) in assay buffer in the same manner that was done to the antibody stains. Observe a 30-min incubation.

Table 2 The different antibodies, probes (PI), and GzmB peptide substrate introduced into the microchambers for the enzymatic activity assay and surface marker staining

Excitation/emission (nm) 360/460

Antibodies (ab), probes, and substrates Cocktail A

Cocktail B

Cocktail B′

Others

GzmB substrate

GzmB substrate

GzmB substrate

GzmB substrate

405/421

CD56 and CD14 (human)

470/525

CD56 (human)

CD8 (human)

535/617 545/605

PI (Texas red) PD1 (human) IgG4 (human)

IgG4 (human)

635/688 620/700

CD3 (human)

TCR α/TCR β and CD56 (human)

CD3 (human)

CD8 (human)

CD3 (human)

Each cocktail contains GzmB substrate, assay buffer, Fc blocker/anti-Fc receptor antibody, and the surface marker antibody. Antibodies and probes were combined as a cocktail (A, B, and B′) in PBMC experiments and individually introduced in model cell experiments

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Fig. 5 Sample images of the microchambers showing (left) gray-scale fluorescence from the cleaving of the peptide substrate and (right) surface marker stains of different cell subsets

For PBMC cells, perfuse a mixture or cocktail of antibodies, probe, and substrate; see Table 2. 9. After incubation, scan and take the image of the entire microchamber field under different optical filters. 10. Measure the fluorescence intensity from the cleaved AFC molecule, imaged through the DAPI (360/460 nm) filter, in each microchamber containing single cell using an image analyzer software. In this work, use the software to create the template file (see Subheading 3.5.2) for auto-identification of the region of interest or ROI (i.e., the area inside the microchamber), and measure the pixel intensity of the acquired DAPI images. 11. Sample images of the assay fluorescence from AFC molecule release and surface marker staining are shown in Fig. 5, and a sample GzmB activity profile is shown in Fig. 6. 3.5.2 Image Analysis Template File Creation

1. Open a sample “.tif” file from the bright-field image folder. 2. Select “Hybrid Cell Count” from the menu panel, and in the Hybrid Cell Count window, select Fluorescence in the “Select image type” and Single extraction in the “Specify target area.” Click start. 3. In the Create Target Area stage, a dialog box asking for color display will appear. Select OFF and click OK. 4. An image of the bright-field file with a color overlay is displayed in the window. Create the target area by adjusting the threshold value to set the extraction area. The mask’s display result can be adjusted according to your preferred transparency value and extract and target color. Click next.

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Fig. 6 Sample GzmB activity profile of a patient PBMC according to identified cell subsets (left) and its percentage distribution (right)

5. Adjust the target area to ensure the ROI results to a circle covering the inside of the microchamber. Remove unwanted areas and correct the target area by using the available editing options. Click next to proceed to the extraction stage. 6. In the extraction stage, select OFF in the Separated Color Display dialog box. Click the open Icon next to “Specify Ext. Area,” and open the DAPI image file that matches the brightfield image. Select fluorescence in the “Select image type” and click Apply. 7. An image of the DAPI image file with an overlay of the target area mask created previously is displayed. Adjust the threshold value to set the extraction area for the fluorescence. If it is the same as the target area created for bright field, set it to zero. Display details for the fluorescence extract area can be modified. Click Next. 8. The resulting view shows the bright-field image with an overlay of the mask created for the target/extraction area. These ROIs are numbered for each identified microchamber. The measurement results can be edited to include measurement of the ROI area and mean pixel intensity in the red, blue, green, etc. Save the template file (*.mcd) by clicking “Save conditions.” Click exit. The following are the steps taken to perform the batch measurements or measure multiple files in a single operation according to the created template.

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9. In the image analyzer software menu panel, click “Hybrid Cell Count.” In the pop-up window, select start under the “Macro Cell Count.” 10. Click “Open” under the Read conditions file and select the created template (*.mcd) file. 11. In the “Target files,” add all the bright-field image files that will be analyzed by either dragging and dropping or selecting the “Add” and “Batch load” options. Next, add all the matching DAPI files in the same manner. 12. Select “Run” to execute the operation in all the files. 13. Click “Save” to save the results in *.csv file, containing the measurement values; *.jpeg file, showing the overlay of the bright-field and DAPI image files; and the ROI mask. 14. Open the *.csv files in MS Excel to create a table. In this work, the mean pixel intensity measured by the software was referred to as relative fluorescence unit (RFU). Subtract the RFU from ROI considered as background (R1) from the measurements of the target ROI (R2) to get ΔR: ΔR = R2 - R1 15. Classify the cell state (viability) through the PI stain and cell subset through the antibody stain with the fluorescence imaging data. Match the stain information with each cell’s corresponding GzmB (ΔR) activity to create the sample’s multi-single-cell profile. 16. Apply statistical treatment and summarize the results in tables and graphs. 3.5.3

Bulk GzmB Assay

Measure the GzmB activity of bulk samples using a 96-well plate as reference for comparison. Use a commercial fluorometric GzmB activity assay kit for this work (see Subheading 2.3): 1. Warm the assay buffer to room temperature. Thaw frozen vials of positive control (granzyme B enzyme, human recombinant), GzmB substrate, and AFC standard. 2. Using a 96-well plate, pipette 1, 5, 10, 20, and 50 μL (or 1–50 μL) of the cell sample per well. Add 2 μL of positive control in selected wells. 3. Add the GzmB assay buffer in each well to a final volume of 50 μL. Add the 50 μL reaction mix to each well with the cell sample and positive control. The reaction mix contains 5 μL of the GzmB substrate and 45 μL of assay buffer. Mix the contents in the wells by pipetting up and down.

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4. Incubate for 30–60 mins at 37 °C, protected from light. Afterward, measure the fluorescence at 380/500 nm (ex/em). 5. Using a fluorescence microscope, take the images of each well at different points. Similar to the discussion in the previous section, use the bright-field and the DAPI (360/460 nm) filter during image acquisition. 6. Measure the pixel intensity in the region of interest (ROI) using an image analysis software. A mask can be made to assign the ROI (see Subheading 3.5.1). The features of the hybrid cell count can be used to generate a measurement result containing the pixel intensity. Open the saved output file (*csv) in MS excel and locate the RFU of the numbered ROI. Determine the RFU for the background (R1) and the target (R2). Subtract R1 from R2 to get ΔR. Interpolate these values from the standard curve to quantify the GzmB expression. 3.5.4

AFC Standard

Obtain the AFC standard curves using the microfluidic device for single-cell measurement and 96-well plate bulk measurement by following the assay kit protocol. Here, prepare different concentrations of the AFC standard in granzyme B assay buffer: 1. Prepare 0, 50, 100, 150, 200, and 250 pmol/μL of AFC standard. This can be done by adding 0, 5, 10, 15, 20, and 25 μl of 10 μM (10 pmol/μL) AFC standard in selected wells of a 96-well plate, adjusting the volume to 100 μL with assay buffer. Mix well through pipetting. 2. Capture bright-field and fluorescence (DAPI) images of each ROI per chamber or well. Measure the pixel intensity (RFU) using an image analyzer software (see Subheading 3.5.2). Subtract the RFU measurement of 0 pmol AFC standard from all readings (see Note 24). Plot the standard curve. Sample standard curves are shown in Fig. 7. 3. Calculate the GzmB activity by interpolating the RFU values from the standard curve. With a high-throughput single-cell assay, better representation of the sample population can be obtained, and at the same time, variations in each cell activity can be investigated [13]. Profiling the cell subsets can show the level of GzmB activity whether it is overexpressed, weakly expressed, or not expressed. This can reveal certain conditions like T-cell exhaustion, having low GzmB activity, present in the patient’s blood sample. Response to treatments or drug screening tests, for example, GzmB inhibitors, can also be reflected in the increase or decrease in the GzmB level.

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Fig. 7 AFC standard curves for bulk sample using (left) 96-well plate and for single-cell analysis using a (right) microfluidic device. The dotted lines in each graph represent the 95% confidence bands of the best fit line 3.6 Cell Trap Rate and Valve Actuation Test 3.6.1 Rate

Single-Cell Trap

Test the fabricated microfluidic device to determine its single-cell trap rate: 1. Using the microfluidic device, pipette a 100 μL cell suspension of Jurkat cells (concentration of 1–5 × 106 /ml) into the inlet of the device. Fluorescent microbeads or other cell types can also be used. 2. Connect the syringe pump. Use varying flow rates (i.e., 1, 2, 5, 10, 20, 30, 40, and 50 μL/min) for each trial. 3. Count the number of cell(s) (or bead) in each trap. Compare the single-cell trap occupancy for each flow rate, and identify the optimized value with the highest occupancy (see Note 25).

3.6.2 Valve Actuation and Microchamber Sealing

To determine the lowest sealing pressure required for the microfluidic device, test the actuation of the valves by applying varying degrees of air pressure at the control inlet. Fill the flow channel with a fluorescent solution (e.g., FITC dextran, 1 mg/ml). Actuation of the valves will cause the defection of the PDMS membrane separating the two layers (control and flow layers) and will displace the fluorescent solution. Confirm that the microchamber is sealed by the absence of the fluorescence or fluorescent solution in the area of valve actuation. Measure the fluorescence intensity in this area under varying degrees applied, and compare it to the intensities of the areas without valve actuation and background. Investigate the effect of the membrane thickness separating the two layers (10 μm–30 μm), as a result of varying the RPM during the PDMS coating on the flow layer mold, to the sealing pressure

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(see Note 26). Similarly, examine the effect of stochastic off-ratio of the PDMS for the flow layer (10:1 and 20:1) and control layer (5:1 and 10:1) to the sealing and rupture pressure (see Note 26). The results of this actuation tests at different parameters have been reported (see Note 27) in our previous work [13].

4

Notes 1. Rectangular-shaped valve can create gaps that will allow fluid to pass or escape the microchamber during actuation resulting in a compartmentalization failure. 2. Keep the maximum value of R at 140 μm to ensure a distance between the mechanical traps not greater than 300 μm. Set this limit in order to avoid further decrease in the single-cell trap occupancy with the widening of the trap distances. 3. Use solid mechanic modules in the simulation of the microchamber block’s membrane deflection. For the elastomer membrane deflection, use the solid mechanics for structural mechanics physics module to understand the relation of the valve diameter and height with the sealing pressure. 4. Since PDMS is a hyperelastic and near-incompressible material, choose the neo-Hookean model with a Lame parameter of 6.67 × 105 N/m2 to describe the behavior of a rubber-like material experiencing a large deformation that is obtained by a statistical-mechanical deformation treatment of freely joined molecular chains. 5. Selecting the boundaries as fixed constraints is to set the identified surfaces as non-movable. 6. Choose the laminar flow for the fluid flow physics for the valve channel using weakly compressible flow to determine the fluid velocity and pressure profile. Use air as well as oil materials in the fluid flow simulation. 7. The values entered in the “range” under parameter value in the Study Extensions set the minimum, step, and maximum pressure to be computed. This range is arbitrary and, thus, depends on the parameters being investigated. 8. Details of this simulation have been reported elsewhere [13]. In summary, the step-like design for the ring valve has been found to achieve a smaller sealing pressure and bigger limit for rupture pressure. In this design, it is found that the valve’s outer radius (R), more specifically the bottom radius Rbottom, is the primary parameter contributing to lowering sealing pressure. Bigger R or Rbottom has resulted to lower sealing pressure. Secondary to R is the height of the cylindrical valve. A higher h contributes to a lower sealing pressure.

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This design is found to reach a higher point of failure or rupture pressure, which means the valve can hold greater pressure before failure. The analysis also shows that this type of design has relatively lower von Mises stress and strain energy density which means a lower risk of material failure. This geometry with an R of 140 μm and height of 50 μm is identified as the optimized design for the fabrication of the photoresist mold and the assembly of the PDMS microfluidic device. 9. Photoresists are light-sensitive materials that consist of a polymer, sensitizer, and solvent. There are two types of photoresists, negative and positive. For a negative photoresist like SU8, areas exposed to light result in polymerization, and they remain on the substrate after wet chemical development [14]. The opposite happens to positive photoresist. Both types can be used in mold fabrication and it depends on the user’s preference. 10. For the thickness of 15 μm, SU8 3010 formulation can be used and spin-coated at about 1500 rpm. For 25 μm thickness, SU8 3025 (3100 rpm) or SU8 3035 (4000 rpm) can be used. A 50 μm thickness can either be achieved by SU8 3035 at 2000 rpm or SU8 3025 at 1600 rpm. 11. The soft bake stage allows the evaporation of the solvent, making the photoresist more solid. The heat pattern (from 65 °C, then to 95 °C plateau, and cooldown) decreases the mechanical stress inside the SU8 photoresist [15]. 12. Save the CAD drawing file (.dwg) to .dxf file. Using a converter software (dfx-to-gds), convert the dxf file to a gds file. Then, convert the gds file to png or bmp image files using another converter software (gds-to-png). Use the png files to create a recipe file (.mer) using a recipe editor software. This type of file is read by the maskless exposure machine. 13. The flow layer needs a softer PDMS membrane that can be easily deflected during pneumatic actuation of the valves. This can be achieved by higher PDMS off-ratios like 10:1 and 20:1. Meanwhile, the control layer needs a stiffer PDMS to hold the air pressure down. In this regard, a 5:1 ratio is preferred. 14. Use IPA to degrease PDMS and leach out additives. This removes un-crosslinked material and strengthens the bond of the two layers during device fabrication. This significantly improves the amount of pressure that the control layer can hold before rupture. 15. The interaction of the excess functional groups in the two layers allows a covalent bond to be formed across the interface resulting in an irreversible bonding [16].

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16. The expose of the bonding surfaces of the PDMS layer and glass substrate to oxygen plasma makes these surfaces extremely clean, which facilitates the bonding process. The use of O2 plasma removes organic, hydrocarbon materials through chemical reaction with highly reactive oxygen radicals and ablation by energetic oxygen ions. This leaves silanol (SiOH) groups on the surface. Conformal contact of the surfaces forms the bridging Si-O-Si bond at the interface, creating an irreversible seal [17, 18]. 17. Storage at a colder temperature such as 4 °C or lower is preferred as colder temperature results to reduced diffusion of low molecular weight species (LMW) in the polymer matrix, thus reducing the hydrophobicity recovery rate [11]. Storage at this temperature also helps remove small air bubbles trapped in the channel. 18. All cells have been provided by collaborators from Osaka University Hospital, and they are expanded from a cryopreserved master cell bank. 19. It should be noted that the size of the pipette tip bore and the pipetting rate may affect cell viability, spreading, and proliferation [19, 20]. A bigger bore size is preferred (i.e., 1000 μL tip). For smaller bore size, a slow mixing action is recommended. 20. All reagents/solutions to be used, except for the running buffer, DNase, and collagenase solutions, need to be prewarmed in a water bath at 37 °C. 21. The cells follow a streamline as they move inside the flow channel. With pillar structures serving as flow guides, some of these streamlines will pass through the hydrodynamic trap gap (hourglass-shaped) that mechanically captures a cell. The width of the gap (maximum width, 15 μm, minimum width, 5 μm) in this device was intended for trapping immune cells. This width can be adjusted during the mold fabrication according to the target cell size. The optimized flow rate is dependent on the channel dimensions. For the 5000-chamber device, 30 μL/ min is the optimized rate, as obtained from the results of trapping test. 22. To measure the GzmB expression from each cell, a tetrapeptide substrate (Ac-IEPD-AFC) containing the protease recognition sequence IEPD or Ile-Glu-Pro-Asp and fluorescent label (AFC) is introduced in each chamber by briefly opening and closing it through the actuation of the valves. GzmB prefers cleaving after the aspartic acid at position P1 of the nonfluorescent substrate [21]. This enzyme catalyzed reaction results in the cleaving of the substrate and release of the fluorescent probe (AFC), whose intensity is proportional to the enzyme activity [22].

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23. The antibodies for immunostaining can be individually introduced or combined as a cocktail. Fc blocker and other probes (i.e., propidium iodide or PI for cell viability) can also be combined together with the antibody in the microchamber. In this work, use anti-CD3, CD8, CD14, CD56, anti-IgG4, anti-PD1, and TCR α/β antibody with the excitation and emission wavelengths shown in Table 2. Use anti-CD3 and anti-CD8 to identify T-cells, anti-CD14 to identify monocyte cells, and anti-CD56 to identify NK cells. Use anti-PD1 to determine the expression of inhibitory receptor programmed death 1 (PD-1) by activated T-cells. Meanwhile, use anti-IgG4 to detect immune checkpoint inhibitors targeting PD-1 on T-cells. Measure the GzmB activities of these cells, particularly the PD-1- and IgG4-positive PBMC cells, and evaluate the activities to understand the possible response of patients to immune checkpoint blockade therapy. All of the substrate-antibody cocktails presented in Table 2 contain 5 μL of GzmB substrate, 5 μL assay buffer, 10 μL of Fc blocker/anti-Fc receptor antibody (see Subheading 2.3, step 4), 15 μL of the surface marker antibody specified in Table 2 (see Subheading 2.3, step 5) at concentrations of IgG4 (0.2 μg/mL), CD3 (100 μg/mL), CD14 (100 μg/mL), TCR α/β (100 μg/mL), and CD56 (200 μg/mL)). In the case of CD56, different fluorescent labels with different excitation/emission wavelengths were assigned to respective antibody cocktails to prevent overlap from other fluorescent labels as the number of available microscope filters is limited; see Table 2. For PBMCs, under the pre-immune checkpoint inhibitor treatment group, use cocktail A containing GzmB substrate, assay buffer, Fc blocker, anti-CD3, anti-CD56, and PI. Moreover, use cocktail B containing GzmB substrate, buffer, Fc blocker, anti-PD1, anti-CD3, and anti-CD8 in the assay experiment for the same sample. On the other hand, under the post-treatment group for the PBMCs, use cocktail A that is previously described and cocktail B which contains GzmB substrate, buffer, Fc blocker, anti-IgG4, anti-CD3, and anti-CD8. Meanwhile for model cells, individually introduce the antibody stains according to the type of cell used in the microfluidic device. For instance, use anti-CD56 for NK-92 cells, antiTCR α/β for Jurkat cells, and anti-CD14 for THP1 cells. 24. The fluorescence measurement is based on the maximum observable intensity after the incubation time. One unit of the granzyme B (GzmB) refers to the amount of the enzyme that hydrolyzes 1 pmol of the substrate per minute. Then, interpolate the fluorescence intensity measurement in each chamber from the standard curve to acquire information on the GzmB expression.

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25. For the cell concentration mentioned and using a 5000chamber chip, the optimized flow rate is found to be at 30 μL/min, achieving about 61% single-cell trap occupancy. The detail of this test result was reported elsewhere [13]. The size of the flow channel affects the flux of cells as well as their speeds. For example, a narrower channel from a 1000-chamber device leads to a 10 μL/min flow rate [22]. It is worth noting that a greater number of chambers require bigger device size as well as the possibility of higher gas release due to gas absorption, tubing connection gas leakage, and PDMS permeability from increased wall surface area [13]. Optimization conditions according to the throughput must be identified to get the highest single-cell trapping rate. Increasing the concentration and volume of the cell suspension that is made to flow in the device can increase the single-cell occupancy and reach about 80% trap occupancy for this device. For assay experiment, only the traps with one cell were included in the data evaluation process. 26. The sealing pressure refers to the pressure applied to the valve with an absence of fluorescence found in the region around the valve or at the same intensity as the background without FITC. On the other hand, the highest amount of the applied pressure that the microchamber unit can hold before compartmentalization fails is referred to as the rupture pressure. This failure can be due to membrane rupture, layer-layer bonding failure, or both. 27. In summary, a sealing pressure of 0.04 MPa is achieved for sealing an array of 5000 chambers. A 20:1 ratio of the PDMS mix for the flow layer and 5:1 ratio for the control layer are preferred as this combination gives the bonded device a greater amount of pressure that can be held before failure. Meanwhile, the membrane thickness is investigated to give a significant impact on the sealing pressure. References 1. Sinha N, Subedi N, Tel J (2018) Integrating immunology and microfluidics for single immune cell analysis. Front Immunol 9:1–16 2. Kulasinghe A, Wu H, Punyadeera C et al (2018) The use of microfluidic technology for cancer applications and liquid biopsy. Micromachines 9(8):397 3. Au AK, Lai H, Utela BR et al (2011) Microvalves and micropumps for BioMEMS. Micromachines 2(4):179 4. Blazek M, Betz C, Hall MN et al (2013) Proximity ligation assay for high-content profiling of cell signaling pathways on a microfluidic chip. Mol Cell Proteomics 12(12):3898–3907

5. Huang Q, Mao S, Khan M et al (2013) Singlecell assay on microfluidic devices. Analyst 144: 808–823 6. Eyer K, Stratz S, Kuhn P et al (2013) Implementing enzyme-linked immunosorbent assays on a microfluidic chip to quantify intracellular molecules in single cells. Anal Chem 85(6): 3280–3287 7. Stratz S, Verboket PE, Hasler K et al (2018) Cultivation and quantitative single-cell analysis of Saccharomyces cerevisiae on a multifunctional microfluidic device. Electrophoresis 39: 540–547

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8. Fan H, Wang J, Potanina A et al (2011) Wholegenome molecular haplotyping of single cells. Nat Biotechnol 29:51–57 9. Streets AM, Zhang X, Cao C et al (2014) Microfluidic single-cell whole-transcriptome sequencing. Biol Sci 111(19):7048–7053 10. Sun Y, Cai B, Wei X et al (2019) A valve-based microfluidic device for on-chip single cell treatments. Electrophoresis 40(6):961–968 11. Jahangiri F, Hakala T, Jokinen V (2020) Longterm hydrophilization of polydimethylsiloxane (PDMS) for capillary filling microfluidic chips. Microfluid Nanofluid 24(2) 12. Lin WC, Mohd Razali NA (2019) Temporary wettability tuning of PCL/PDMS micro pattern using the plasma treatments. Materials 12(4):644 13. Briones J, Espulgar W, Koyama S et al (2021) A design and optimization of a high throughput valve based microfluidic device for single cell compartmentalization and analysis. Sci Rep 11: 12995 14. Quero AJM, Perdigones F, Aracil C (2021) Microfabrication technologies used for creating smart devices for industrial applications. In: Nihtianov S, Luque A (eds) Woodhead publishing series in electronic and optical materials, smart sensors and MEMs, 2nd edn. Woodhead Publishing, Cambridge, pp 291–311 15. How to make an epoxy SU-8 mold? The SU-8 mold fabrication process: tips and tricks (2021)

ElveFlow. http://www.elveflow.com/micro fl u i d i c - r e v i e w s / s o f t - l i t h o g r a p h y microfabrication/su-8-mold-lithography. Accessed 25 Nov 2021 16. Lai A, Altemose N, White JA et al (2019) On-ratio PDMS bonding for multilayer microfluidic device fabrication. J Micromech Microeng 29(10):107001 17. PDMS BONDING (2021) Harrick Plasma. https://harrickplasma.com/pdms-bonding. Accessed 25 Nov 2021 18. Tan SH, Nguyen NT, Chua YC et al (2010) Oxygen plasma treatment for reducing hydrophobicity of a sealed polydimethylsiloxane microchannel. Biomicrofluidics 4(3):32204 19. Hanamsagar R, Reizis T, Chamberlain M et al (2020) An optimized workflow for single-cell transcriptomics and repertoire profiling of purified lymphocytes from clinical samples. Sci Rep 10:2219 20. Agashi K, Chau DY, Shakesheff KM (2009) The effect of delivery via narrow-bore needles on mesenchymal cells. Regen Med 4(1):49–64 21. Martins CDF, Raposo MMM, Costa SPG (2021) A new fluorogenic substrate for granzyme B based on fluorescence resonance energy transfer. Chem Proc 3(1):6 22. Briones JC, Espulgar WV, Koyama S et al (2020) A microfluidic platform for single cell fluorometric granzyme B profiling. Theranostics 10(1):123–132

Chapter 11 Single-Cell Exogenous Gene Transfection Analysis Chip Haiyang Xie and Xianting Ding Abstract Fast and accurate profiling of exogenous gene expression in host cells is crucial for studying gene function in cellular and molecular biology. This is achieved by co-expression of target genes and reporter genes, but we still have to face the challenge of incomplete co-expression of the reporter and target genes. Here, we present a single-cell transfection analysis chip (scTAC), which is based on the in situ microchip immunoblotting method, for rapid and accurate analysis of exogenous gene expression in thousands of individual host cells. scTAC not only can assign information of exogenous gene activity to specific transfected cells but can also enable the acquisition of continuous protein expression even in incomplete and low co-expression scenarios. Key words In situ microchip, Immunoblotting, Incomplete co-expression, Transfection, Single cell

1

Introduction Transient transfection of fusion plasmids is a commonly utilized method for studying specific gene functions in molecular biology [1]. With a promoter and a terminator, the reporter gene and target gene form a fusion plasmid, and it co-expresses a fusion protein [2]. Fusion proteins with fluorescent reporter protein tags are important for evaluating the efficiency of editing genes [3], tracing protein signals [4], and nondestructively studying cell biological functions such as the cell cycle [5]. However, even in successfully transfected cells, incomplete co-expression of the fusion gene could occur and generate false-positive signals. This may probably be due to the complex enzymatic environment and translation modification mechanism [6]. In the existing methods, quantitative RT-PCR (qRT-PCR) and fluorescence-activated cell sorting (FACS) can reveal the expression of fusion genes in transfected cells [7]. However, qRT-PCR, which evaluates the mRNA level rather than the protein expression level, does not reflect real transfection efficiency due to inconsistencies

Paul C.H. Li and Angela Ruohao Wu (eds.), Single-Cell Assays: Microfluidics, Genomics, and Drug Discovery, Methods in Molecular Biology, vol. 2689, https://doi.org/10.1007/978-1-0716-3323-6_11, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Hydrophobic treatment is performed on the isolation area to limit sample cross-contamination. The cell samples are loaded into the sample loading area. Cells expressing the reporter gene are located with confocal fluorescence microscope and assigned with corresponding coordinates. Transfection quality is examined with single-cell immunoblotting analysis showing the correctly expressed proteins. After the target protein is successfully verified, the coordinates are recorded. Different fluorescent antibodies are used to detect protein expression, in which distinct colors represent different protein signals. Subsequent proteomic analyses are performed only for successfully transfected cells

between transcriptional and posttranslational levels [8]. Furthermore, low co-expression of fusion proteins in cells could cause false positives in FACS analysis, interfering with downstream protein expression analysis of the introduced gene in the signaling pathway [9]. Particularly, for some cells that are difficult to transfect, the efficiency of transient transfection is considerably low, resulting in a sparse number of cells inadequate for analysis in each process batch [10]. These limitations not only hinder the functional study of proteins but may also result in incorrect conclusions [11]. To enable simultaneous screening of exogenous gene expression and subsequent protein expression analysis in individual host cells, we herein develop a single-cell transfection analysis chip (scTAC) (Fig. 1), a variation of scWB (single-cell Western blotting) method that combines in situ immunoblotting and photoclick bioorthogonal chemistry [12].

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Materials Prepare all solutions using ultrapure water (prepared by purifying deionized water, to attain a resistivity of 18 MΩ–cm at 25 °C) and analytical grade reagents. Prepare and store all reagents at room temperature (unless indicated otherwise). Diligently follow all waste disposal regulations when disposing waste materials. 2.1. 10% (w/v) Ammonium Persulfate (APS) Dissolve 10 mg of APS in 100 μL of purifying deionized water; store this solution at 4 °C for a short-term period (